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First published online 13 March 2007
doi: 10.1242/jcs.03420

Journal of Cell Science 120, 1209-1215 (2007)
Published by The Company of Biologists 2007
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Research Article

Crystal cell rupture after injury in Drosophila requires the JNK pathway, small GTPases and the TNF homolog Eiger

Gawa Bidla1, Mitchell S. Dushay2 and Ulrich Theopold1,*

1 Department of Molecular Biology and Functional Genomics, University of Stockholm, Svante Arrheniusväg 16-18, SE 10691 Stockholm, Sweden
2 Department of Comparative Physiology, Uppsala University, Norbyvägen 18A, SE 75236 Uppsala, Sweden

* Author for correspondence (e-mail: uli{at}molbio.su.se)

Accepted 1 February 2007


   Summary
Top
 Summary
Introduction
 Results
 Discussion
 Materials and Methods
 References
 
The prophenoloxidase-activating cascade is a key component of arthropod immunity. Drosophila prophenoloxidase is stored in crystal cells, a specialized class of blood cells from which it is released through cell rupture. Within minutes after bleeding, prophenoloxidase is activated leading to visible melanization of the clot matrix. Using crystal cell rupture and melanization as readouts to screen mutants in signal transduction pathways, we show that prophenoloxidase release requires Jun N-terminal kinase, small Rho GTPases and Eiger, the Drosophila homolog of tumor necrosis factor. We also provide evidence that in addition to microbial products, endogenous signals from dying hemocytes contribute to triggering and/or assembly of the prophenoloxidase-activating cascade, and that this process can be inhibited in vitro and in vivo using the viral apoptotic inhibitor p35. Our results provide a more comprehensive view of immune signal transduction pathways, with implications for immune reactions where cell death is used as a terminal mode of cell activation.

Key words: Innate immunity, Phenoloxidase, Apoptosis, Hemocytes, JNK, TNF


   Introduction
Top
 Summary
 Introduction
Results
 Discussion
 Materials and Methods
 References
 
Innate immune responses are a common feature of metazoan organisms, protecting against pathogens and opportunistic microbial infections. Innate responses are controlled by signaling pathways that are remarkably well conserved. Genetic analysis in Drosophila melanogaster has revealed three such conserved immune pathways; the imd/Relish, Toll and JNK pathways that contribute to induction of humoral immune effectors such as antimicrobial peptides (AMPs) (Brennan and Anderson, 2004Go; Hultmark, 2003Go; Leclerc and Reichhart, 2004Go). Immune responses are triggered by recognition of conserved microbial patterns that are detected by dedicated receptors (pattern recognition receptors, PRRs). In Drosophila, bacterial peptidoglycan has been identified as a key elicitor and regulator of immune reactions. Recognition of peptidoglycans is mediated through a family of peptidoglycan recognition proteins (PGRPs) (Girardin and Philpott, 2006Go; Royet et al., 2005Go; Steiner, 2004Go).

Melanization constitutes another important branch of invertebrate immunity. It involves the proteolytic activation of prophenoloxidase (PPO) that catalyses several reactions that contribute to crosslinking of proteins, the production of reactive intermediates with potential cytotoxic activity (Nappi and Ottaviani, 2000Go), and ultimately to the production of melanin (Cerenius and Söderhäll, 2004Go). In Drosophila, phenoloxidase was recently shown to cooperate with other immune mechanisms to fight off microbial infections (Tang et al., 2006Go). Drosophila PPO is released from specialized cells (crystal cells) (Meister, 2004Go; Rizki et al., 1985Go) and subsequently activated by assembly of a proteolytic cascade and concomitant breakdown of the serine protease inhibitor Serpin27A (Spn27A). Spn27A degradation and the production of several of the proteases present in the PPO-activating cascade (PPO-AC) depend on transcriptional induction of the corresponding genes in the fat body, the insect equivalent of the liver (De Gregorio et al., 2002Go; De Gregorio et al., 2001Go; Irving et al., 2001Go; Leclerc et al., 2006Go; Ligoxygakis et al., 2002Go). Both the Toll and the imd/relish pathways, as well as PRGP-LE, have been implicated in this transcriptional induction (De Gregorio et al., 2002Go; Ligoxygakis et al., 2002Go; Takehana et al., 2004Go).

