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.).

1. Hemocyte Bleed/Scrape Assay
<ol>
	<li>Preparation of slides:
	<ol>
		<li>Option 1 for microscopes without tile scanning function: For each larva to be analyzed, prepare one glass slide with about 5 Pap-pen wells of 2 mm squares each, corresponding to the field viewing area of the microscope; add approximately 5 - 10 &#181;l of S2 media to each (Figure 1A). Keep slides in moist chamber to prevent wells from drying out.</li>
		<li>Option 2 for microscopes with tile scanning function: For each larva to be analyzed, prepare one glass slide with 3 to 4 Pap-pen wells of ~3 - 4 mm squares each; add approximately 15 - 20 &#181;l of S2 media to each well (Figure 1B). Keep slides in moist chamber to prevent wells from drying out. 
		NOTE: The above recommended number of wells is sufficient for totals of up to 3,000 hemocytes per larva (late 2nd instar larvae, ~2.5 - 3 mm length, transgene labeling the majority of larval blood cells). When assessing larger blood cell numbers, more wells might be needed to avoid over-crowding.</li>
	</ol>
	</li>
	<li>Collection of larvae:
	<ol>
		<li>Squirt water into a fly vial containing larvae and flush larvae into a Petri dish, or scoop some food that contains larvae into a Petri dish and dilute with water using a squirt bottle.</li>
		<li>Gently pick larvae out of the Petri dish using a paintbrush and place them in water in a cavity dish or on a slide on a cold block. 
		NOTE: Larvae can be kept for a limited time in water or on a cold block; use specimens within 45 min or less to avoid larval death or unwanted effects on hemocytes.</li>
	</ol>
	</li>
	<li>Dissection:
	<ol>
		<li>Select larvae under a fluorescence microscope on a cold metal block. Measure sizes and image larvae if desired.</li>
		<li>Isolation of circulating hemocytes (&#8220;Bleed&#8221;):
		<ol>
			<li>Once larvae are selected, place one larva in the first Pap-pen well (Figure 1C, 2A).</li>
			<li>Use 2 clean needles or dissecting scissors and forceps to make an incision at both the posterior and anterior ends of the larva. To avoid disturbing resident hemocytes, it is best to make these incisions on the ventral side of the larva. For consistent results, make the incisions in the same locations for every larva. For 1st instar larvae, 1 incision (in the ventral anterior) is sufficient.</li>
			<li>Allow larva to bleed for a few seconds without any pressure or physical agitation (Figure 2A). 
			NOTE: If working on multiple larvae it is better to make these incisions for each one before proceeding to the next step to avoid keeping larvae on ice too long which could affect the samples&#8217; integrity.</li>
			<li>Gently lift the larva with the needles or forceps and dip it into the second well to rinse any remaining circulating hemocytes. After that, follow with the release of resident hemocytes.</li>
		</ol>
		</li>
		<li>Isolation of resident hemocytes (&#8220;Scrape&#8221;):
		<ol>
			<li>Gently transfer the larva to the next well (Figure 2C).</li>
			<li>Identify the lymph gland of the larva, which typically is located approximately 1/3 from the anterior end of the larva, and which may fluoresce dorsally through the larval body wall. Avoid the lymph gland while releasing resident hemocytes by pinning down the larva with a needle as near as possible to the lymph gland to avoid puncturing (Figure 2C). 
			NOTE: During normal development the maturation of lymph gland hemocytes is delayed compared to larval hemocytes, and fluorescent reporters of differentiated hemocytes may not show a signal in the lymph gland of young larvae. In these instances, less attention needs to be paid to the lymph gland as no contamination of differentiated fluorescently-labeled larval hemocytes by lymph gland hemocytes is expected.</li>
			<li>Release resident blood cells in a dissection process of scraping and/or jabbing. Use one needlepoint to effectively pin down the larva near the lymph gland (see above) or other body areas as needed. Use another needle to jab at the clusters of hemocytes that are visible through the larval body wall (Figure 2C,E), aiming to separate the hemocytes. Hemocytes can also be released in a scraping motion. However, tearing the epidermis early may release big clusters of blood cells, which could make automated counting more challenging. 
