Drosophila blood cells, or hemocytes, cycle between resident sites and circulation. In the larva, resident (sessile) hemocytes localize to inductive microenvironments, the Hematopoietic Pockets, while circulating hemocytes move freely in the hemolymph. The goal of this protocol is the standardized isolation and quantification of these two, behaviorally distinct but interchanging, hemocyte populations.
In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism Drosophila melanogaster, inductive microenvironments known as larval Hematopoietic Pockets (HPs) have been identified as anatomical sites for the development and regulation of blood cells (hemocytes), in particular of the self-renewing macrophage lineage. HPs are segmentally repeated pockets between the epidermis and muscle layers of the larva, which also comprise sensory neurons of the peripheral nervous system. In the larva, resident (sessile) hemocytes are exposed to anti-apoptotic, adhesive and proliferative cues from these sensory neurons and potentially other components of the HPs, such as the lining muscle and epithelial layers. During normal development, gradual release of resident hemocytes from the HPs fuels the population of circulating hemocytes, which culminates in the release of most of the resident hemocytes at the beginning of metamorphosis. Immune assaults, physical injury or mechanical disturbance trigger the premature release of resident hemocytes into circulation. The switch of larval hemocytes between resident locations and circulation raises the need for a common standard/procedure to selectively isolate and quantify these two populations of blood cells from single Drosophila larvae. Accordingly, this protocol describes an automated method to release and quantify the resident and circulating hemocytes from single larvae. The method facilitates ex vivo approaches, and may be adapted to serve a variety of developmental stages of Drosophila and other invertebrate organisms.
Research in the invertebrate model Drosophila melanogaster has driven the discovery of innate immunity1, and has facilitated the understanding of various aspects of blood cell development2-4. Drosophila hematopoiesis can be divided into the lineage of embryonic/larval hemocytes, which originate in the embryo and expand in the larva, and the lineage of lymph gland hemocytes4,5. Here, we present a protocol that focuses on the lineage of embryonic/larval hemocytes, which in the Drosophila larva mainly comprises plasmatocytes (macrophages) and few crystal cells4. In the larva, hemocytes of the embryo persist and colonize segmentally repeated and terminal Hematopoietic Pockets (HPs) located between the epidermis and muscle layers of the larval body wall5,6. Based on their nature as self-renewing macrophages6, their predominant residence in local tissue microenvironments6,7, and their lineage from the earliest blood cells emerging during development6,8, this blood cell population is considered similar to vertebrate self-renewing tissue macrophages, an independent myeloid lineage recently identified in a variety of species4,9,10. However, in Drosophila, some or all of these resident cells also show plasticity to give rise to other blood cell types such as crystal cells11,12.
Larval hemocytes are predominantly resident (sessile), but are in a dynamic steady-state between various HPs. They are progressively released into circulation, in particular as the 3rd instar larva approaches pupariation5-7. Immune challenges, injury or mechanical disturbance lead to a premature, in the latter case reversible, mobilization of resident hemocytes into the hemolymph4,6,13.
Previous studies have suggested that resident and circulating larval hemocytes are of the same lineage, but differ in their adhesive or homing properties6,7,13,14. Selective isolation of circulating versus resident hemocytes revealed elevated levels of proliferation in the resident hemocyte population, suggesting their exposure to inductive cues from the HPs6. Drosophila larval HPs are lined by epidermis and muscle layers and further harbor sensory neuron clusters of the Peripheral Nervous System (PNS) and liver function resembling oenocytes6. Functionally, mutant and genetic cell ablation experiments have demonstrated that sensory neurons present in the HPs support the trophic survival and localization of larval hemocytes6.
Here we describe a method for the specific isolation and quantification of resident and circulating hemocytes from single Drosophila larvae, and a protocol for mechanical hemocyte mobilization. The methods can be used for the ex vivo study of hemocytes and can further be adapted to other Drosophila developmental stages such as the pupa and adult, and other invertebrate systems. Since previous studies did not distinguish between resident and circulating hemocytes, this protocol provides a common standard for the study of resident blood cells and will help to increase the consistency of invertebrate blood cell research.