PPO can also be activated in a more local manner, for example during the encapsulation of large intruders such as parasitoid eggs, the coagulation of hemolymph and as part of wound healing (Bidla et al., 2005Go; Rämet et al., 2002Go; Schmidt et al., 2001Go; Wertheim et al., 2005Go). However, it is not known whether this activation is regulated at the transcriptional level or post translationally, as suggested mostly through work on larger insects (Cerenius and Söderhäll, 2004Go; Leclerc and Reichhart, 2004Go). Here we show that rapid rupture of crystal cells and subsequent local melanization in the Drosophila clot depend on the JNK pathway and on Eiger, the Drosophila homolog of tumor necrosis factor (TNF). Supporting the role of the cytoskeleton during crystal cell rupture, the release of PPO is also affected in mutants with defects in small GTPases. To our knowledge, this is the first time crystal cell activation has been linked to a particular signal transduction pathway. In addition, we present evidence that endogenous signals released from crystal cells and plasmatocytes undergoing apoptosis followed by secondary necrosis may be as effective as microbial elicitors in triggering assembly and/or activation of the PPO-AC. We show that melanization can be induced in vivo by ectopic expression of the proapoptotic protein Grim, and that this Grim-induced melanization can be inhibited by the caspase inhibitor p35. Through rupture, crystal cells provide most, if not all of the components required for clot melanization in a fast and highly effective way. This ensures that PPO activation at the pre-infection stage can occur independently of microbial elicitors and transcriptional gene activation.


Figure 1
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  		Fig. 1. Crystal cell activation involves cell rupture. (A-D) Rupture of crystal cells. Crystal cells immediately (A) and at different time points (4, 6 and 8 minutes respectively) after bleeding (B-D). The cell is shown in phase contrast (A,C) and fluorescence (B,D) showing membrane structures labeled with CD8-GFP. Note that the cell membrane ruptures starting at the top of the cell (indicated by the arrow in B) and the crystals dissolve. A fat body fragment adhering to the crystal cell is indicated (*). (E) Activation of PPO in crystal cells can also be achieved by incubating larvae at 65°C for 10 minutes (e.g. Galko and Krasnow, 2004Go), leading to the appearance of melanotic spots. Phase-contrast microscopy of the melanotic spots after bleeding reveals activation of PPO in the crystals (insets show a single melanized cell (left) and a group of melanized cells (right). Note that crystal cells appear not to rupture and the crystal not dissolved, although PPO is activated. Also, this activation of PPO in the crystal is not enhanced in a Spn27A mutant background (data not shown). Bars, 10 µm.

 

   Results
Top
 Summary
 Introduction
 Results
Discussion
 Materials and Methods
 References
 
Melanization of the Drosophila clot
In Drosophila larvae, the release of PPO involves both rupture of crystal cells (Fig. 1A-D) and dissolution of the crystals (Fig. 1C,D) that contain PPO (Rizki et al., 1985Go) (Fig. 1E, see also supplementary material Movie 1). To study the activation of PPO after wounding, we prepared hemolymph clots from noninfected larvae using a previously described ex vivo method (Bidla et al., 2005Go). We found melanization to be strongest in folds of the clot matrix and around cellular fragments that were most likely derived from lysed crystal cells (Fig. 2A-D). Similarly some plasmatocytes in the clot were melanized, although less intensely than cellular fragments, whereas cells outside the clot were not affected (Fig. 2E,F). As expected, melanization of the clot correlates with phenoloxidase (PO) activity, which was measured using a dot blot assay with a PO substrate (Sorrentino et al., 2002Go) (see Fig. 2B inset). Both melanization and PO activity were detected in preparations from wild-type animals, in spite of the presence of Spn27A in the hemolymph at normal concentration. By contrast, melanization was completely inhibited when Spn27A was overexpressed either ubiquitously [using the daughterless (da) GAL4 driver, Fig. 2G,H and Table 1], or in crystal cells [using the lozenge (lz) GAL4 driver, see Table 1]. The absence of melanization after ectopic expression of Spn27A correlated with the complete loss of PO activity measured with the dot blot assay (Fig. 2H, inset). This indicates that PO activity in the clot is regulated proteolytically and can be inhibited by Spn27A if the inhibitor is present at sufficiently high concentrations. Note that Spn27A overexpression did not block coagulation per se; clot fibers still formed (Fig. 2G,H) and most of the characteristic clot components were not affected in clots from larvae overexpressing Spn27A (data not shown). This is consistent with our previous finding that PO is not required for clot formation (Bidla et al., 2005Go).