			NOTE: Depending on the age and genotype of the larva, the number of total hemocytes will vary. Distribute the release process described above over several wells to avoid overcrowding of some wells with blood cells, which could make single cell image analysis more difficult.</li>
			<li>If few hemocytes remain in the final carcass, count these hemocytes by observation through the microscope and use of a manual tally counter (Figure 2E). To facilitate counting, place the carcass on a clean area of the same slide and spread it as thinly as possible to reduce the number of optical planes.</li>
			<li>Once the dissection is complete, wait between 5 - 10 min for the cells to settle (but not necessarily adhere) before imaging the wells. Incubate the slide in a moist chamber to avoid drying, and avoid rough handling of the slides, which could disturb the settled hemocytes. 
			NOTE: When determining hemocyte counts, released cells are not fixed and the cells must be imaged shortly after dissection, preferably within 30 min after release from the larva. Depending on the volume of medium and cell properties, the vast majority of cells will have settled within 5 - 10 min, which should be confirmed by focusing through the optical planes of the medium in the well. However, only a fraction of blood cells will have adhered to the slide surface by this time, a fact that needs to be considered if modifying this protocol for cell fixation-based approaches.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Quantification:
	<ol>
		<li>Take images of the settled hemocytes under a fluorescent microscope (Figure 2B,D,F). Follow with quantification of hemocytes using ImageJ software.</li>
		<li>Prepare image for ImageJ cell counting algorithm:
		<ol>
			<li>Open image of well using ImageJ: File &#8594; Open&#160;&#8594; (locate file and select).</li>
			<li>Ensure that the image(s) is 8-bit or 16-bit. Adjust the threshold for the image by selecting Image then click Adjust and select Threshold. Observe the &#8220;Threshold window&#8221; (Figure 3A).</li>
			<li>Check the &#8220;Dark Background&#8221; option. Select &#8220;Red&#8221; and increase the Lower Threshold Level (see black arrow) until each cell in the image is marked with a red dot (cells that are not being covered will be seen in grayscale; Figure 3B). As the Lower Threshold is increased some cells will become unmarked. This can be the indicator for how far to set the Lower Threshold. 
			NOTE: Occasionally clusters of cells cannot be resolved and would be counted as one by the particle counter. In such cases, the number of cells in a cluster can be estimated by examining the image (zoom in if needed) and manual counting using a tally counter. Alternatively, the Lower Threshold can be increased to resolve clusters of cells; any unmarked cells resulting from this manipulation can then be counted using a tally counter.</li>
		</ol>
		</li>
		<li>Analyze cell number using ImageJ:
		<ol>
			<li>Launch the Particle Analyzer to count the cells (Figure 3C). Select Analyze and click on Analyze Particle. Optionally select &#8220;Overlay Outlines&#8221; to see the particles the algorithm count (Figure 3D). Alternatively, set a limit to the size or pixel area of a unit (e.g., cell, clump of cells, etc.) for the algorithm to count.</li>
			<li>Click OK. Observe a summary window with the count (Figure 3E).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Hemocyte Disturbance Assay
<ol>
	<li>To disturb hemocytes, select larvae and place them in a 2 ml microcentrifuge tube with approximately 0.5 g of glass beads (212 - 600 &#181;m) and add 0.5 ml water.</li>
	<li>Vortex the tube, by hand, at speed 10 for 1 min.</li>
	<li>Retrieve the larvae from the glass beads by spilling the contents of the microcentrifuge tube into a Petri dish and picking out the larvae with a paintbrush.</li>
	<li>For the recovery phase, place larvae in previously prepared Petri dishes with small amounts of fly food. Allow the larvae to re-establish their hemocyte pattern for a period of 45 min or as desired. 
	NOTE: Discard any larvae that have stopped moving, as they have died in the process. However, we typically see little damage after 1 min of vortexing (see below and Supplemental Figure 1).</li>
	<li>After the recovery period, continue with the Bleed/Scrape Assay as described above in Section 1.</li>
</ol>

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The authors have nothing to disclose.