First, the Hemocyte Bleed/Scrape Assay describes the differential isolation and automated quantification of fluorescent protein-marked resident and circulating hemocyte populations from single Drosophila larvae; the protocol provides two options for regular and tile scan-equipped microscopes (Figure 1). As a result, the percentage of circulating hemocytes and the total number of hemocytes per larva are obtained. The method relies on transgenic Drosophila larvae that express fluorescent protein among their blood cell population. The choice of hemocyte driver or reporter determines the outcome, i.e., which population of blood cells is visualized and quantified. To label mainly macrophages (plasmatocytes), which comprise the vast majority of the resident and circulating hemocyte population of the Drosophila larva6, suitable transgenes include HmlΔ-DsRed 6, HmlΔ-GAL4 15, Pxn-GAL4 16, Crq-GAL4 (by H. Agaisse16), or eater-GAL4 17; for labeling the relatively small population of crystal cells, suitable lines are BcF6-CFP and -GFP 18, or lz-GAL4 (by J. Pollock19); for labeling lamellocytes, a specialized cell type mainly induced by immune challenges and injury13, e.g., MSNF9mo-mCherry may be used17. Some transgenic drivers are expressed in a range of differentiated blood cells and progenitors, such as He–GAL4 20, which labels about 80% of all larval blood cells20. Please note that in all cases where GAL4 drivers are used, combination with UAS-GFP or another fluorescent protein UAS-transgene is required. In the Results section, this method is used to monitor blood cell number and circulation behavior over the course of larval development.
Second, the Hemocyte Disturbance Assay describes a preceding step designed to detach resident hemocytes by external manipulation, which subsequently allows the evaluation of the ability of hemocytes to re-adhere and home to HPs within a limited time frame (30 – 60 min)6. Typically this assay is followed by the Bleed/Scrape Assay to determine the percentage of circulating hemocytes per larva. We present a simplified protocol for this assay (Figure 1D), which uses disturbance by vortexing with glass beads, rather than manipulation of single larva with a paint brush as described previously6. In the Results section, this assay is used to demonstrate that transiently detached hemocytes float in the hemolymph and can be recovered in the fraction of circulating hemocytes. The assay is also useful to quantify differences of hemocytes in their homing/adhesion to resident sites, comparing e.g., various genetic backgrounds or stimulation conditions. Please note that this mechanical manipulation reflects a reversible process and is distinct from infection- or injury-induced resident hemocyte mobilization, which typically are not reversible in a short time frame4,13.
1. Hemocyte Bleed/Scrape Assay
2. Hemocyte Disturbance Assay
To illustrate typical outcomes of the described methods, we first used the Hemocyte Bleed/Scrape Assay to outline the progression of larval hemocyte numbers and their residence over the course of larval development (Figure 4). Resident and circulating larval hemocyte populations were isolated from single larvae (HmlΔ-GAL4, UAS-GFP; He-GAL4 to label the vast majority of larval hemocytes) and quantified using ImageJ. Cohorts of larvae sized 1.2 mm (~48 hr AEL or 1st instar), 2.5 mm (~80 hr AEL or late 2nd instar), and 3.5 mm (~96 hr AEL or 3rd instar) were examined (Figure 4). Hemocyte numbers expanded over the course of larval development, correlating with and exceeding previous estimates based on light microscopy of dye stained larvae7 and live counting of fluorescent protein labeled hemocytes through the larval cuticle6. In 1st instar larvae almost all hemocytes were resident, while the fraction of circulating hemocytes progressively increased over the course of larval development (Figure 4B,C), consistent with previous publications6,7.
Next we examined whether the method faithfully monitors the transition of hemocytes between the resident and circulating populations. Taking advantage of the phenomenon that resident hemocytes can be transiently detached by mechanical disturbance and they re-adhere to their resident sites spontaneously6, we dispersed resident hemocytes by vortexing with glass beads as described in the Hemocyte Disturbance Assay. Indeed, mechanical disturbance of larvae led to a dramatic increase in the population of circulating hemocytes at the expense of resident hemocytes (Figure 5). After a recovery period of 45 min, hemocytes had largely returned to their adherent state, both by visual inspection and by the assessed percentage of circulating cells (Figure 5D,E). As expected, total hemocyte numbers remained stable over time, despite the shift of hemocytes between the circulating and resident populations.
Several additional considerations were taken into account. To confirm that vortexing did not cause major tissue damage, vortexing with glass beads was performed in the presence of trypan blue (Sigma) for various time periods (1, 5, 20 min). Both 1 and 5 min vortexing did not cause any obvious tissue disruption, while 20 min vortexing resulted in small areas of damage, resembling damage caused by needle stitches used as positive control (Supplemental Figure 1). While internal damage of epidermis or other tissues without cuticle damage cannot be excluded, this scenario seems rather unlikely as hemocytes of 1 min and 5 min-treated larvae re-adhered in the expected pattern and time frame, suggesting larval integrity was not compromised (Supplemental Figure 1). In contrast, larvae vortexed for 20 min suffered from a lack of re-adhesion, and did not even show attachment of circulating hemocytes to epidermal wound sites, as has been described previously14.