Figure 2
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  		Fig. 2. Local activation of phenoloxidase in the Drosophila clot. Drosophila hemolymph clots were prepared as described (Bidla et al., 2005Go) and examined with phase-contrast (A,C,E) and bright field microscopy (B,D,F). Melanization is mostly restricted to cell fragments (asterisks) and folds in the clot matrix (arrowheads indicate clot folds, the arrows indicate plasmatocytes). Bars, 10 µm. (E,F) Plasmatocytes inside the clot (filled arrows) show melanization beginning on their surface. Plasmatocytes outside the clot (open arrow) are not melanized. (G,H) Overexpression of Spn27A (da-GAL4 driven) leads to complete loss of melanization, although clot fibers are formed (visible in phase-contrast, also visible in bright field images G,H). The insets in B and H show PO activity analyzed with a dot-blot assay (see Materials and Methods).

 

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  		Table 1. Effect of different mutations on PO activation in the clot

 


Figure 4
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  		Fig. 4. Melanization defects in egr mutants. (A,B) an egr1/def mutant crystal cell is shown 10 minutes (A) and 60 minutes (B) after bleeding. Crystals are still visible in A (arrows) and not in B without leading to visible melanization. Bar, 10 µm. (C,D) Melanization in egr mutants was measured photometrically. (C) Melanization is reduced in egr1/def mutants compared to wild-type (wt) activity (n=3, *P=0.014). When Egr is overexpressed using he-Gal4 in an egr1 homozygous background melanization is restored to levels above wild-type activity (n=3, **P=0.007 for comparison with the wild type). (D) Melanization in UAS-egr control larvae and in larvae overexpressing Eiger in the fat body (lsp2-Gal4>UAS-egr) or in hemocytes (he-Gal4>UAS-egr). Only expression in hemocytes leads to significant induction (n=3, **P=0.006). Note that the crosses were kept at 25°C. Data are mean ± s.d.

 
Although systemic PPO activation is known to depend on microbial elicitors, plasmatocytes as well as crystal cell fragments that are melanized in the clot (Fig. 2) do not display any microbial patterns on their surface. Thus some other, nonmicrobial feature must activate PPO on these cells in the clot. Since our mutant analysis (see below) also suggested that the recognition of microbial elicitors does not play an essential role in clot melanization (Table 1), we hypothesized that PPO activation might be triggered by endogenous signals exposed on dying crystal cells and plasmatocytes. To test whether hemocytes in the clot die, we analyzed clot preparations using the TUNEL assay and found some cells were labeled (Fig. 3A,B). We confirmed this with an independent assay for apoptosis based on the exposure of phosphatidylserine on the cell surface (Fig. 3C-E). Just as in the TUNEL assay, some cells in clot preparations showed a positive reaction whereas others did not. At an early point after bleeding, a few cells were necrotic (Fig. 3C-E), and the number of these cells increased over time (data not shown). Apoptosis of hemocytes was also observed in clot preparations from lz mutants which lack crystal cells and hemolymph PO, demonstrating that programmed cell death is not restricted to crystal cells and is not a consequence of melanization (data not shown).


Figure 3
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  		Fig. 3. Hemocytes in the clot show hallmarks of apoptosis. Bright field (A) and fluorescence (B) microscopy of a TUNEL assay performed on a clot preparation showing TUNEL-positive (filled arrows) and TUNEL-negative plasmatocytes (open arrows) in the clot. The border of the clot is indicated by arrowheads. (C-E) Phase contrast (C), annexin V (D) and propidium iodide (E) labeling of plasmatocytes 7 minutes after bleeding showing live (*) as well as apoptotic (open arrow, only annexin V positive) and necrotic cells (solid arrow, reacting with both annexin V and propidium iodide).