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.

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 Hml&#916;-DsRed 6, Hml&#916;-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.

<table><tbody><tr><td>6 cm/9 cm Petri dishes</td><td></td><td></td><td>One for each genotype to be evaluated</td></tr><tr><td>Water squirt bottle</td><td></td><td></td><td></td></tr><tr><td>Metal spoon/spatula</td><td></td><td></td><td></td></tr><tr><td>Thin paintbrush</td><td></td><td></td><td>&lt;em&gt;e.g.,&lt;/em&gt; a &quot;liner&quot;</td></tr><tr><td>Glass cavity dish</td><td></td><td></td><td></td></tr><tr><td>PAP pen: Super PAP PEN IM3580</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Glass slides</td><td></td><td></td><td>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; &lt;em&gt;e.g.,&lt;/em&gt; 2 mm squares.</td></tr><tr><td>Moist chamber</td><td></td><td></td><td>This will be used to prevent slides and wells from drying out: sealed container with wet paper towels lining the sides/bottom</td></tr><tr><td>Schneider&amp;rsquo;s &lt;em&gt;Drosophila&lt;/em&gt; cell culture media</td><td>Invitrogen</td><td></td><td></td></tr><tr><td>Cold block</td><td></td><td></td><td>This is a metal block (a.k.a. heating block) chilled in bucket containing ice; preferably black-colored or other dark, non-reflective color</td></tr><tr><td>2 x 1 ml syringes with needles (27 G &amp;frac12;&quot;)</td><td>Becton Dickinson</td><td></td><td>For dissections.</td></tr><tr><td>Optional: Surgical spring scissors (cutting edge 2 mm)</td><td>Fine Science Tools</td><td></td><td></td></tr><tr><td>Glass beads, 212 - 600 &amp;mu;m</td><td>Sigma</td><td></td><td></td></tr><tr><td>2 ml Eppendorf tubes</td><td>Eppendorf</td><td></td><td>One per genotype evaluating</td></tr><tr><td>Vortex mixer</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Transgenic Drosophila larvae with fluorescently marked hemocytes.</td><td></td><td></td><td>Suitable transgenes include: Hml&amp;Delta;-DsRed (Makhijani &lt;em&gt;et al.,&lt;/em&gt; 2011), MSNF9mo-mCherry (Tokusumi &lt;em&gt;et al.,&lt;/em&gt; 2009), BcF6-CFP and -GFP (Gajewski &lt;em&gt;et al.,&lt;/em&gt; 2007), or Hml&amp;Delta;-GAL4 (Sinenko and Mathey-Prevot, 2004), Pxn-GAL4 (Stramer &lt;em&gt;et al.,&lt;/em&gt; 2005), He-GAL4 (Zettervall &lt;em&gt;et al.,&lt;/em&gt; 2004), Crq-GAL4 (by H. Agaisse (Stramer &lt;em&gt;et al.,&lt;/em&gt; 2005)), or eater-GAL4 (Tokusumi &lt;em&gt;et al.,&lt;/em&gt; 2009) combined with UAS-GFP or other fluorescent protein transgenes.</td></tr><tr><td>Fluorescence dissecting microscope</td><td>Leica</td><td></td><td>Here: Leica M205, optional with camera, imaging software and measuring module</td></tr><tr><td>Inverted fluorescence microscope with camera attachment</td><td>Leica or Keyence</td><td></td><td>With or without tile scanning function (&lt;em&gt;e.g.,&lt;/em&gt; Leica DMI series, Keyence BIOREVO BZ-9000 series)</td></tr></tbody></table>

assaying blood cell populations of the drosophila melanogaster larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;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.&#160;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.