Lastly, to demonstrate reproducibility of the method, we compared biological replicates of 2.5 mm larvae from the above two experiments, which were conducted by distinct experimenters. As illustrated in Supplemental Figure 2, both cohorts showed comparable total numbers of hemocytes per larva, and the percentage of circulating hemocytes. Student’s t testing showed no statistically significant differences, suggesting that the method is reproducible and broadly applicable.
Figure 1. Hemocyte Bleed/Scrape and Disturbance Assay setup and schematic. (A) Single Image Slide Setup: five 2mm squares for imaging with a 5X objective. (B) Tile Scan Slide Setup: four 3 mm squares for imaging bleed/scrapes of ≤2.5 mm larvae with a tile scan microscope. Recommended objectives for imaging are 5X or 10X. (C) Bleed/Scrape Assay schematic and resulting quantifications using ImageJ. (D) In the Disturbance Assay, the hemocyte pattern is mechanically disrupted by vortexing larvae with glass beads. Larvae are allowed to recover over a period of 45 min during which hemocytes re-adhere to the Hematopoietic Pockets. The adhesive properties of hemocytes can be assessed by this method, quantifying the percentage of hemocytes in circulation after disturbance. Please click here to view a larger version of this figure.
Figure 2. Bleed/Scrape Assay to release circulating and resident hemocytes. (A) To bleed a larva, ventral incisions at the posterior and anterior ends of the larva are made (scissors symbol). (B) Hemocytes in circulation will flow out of the incisions and settle on the surface of the slide. (C) The lymph gland (LG) is located and pinned down, without puncturing it. Resident hemocytes are released by jabbing and/or scraping the larva with a needle. (D) Resident hemocytes on slide. (E,F) The Scrape process is repeated until all resident hemocytes are released. The larval carcass containing the intact lymph gland is left behind. Please click here to view a larger version of this figure.
Figure 3. Automated quantification of hemocytes using ImageJ. (A,B) After opening a hemocyte image file in ImageJ, the Lower Threshold level is adjusted to account for all the cells in the image. (C,D) Analyze Particles requires setting the cell pixel size, circularity, and the result readout format (e.g., Overlay Outlines). (E) Summary window displaying the number of hemocytes. Please click here to view a larger version of this figure.
Figure 4. Representative Results (1). Hemocyte number and resident state over the course of larval development. (A) Overview of the larval stages used; 1st instar (48 hr AEL; ~1.2 mm length); 2nd instar (80 hr AEL; 2.5 mm length); 3rd instar (96 hr AEL; ~3.5 mm length). Genotype is HmlΔ-GAL4, UAS-GFP; He-GAL4. Stages were confirmed by assessing larval mouthhooks. (B) Bar diagram of circulating and resident hemocyte numbers at the respective larval stages. (C) Percentage of circulating hemocytes. Note that the fraction of circulating hemocytes increases disproportionally over the course of larval development. (D) Total hemocytes, resulting from the sum of circulating and resident hemocytes per larva. Hemocytes were quantified using the Bleed/Scrape method; n ≥ 6 larvae/condition, error bars show standard deviation, findings confirmed in 3 independent replicate experiments. Please click here to view a larger version of this figure.
Figure 5. Representative Results (2). Effects of mechanical disturbance on hemocyte residence. (A-C) Example of a larva before and after vortexing with glass beads, followed by 45 min recovery. (A) No disturbance control; hemocytes are localized in Hematopoietic Pockets. (B) Disrupted hemocyte pattern at 0 min after vortexing larvae in a suspension of glass beads and water. (C) Hemocyte pattern at 45 min of recovery post-disturbance; many hemocytes have relocated to the Hematopoietic Pockets; note enlarged dorsal-vessel associated clusters and dorsal stripes which are predominant sites of early post-disturbance accumulation (arrows). Genotype is HmlΔ-GAL4, UAS-GFP; He-GAL4 x yw. (D) Percentage of circulating hemocytes quantified by the Bleed/Scrape method. (E) Total hemocytes, resulting from the sum of circulating and resident hemocytes per larva. n ≥ 4 larvae/condition, error bars show standard deviation, findings confirmed in 3 independent replicate experiments. Student’s t-test to confirm significance, NS (not significant), ** (p ≤ 0.05), ** (p ≤ 0.01). Please click here to view a larger version of this figure.
Here, we describe the first method to quantitatively recover resident and circulating blood cells from single Drosophila larvae, and quantify these two hemocyte populations. The protocol comprises the sequential release of circulating and resident blood cells, followed by imaging and automated cell counting. Larval resident hemocytes can be transiently mobilized into circulation by mechanical disturbance, a process that is known to be largely reversed within a 30 – 60 min recovery period6. Accordingly, this protocol was tested in two ways, (1) by assessing the total hemocyte number per larva and fraction of circulating hemocytes over the course of larval development, and (2) by experimentally dislodging resident hemocytes using an automated method, which confirmed the tight correlation of hemocyte localization and hemocyte number in the resident and circulating populations. In addition, the reproducibility of the method was demonstrated by comparing two datasets of biological replicates.