 
Drosophila JNK, small GTPases and Eiger are required for PPO release from crystal cells
Having established a reliable assay for clot melanization, we went on to screen Drosophila strains bearing mutations in known immune pathway genes for possible effects on clot melanization (Table 1, see also Fig. 6). Our screen included Bc and lz mutants, which lack hemolymph phenoloxidase as negative controls. In hml mutants where hemolymph does not coagulate (Goto et al., 2003Go) we still observed melanization, demonstrating that clotting itself is not required for PPO activation. An RNAi mutant in a PPO-activating protease (CG3066/MP2), which is expressed in crystal cells (Castillejo-Lopez and Häcker, 2005Go; Tang et al., 2006Go), had reduced clot melanization, supporting the notion that PPO in the clot is regulated by the same proteases as in the hemolymph (see also Discussion). By contrast, clot melanization was not altered in larvae bearing mutations in genes coding for known PRRs. This included mutants in a gram-negative binding protein (GNBP1) as well as three PGRPs, including PGRP-LE, which mediates the transcriptional induction that leads to systemic activation of PPO (Table 1) (Takehana et al., 2004Go). PPO activation in the clot thus appears to be independent of transcriptional induction. This observation was also in line with results from mutants in the two main immune signaling pathways; mutants in both the Toll (spz) and the imd/Relish pathway (Relish), as well as dTAK mutants (Boutros et al., 2002Go; Park et al., 2004Go) all showed normal melanization (Table 1). Conversely, an RNAi knockdown mutant in Drosophila JNK (basket, bsk), which acts downstream of dTAK during transcriptional activation, showed a complete lack of clot melanization (Table 1 and supplementary material Fig. S1A-E). In accordance with this, most crystal cells from bsk mutants had not ruptured even 1 hour after bleeding, although at this stage some cells lysed, probably owing to incomplete inhibition of bsk by the RNAi construct (supplementary material Fig. S1A). JNK pathway involvement in clot melanization was supported by finding puc lacZ staining in crystal cells, which indicates JNK pathway activity in these cells (supplementary material Fig. S1F,G). Also, RNAi knockdown of hemipterous (hep, dJNKK), which encodes a Drosophila Jun N-terminal kinase (JNK) upstream of Bsk (Glise et al., 1995Go), prevented clot melanization (Table 1).


Figure 6
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  		Fig. 6. Drosophila signal transduction pathways involved in immunity and their contribution to crystal cell rupture. Mutants in the factors that are shown in red were tested, mutants that show defects crystal cell rupture are also underlined (see Brennan and Anderson, 2004Go; Delaney et al., 2006Go). In addition to regulating the early phase of transcriptional immune and stress responses (Boutros et al., 2002Go; Park et al., 2004Go), Bsk acts in a transcription-independent way downstream of Rho GTPases and upstream of cytoskeletal regulators. Dl/Dif, dorsal/dorsal-related immunity factor; GNBP1, gram-negative binding protein 1; IMD, immune deficiency; PGRP-LC, peptidoglycan recognition protein LC; PGRP-LE, peptidoglycan recognition protein LE; PGRP-SA, peptidoglycan recognition protein SA; TAK, HIV Tat-associated kinase; Tl, toll.

 

Small Rho GTPases are known regulators of JNK (Johndrow et al., 2004Go), so we also tested mutations in genes coding for these. We found that larvae expressing a dominant active form of RhoA (RhoA.DA) showed no clot melanization (Table 1, supplementary material Fig. S2A,B). Consistent with the lack of melanization, mutant crystal cells appeared unable to rupture and contained undissolved crystals even after prolonged incubation (supplementary material Fig. S2C,D). Although heat treatment to identify crystal cells (Galko and Krasnow, 2004Go) revealed the presence of comparable numbers of crystal cells in control and mutant larvae, melanization itself was much weaker in mutants, further supporting the idea that crystal cell activation is defective in these animals (supplementary material Fig. S2F,G, see also the Discussion for further details on the analysis of GTPase mutants). To test whether the TNF homolog Eiger, which is known to act upstream of the JNK pathway during eye development (Igaki et al., 2002Go; Moreno et al., 2002Go) is also required for crystal cell lysis, we analyzed two strongly hypomorphic egr alleles (egr1 and egr3) (Igaki et al., 2002Go) as well as transheterozygotes between the stronger allele and a deletion uncovering the same region. All egr mutants showed reduced clot melanization, with the strongest effects in the transheterozygotes (Table 1) in which we also observed defects in crystal cell rupture (Fig. 4A,B). Consistent with these histological observations, we found less PO activity in egr mutants using a photometric assay. Overexpressing Eiger in the mutant background compensated or even overcompensated this defect (Fig. 4C). Eiger overexpression in wild-type larvae showed that protein expression is sufficient in hemocytes but not in the fat body to induce melanization, since expression driven by he-GAL4, but not the fat-body-specific lsp2-GAL4 activated melanization (Fig. 4D).