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 (Hml&#916;-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&#8217;s t testing showed no statistically significant differences, suggesting that the method is reproducible and broadly applicable.
<img alt="Figure 1" src="/files/ftp_upload/52733/52733fig1highres.jpg" width="500" /> 
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 &#8804;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. <a href="https://www.jove.com/files/ftp_upload/52733/52733fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52733/52733fig2highres.jpg" width="700" /> 
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. <a href="https://www.jove.com/files/ftp_upload/52733/52733fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52733/52733fig3highres.jpg" width="600" /> 
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. <a href="https://www.jove.com/files/ftp_upload/52733/52733fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52733/52733fig4highres.jpg" width="700" /> 
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 Hml&#916;-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&#160;&#8805; 6 larvae/condition, error bars show standard deviation, findings confirmed in 3 independent replicate experiments. <a href="https://www.jove.com/files/ftp_upload/52733/52733fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52733/52733fig5highres.jpg" width="700" /> 
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 Hml&#916;-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&#160;&#8805; 4 larvae/condition, error bars show standard deviation, findings confirmed in 3 independent replicate experiments. Student&#8217;s t-test to confirm significance, NS (not significant), ** (p&#160;&#8804; 0.05), ** (p&#160;&#8804; 0.01). <a href="https://www.jove.com/files/ftp_upload/52733/52733fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Assaying Blood Cell Populations of the Drosophila melanogaster Larva at JoVE.com

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

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.

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila ...

assaying-blood-cell-populations-of-the-drosophila-melanogaster-larva

Research

JoVE Journal

Developmental Biology

12.3K Views.  University of California San Francisco. 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.

Video: Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Neuroscience

A Direct and Simple Method to Assess Drosophila melanogaster's Viability from Embryo to Adult

In Drosophila melanogaster, viability assays are used to determine the fitness of certain genetic backgrounds. Allelic variations can result in partial or complete loss of viability at different stages of development. Our lab has developed a method to assess viability in Drosophila from embryo to fully mature adult. The method relies on quantifying the number of progeny present at different stages during development, starting with hatched embryos. After embryos have been quantified, additional stages are counted, including L1/L2, pupae, and mature adults. After all stages have been examined, a statistical analysis such as the chi-square test is used to determine if there is a significant difference between the starting number of progeny (hatched embryos) and later stages culminating in the observed number of adults, thus rejecting or accepting the null hypothesis (that the number of hatched embryos will be equal to the number of larvae, pupae, and adults recorded throughout the stages of development). The primary advantage of this assay is its simplicity and accuracy, as it does not require an embryo rinse to transfer them to the food vial, avoiding losses from technical errors. Although the protocol described here does not directly examine L2/L3 larvae, additional steps can be added to account for these. Comparing the number of hatched embryos, L1, pupae, and adults can help determine if viability was compromised during the L2/L3 stages for further studies (the use of colored food helps with visual identification of larvae). Overall, this method can help Drosophila researchers and educators determine when viability is compromised during the fly life cycle. Routine assessment of stocks using this assay can prevent accumulation of secondary mutations that may affect the phenotype of the originally isolated mutant, especially if the original mutations affect fitness. For this reason, our lab maintains multiple copies of each of our Dm ime4 alleles and routinely checks the purity of each stock with this method in addition to other molecular analyses.&#160;

A Direct and Simple Method to Assess Drosophila melanogaster's Viability from Embryo to Adult

This protocol is designed to assess the viability of Drosophila at every developmental stage, from embryo to adult. The method can be used to determine and compare the viability of different genotypes or growth conditions.