In the past, laboratories have used a range of techniques to quantify larval hemocytes6,13,21. This protocol establishes a common standard to retrieve and quantify resident and circulating blood cell populations from Drosophila larvae, providing an easily adaptable platform. The method described is critical for studies that focus on the role of resident hemocytes and their microenvironment, the Hematopoietic Pockets4-6, and is suitable to study fluorescent protein transgene-carrying Drosophila strains in wild type and genetically modified backgrounds. The protocol is also relevant for studies that focus on hemocyte mobilization after immune challenge or injury, and genetically or environmentally induced signaling that triggers mobilization of resident hemocytes or changes in total hemocyte number (reviewed in 4). It should be noted that, in cases of premature differentiation and release of hemocytes from the lymph gland, distinguishing embryonic/larval versus lymph gland lineages may be limited by the expression pattern of the fluorescent hemocyte reporter used.
The protocol presented here relies on imaging live, fluorescently-labeled hemocytes. In the future, it may be modified to permit the detection of released cells after fixation, e.g., using immunocytochemistry. In this case, the protocol may need to be adapted to ensure complete adhesion of the blood cells, for example by increasing adhesion incubation times, and adding adhesive slide coating, such as concanavalin A. Since the method allows retrieval of hemocytes and their manipulation ex vivo, it will benefit a wide range of developmental, cell biological and biochemical studies. Resident and circulating blood cells are found during all postembryonic developmental stages of Drosophila and other invertebrates22, suggesting that adaptation of this method will benefit a wide range of studies beyond the Drosophila larval hematopoietic system.
The authors have nothing to disclose.
We thank Jesper Kronhamn and Dan Hultmark, Michael Galko, and the Bloomington Stock Center for the fly stocks. Special thanks to Courtney Onodera for advice with statistical analysis. We thank Katrina Gold for critical reading of the manuscript, and Kalpana Makhijani, Katrina Gold, members of the Derynck laboratory, and members of the Nystul laboratory for discussion and comments on the manuscript. This work was supported by grants from the UCSF Program for Breakthrough Biomedical Research (PBBR), Broad Center, Hellman Foundation, American Cancer Society RSG DDC-122595, American Heart Association 13BGIA13730001, National Science Foundation 1326268, National Institutes of Health 1R01GM112083-01 and 1R56HL118726-01A1 (to K.B.).
6cm/9cm Petri dishes | One for each genotype to be evaluated | ||
Water squirt bottle | |||
Metal spoon/spatula | |||
Thin paintbrush | e.g. a "liner" | ||
Glass cavity dish | |||
PAP pen: Super PAP PEN IM3580 | Beckman Coulter | ||
Glass slides | Each slide will have 5 or more PAP PEN squares drawn on them. Size of squares depends on the imaging objective and magnification of the microscope camera; e.g. 2mm squares. | ||
Moist chamber | This will be used to prevent slides and wells from drying out: sealed container with wet paper towels lining the sides/bottom | ||
Schneider’s Drosophila cell culture media | Invitrogen | ||
Cold block | This is a metal block (a.k.a. heating block) chilled in bucket containing ice; preferably black-colored or other dark, non-reflective color | ||
Two 1ml syringes with needles (27G ½) | Becton Dickinson | For dissections. | |
Optional: Surgical spring scissors (cutting edge 2mm) | Fine Science Tools | ||
Glass beads, 212-600 micron | Sigma | ||
2 ml Eppendorf tubes | Eppendorf | One per genotype evaluating | |
Vortex Mixer | Fisher Scientific | ||
Transgenic Drosophila larvae with fluorescently marked hemocytes. Suitable transgenes include: HmlΔ-DsRed (Makhijani et al., 2011), MSNF9mo-mCherry (Tokusumi et al., 2009), BcF6-CFP and -GFP (Gajewski et al., 2007), or HmlΔ-GAL4 (Sinenko and Mathey-Prevot, 2004), Pxn-GAL4 (Stramer et al., 2005), He-GAL4 (Zettervall et al., 2004), Crq-GAL4 (by H. Agaisse (Stramer et al., 2005)), or eater-GAL4 (Tokusumi et al., 2009) combined with UAS-GFP or other fluorescent protein transgenes. | |||
Fluorescence dissecting microscope | Leica | Here: Leica M205, optional with camera, imaging software and measuring module | |
Inverted fluorescence microscope with camera attachment | Leica or Keyence | With or without tile scanning function (eg. Leica DMI series, Keyence BIOREVO BZ-9000 series) |