Since plasmatocytes in the clot show some hallmarks of apoptosis (Fig. 3), we also tested the influence of the viral apoptosis inhibitor p35 after ectopic expression in hemocytes and found strong reduction in clot melanization comparable with the strongest egr alleles (Table 1). To obtain in vivo evidence for a role of hemocyte death during PO activation and to exclude microbial contamination as a cause of melanization in ex vivo preparations, we expressed the apoptotic inducer Grim ectopically in larval hemocytes (Wing et al., 2001Go). Consistent with our previous results, induction of apoptosis led to the formation of melanotic aggregates that were confirmed to contain hemocytes (Fig. 5A,D,E). When Grim expression was induced using heat-shock driver lines, extensive melanization was observed throughout the animal (data not shown). Just as in our ex vivo experiments (Fig. 2G,H), the number and extent of the aggregates induced in hemocytes was regulated by Spn27A, since the severity of the melanotic phenotype was enhanced in a spn27A mutant background (Fig. 5B). Conversely, when Spn27A was co-expressed with Grim in hemocytes, surviving larvae showed almost no melanization (Fig. 5C). In addition, Grim-induced melanization was completely inhibited when p35 was co-expressed (data not shown), paralleling the reduced melanization in clots from larvae that expressed p35 in hemocytes in a wild-type background (Table 1).


Figure 5
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  		Fig. 5. Induction of apoptosis activates PO in vivo in a Spn27A-dependent way. (A) Expression of the apoptotic inducer Grim together with GFP in hemocytes (he-GAL4 UAS-GFP.nls>UAS-grim) leads to formation of melanotic aggregates. Note that he is not expressed in all hemocytes, leaving some cells unaffected. (B) In a spn27a mutant background both the number and severity of the melanotic spots after expressing Grim in hemocytes is increased. (C) Spn27A co-expression with Grim in hemocytes (using the same driver but without UAS-GFPnls to compensate for GAL4 dosage in A and B) abolishes melanotic spot formation almost completely. (D,E) A preparation of an aggregate dissected from a larva such as those shown in A shows the presence of hemocytes which were labeled with GFP. Bar, 10 µm. Note that adults emerging from the cross in Fig. 5A show defects in wing morphology and co-expression of Grim and viral p35 protein abolishes the phenotypes attributable to Grim (not shown).

 

   Discussion
Top
 Summary
 Introduction
 Results
 Discussion
Materials and Methods
 References
 
Here we present evidence that the Drosophila JNK homolog Bsk, the dJNK kinase Hep, small GTPases and the TNF homolog Eiger are all involved in the lysis of crystal cells. Fig. 6 summarizes our results and presents an extended model for immune activation that incorporates these findings. The involvement of small Rho GTPases, which are mostly known as cytoskeletal regulators, the lack of an effect of imd, Rel or Toll signal pathway mutations on clot melanization, and the fast kinetics of crystal cell activation all support the idea that this process is independent of transcriptional activation. We propose that during activation of crystal cells, the JNK pathway is used solely to regulate events at the post-translational level leading to cell rupture. This differs from the induction of apoptosis in the developing eye, where a pathway that includes both Eiger and JNK activates transcription (Igaki et al., 2002Go; Moreno et al., 2002Go), and from JNK-dependent transcriptional induction of immune genes (Boutros et al., 2002Go; Delaney et al., 2006Go; Park et al., 2004Go). Morphologically, crystal cell lysis shows more similarity to necrotic forms of cell death (such as swelling, the lack of apoptotic bodies and cell rupture; see Movie 1 in supplementary material and Fig. 1) than to apoptosis. Our findings add to the growing evidence that necrotic cell death - similarly to apoptosis - can be induced endogenously as part of inflammatory and repair responses (Zong and Thompson, 2006Go). The proposed role of the JNK pathway as a cytoskeletal regulator in crystal cells is somewhat reminiscent of its role in planar cell polarity in the wng/wnt pathway (Weston and Davis, 2002Go). Similar to our findings, the role of JNK in the non-canonical Wng/Wnt pathway also involves small Rho GTPases (Weston and Davis, 2002Go). Our analysis of loss-of-function mutants in small Rho GTPases suggests that Rac2 plays a key role, since the clot did not melanize in Rac2 mutants (data not shown). Consistent with the lack of melanization in Rac2 mutant clots, capsules in Rac2 mutant larvae are also not melanized (Williams et al., 2005Go). Thus Rac2 is the most likely Rho GTPase to activate melanization in these two immune reactions. Further support for a role for small GTPases in immunity comes from the identification of Rho GTPase inhibitors as virulence factors in both prokaryotic and eukaryotic insect parasites (Avet-Rochex et al., 2005Go; Labrosse et al., 2005aGo; Labrosse et al., 2005bGo; Reineke et al., 2002Go).