In Drosophila melanogaster, viability assays are used to determine the fitness of certain genetic backgrounds. Allelic variations can result in partial or ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Biology

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous system and protects neural tissue from toxins and pathogens. Defects in the formation of this barrier have been correlated with neuropathies, and the breakdown of this barrier has been observed in many neurodegenerative diseases. Therefore, it is critical to identify the genes that regulate the formation and maintenance of the blood-brain barrier to identify potential therapeutic targets. In order to understand the exact roles these genes play in neural development, it is necessary to assay the effects of altered gene expression on the integrity of the blood-brain barrier. Many of the molecules that function in the establishment of the blood-brain barrier have been found to be conserved across eukaryotic species, including the fruit fly, Drosophila melanogaster. Fruit flies have proven to be an excellent model system for examining the molecular mechanisms regulating nervous system development and function. This protocol describes a step-by-step procedure to assay for blood-brain barrier integrity during the embryonic and larval stages of D. melanogaster development.

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Blood-brain barrier integrity is critical for nervous system function. In Drosophila melanogaster, the blood-brain barrier is formed by glial cells during late embryogenesis. This protocol describes methods to assay for blood-brain barrier formation and maintenance in D. melanogaster embryos and third instar larvae.

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous ...

Extraction of Hemocytes from Drosophila melanogaster Larvae for Microbial Infection and Analysis

During the pathogenic infection of Drosophila melanogaster, hemocytes play an important role in the immune response throughout the infection. Thus, the goal of this protocol is to develop a method to visualize the pathogen invasion in a specific immune compartment of flies, namely hemocytes. Using the method presented here, up to 3 &#215; 106 live hemocytes can be obtained from 200 Drosophila 3rd instar larvae in 30 min for ex vivo infection. Alternatively, hemocytes can be infected in vivo through injection of 3rd instar larvae followed by hemocyte extraction up to 24 h post-infection. These infected primary cells were fixed, stained, and imaged using confocal microscopy. Then, 3D representations were generated from the images to definitively show pathogen invasion. Additionally, high-quality RNA for qRT-PCR can be obtained for the detection of pathogen mRNA following infection, and sufficient protein can be extracted from these cells for Western blot analysis. Taken together, we present a method for definite reconciliation of pathogen invasion and confirmation of infection using bacterial and viral pathogen types and an efficient method for hemocyte extraction to obtain enough live hemocytes from Drosophila larvae for ex vivo and in vivo infection experiments.

Extraction of Hemocytes from Drosophila melanogaster Larvae for Microbial Infection and Analysis

This method demonstrates how to visualize pathogen invasion into insect cells with three-dimensional (3D) models. Hemocytes from Drosophila larvae were infected with viral or bacterial pathogens, either ex vivo or in vivo. Infected hemocytes were then fixed and stained for imaging with a confocal microscope and subsequent 3D cellular reconstruction.

During the pathogenic infection of Drosophila melanogaster, hemocytes play an important role in the immune response throughout the infection. Thus, the ...

Immunology and Infection

Assessing the Cellular Immune Response of the Fruit Fly, Drosophila melanogaster, Using an In Vivo Phagocytosis Assay

In all animals, innate immunity provides an immediate and robust defense against a broad spectrum of pathogens. Humoral and cellular immune responses are the main branches of innate immunity, and many of the factors regulating these responses are evolutionarily conserved between invertebrates and mammals. Phagocytosis, the central component of cellular innate immunity, is carried out by specialized blood cells of the immune system. The fruit fly, Drosophila melanogaster, has emerged as a powerful genetic model to investigate the molecular mechanisms and physiological impacts of phagocytosis in whole animals. Here we demonstrate an injection-based in vivo phagocytosis assay to quantify the particle uptake and destruction by Drosophila blood cells, hemocytes. The procedure allows researchers to precisely control the particle concentration and dose, making it possible to obtain highly reproducible results in a short amount of time. The experiment is quantitative, easy to perform, and can be applied to screen for host factors that influence pathogen recognition, uptake, and clearance.