In contrast to the induction of AMPs where dTAK genetically interacts with bsk (Delaney et al., 2006Go), we find that dTAK is not required for crystal cell lysis. This is supported by earlier observations on dTAK mutants that show immune defects but are otherwise perfectly viable (Vidal et al., 2001Go), whereas bsk mutants have cuticular defects and defective wound healing (Rämet et al., 2002Go).

As a result of crystal cell rupture, PPO is solubilized and forms complexes with proteases and protease-like proteins (Gupta et al., 2005Go; Piao et al., 2005Go). Possible candidates for proteases in the PPO-AC include the CG3066 product (or MP2), which is expressed in hemocytes and appears to affect clot melanization (Table 1), and other proteases downstream of CG3066, such as MP1 (Castillejo-Lopez and Häcker, 2005Go; Leclerc et al., 2006Go; Tang et al., 2006Go). We suggest that CG3066/MP2 in the clot is protected from serpins such as Spn27A in the hemolymph because the inhibitor has limited access to the clot matrix. Most, if not all components required for immediate assembly of the PPO-AC may be released from crystal cells. This ensures rapid clot melanization independently of both microbial elicitors and transcriptional activation. Only in those cases where microbes get past the clot and infect the hemocoel are immune genes activated transcriptionally. This leads to a systemic humoral response, including breakdown of Spn27A and activation of PPO even outside the clot (Ligoxygakis et al., 2002Go). Our observations indicate that assembly and/or activation of the PPO-AC in the clot may occur not only on microbial surfaces, but also on cell fragments from crystal cells themselves (Fig. 1) or on the surface of plasmatocytes (see below). Other assembly sites and activators of PPO include components from damaged basement membranes or damaged cells at the wound site (Brennan and Anderson, 2004Go; Schmidt et al., 2001Go).

In addition to rupturing crystal cells, plasmatocytes in the clot also undergo a form of programmed cell death. We demonstrate the presence of some of the hallmarks of apoptosis such as exposure of PS and DNA fragmentation in plasmatocytes within the clot (Fig. 4), although it appears that the apoptotic cells quickly start swelling and their membranes become leaky (Fig. 4C-E). We assume that apoptosis of plasmatocytes is soon followed by secondary necrosis (Zong and Thompson, 2006Go). A role for apoptosis during activation of PPO is also supported by our ability to inhibit clot melanization in histological preparations (Table 1) and Grim-induced melanization (Fig. 5) in vivo using the caspase inhibitor p35 (not shown). Note that apoptotic cell death is part of many developmental processes, and that in many cases apoptosis per se does not lead to melanization. Possible explanations for the different consequences of apoptotic death are that apoptotic cells are not exposed to the hemolymph during development, that additional signals are required to activate PPO, or that melanization is triggered in the clot only when apoptosis is followed by secondary necrosis. At present, we favor the last explanation. We propose that plasmatocyte death is initially apoptotic, since this ensures exposure of PS, attracting the PPO-AC, and subsequently necrotic. This would be an effective way to eliminate potential host cells for intracellular parasites that may enter via the wound. In this respect there are striking parallels between clot melanization and plant resistance responses which also include the production of reactive oxidative intermediates used to kill bacteria and reinforce extracellular matrices, as well as the induction of programmed cell death that is difficult to categorize as either apoptotic or necrotic (Chisholm et al., 2006Go).

The assembly of the PPO-AC on cell-derived membrane vesicles resembles the assembly of the mammalian prothrombinase complex, which also occurs on PS-containing surfaces derived from platelets and their fragments (Heemskerk et al., 2002Go). We hypothesize that in wounded Drosophila larvae, formation and activation of the PPO-AC plays a similar role to vertebrate secondary hemostasis, whereas the formation of the clot itself is equivalent to primary hemostasis. Both processes together ensure proper wound closure and prevent dissemination of microbial intruders (Herwald et al., 2004Go; Scherfer et al., 2006Go; Sun et al., 2004Go).