Assessing the Cellular Immune Response of the Fruit Fly, Drosophila melanogaster, Using an In Vivo Phagocytosis Assay

This protocol describes an in vivo phagocytosis assay in adult Drosophila melanogaster to quantify phagocyte recognition and clearance of microbial infections.

In all animals, innate immunity provides an immediate and robust defense against a broad spectrum of pathogens. Humoral and cellular immune responses are ...

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

Flow cytometry has been widely used to obtain information about DNA content in a population of cells, to infer relative percentages in different cell cycle phases. This technique has been successfully extended to the mitotic tissues of the model organism Drosophila melanogaster for genetic studies of cell cycle regulation in vivo. When coupled with cell-type specific fluorescent protein expression and genetic manipulations, one can obtain detailed information about effects on cell number, cell size and cell cycle phasing in vivo. However this live-cell method has relied on the use of the cell permeable Hoechst 33342 DNA-intercalating dye, limiting users to flow cytometers equipped with a UV laser. We have modified this protocol to use a newer live-cell DNA dye, Vybrant DyeCycle Violet, compatible with the more common violet 405nm laser. The protocol presented here allows for efficient cell cycle analysis coupled with cell type, relative cell size and cell number information, in a variety of Drosophila tissues. This protocol extends the useful cell cycle analysis technique for live Drosophila tissues to a small benchtop analyzer, the Attune Acoustic Focusing Cytometer, which can be run and maintained on a single-lab scale.

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

A protocol for cell cycle analysis of live Drosophila tissues using the Attune Acoustic Focusing Cytometer is described. This protocol simultaneously provides information about relative cell size, cell number, DNA content and cell type via lineage tracing or tissue specific expression of fluorescent proteins in vivo.

Flow cytometry has  been widely used to obtain information about DNA content in a population of  cells, to infer relative percentages in different cell ...

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to recognize a wide range of particles and destroy microbial pathogens. As organisms age, this process begins to decline, yet little is known about the underlying mechanisms or the genetic basis of immunosenescence. Here, an injection based in vivo phagocytosis assay is used to assess age related changes in different aspects of phagocytosis, such as binding, engulfment, and degradation of internalized particles, by quantifying phagocytic events in hemocytes in adult Drosophila. Drosophila melanogaster has become an ideal model to investigate age related changes in innate immune function for many reasons. For one, many genetic components and functions of the innate immune response, including phagocytosis, are evolutionarily conserved between Drosophila and mammals. Because of that, results obtained from using this protocol are likely to be widely relevant to understanding the age related changes in immune function in a variety of organisms. Additionally, we note that this method provides quantitative estimates of hemocyte phagocytic ability, which could be useful for a variety of research topics, and need not be limited to studies of aging.

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

This protocol describes an in vivo assay of phagocytosis used to assess and quantify the ability of young and aged Drosophila melanogaster hemocytes to phagocytose bacteria.

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to ...

Cuticle Disruption: A Method to Collect Hemolymph from Drosophila Larvae

Source: Reimels, T. A. and Pfleger, C. M. <a href="https://www.jove.com/video/54544/" target="_blank">Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae</a>. J. Vis. Exp. (2016).
The Drosophila hemolymph serves as both blood and interstitial fluid, circulating throughout the body during the larva and adult stages. This video describes a protocol to collect larval hemolymph in order to quantify the circulating cells.

Source: Reimels, T. A. and Pfleger, C. M. Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae. J. Vis. Exp. (2016).
The Drosophila ...

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Summary

Explore More Videos

Chapters in this video

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Functional Analysis of the Larval Feeding Circuit in Drosophila

A Direct and Simple Method to Assess Drosophila melanogaster's Viability from Embryo to Adult

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Extraction of Hemocytes from Drosophila melanogaster Larvae for Microbial Infection and Analysis

Assessing the Cellular Immune Response of the Fruit Fly, Drosophila melanogaster, Using an In Vivo Phagocytosis Assay

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

Cuticle Disruption: A Method to Collect Hemolymph from Drosophila Larvae