In summary our results show that in spite of the obvious differences between insects and vertebrates such as the lack of a PPO-AC in the latter, there are striking similarities at the cellular level. The identification of the pathways involved in the rupture of crystal cells and subsequent release of the PPO-AC may have implications for the analysis of the terminal activation of other immune cells such as neutrophils and eosinophils, which can also involve cytolysis (Brinkmann et al., 2004Go; Erjefält and Persson, 2000Go).


   Materials and Methods
Top
 Summary
 Introduction
 Results
 Discussion
 Materials and Methods
References
 
Fly stocks
Flies were kept at 25°C in a 12 hour light/12 hour dark cycle on potato sucrose medium. Washed third instar larvae were used. The Drosophila strains used were: w1118 (wild type), y1 w*; P{UAS-mCD8::GFP.L}LL5 (GFP-tagged mouse transmembrane protein CD8, which labels the cell surface), w*; wgSp-1/CyO; ry506Sb1P{{Delta}2-3}99B/TM6B,Tb+ (transposase stock), P{GawB}lzgal4, P{UAS-GFP.S65T}T2, P{lArB}pucA251.1F3 ry506/TM3, Sb1 (JNK pathway reporter), Df(2R)stan1, P{neoFRT}42D cn1 sp1/CyO (deficiency that uncovers egr), w*; P{UAS-p35.H}BH1, w*; P{UAS-p35.H}BH2, Act5C-GAL4/TM6B, Tb1, hmlf03374 and Bc1 (Bloomington Stock Center); dTAK1 (Vidal et al., 2001Go), UAS-bsk-IR (Ishimaru et al., 2004Go) and daGAL4 (kindly provided by B. Lemaitre, Gif-sur-Yvette); UAS-spn27A (Ligoxygakis et al., 2002Go), GNPB-1osi, PGRP-SAseml and PGRP-LCE12 (Gobert et al., 2003Go) (kindly provided by J.-M. Reichhart, Strassbourg); Hemese-GAL4, Hemese-GAL4 UAS-GFP.nls (Zettervall et al., 2004Go) (kindly provided by D. Hultmark, Umeå); PGRP-LE112 (Takehana et al., 2004Go) (kindly provided by S. Kurata, Sendai); lzr15/XÙX (kindly provided by M. Meister, Strassbourg); UAS-RhoA V14/CyO-GFP (Billuart et al., 2001Go), CG3066-IR (Castillejo-Lopez and Häcker, 2005Go), UAS-grim (Wing et al., 2001Go) and egr1/egr1, egr3/egr3, UAS-egr (Igaki et al., 2002Go) (kindly provided by M. Miura, Tokyo). Flies bearing a UAS-grim and UAS-egr insertion on chromosome three were generated by standard genetic techniques after crossing UAS-grim (Wing et al., 2001Go) and UAS-egr (Igaki et al., 2002Go), respectively to the transposase stock. UAS-grim, UAS-spn27A flies were obtained by recombining UAS-grim and UAS-spn27A (Ligoxygakis et al., 2002Go) on the third chromosome. Gal4>UAS experiments were performed at 29°C unless noted otherwise.

Histology
The preparation of clot samples was essentially as previously described (Bidla et al., 2005Go). Hemolymph from five larvae was incubated upside down for 30 minutes in a humid chamber at 25°C. The clot was then caught on a coverslip, and images were taken with a COOLPIX 4500 digital camera (Nikon) adapted to a Leitz Orthoplan microscope.

PO activation
For the measurement of PO activity by dot blots, 5 µl hemolymph was applied to a filter paper pre-soaked with 20 mM DOPA in phosphate buffer pH 6.6 (Sorrentino et al., 2002Go), incubated for 20 minutes at 37°C and briefly heated in a microwave to dry the paper. For photometric measurement of PO activity, 30 µl hemolymph from each strain was pooled on ice by quickly bleeding 10-12 larvae and withdrawing 6 µl hemolymph each time into 10 µl of insect Ringer with EDTA instead of Ca2+ (anticoagulant) (Scherfer et al., 2004Go). After pipette mixing, 8 µl of each hemolymph was aliquoted and activated at 25°C for 10 minutes. Optical density (OD) was read at 490 nm with a VmaxTM Kinetic Microplate Reader after adding 25 µl DOPA at 10 and 30 minutes. Activation of PO was measured as the relative change in OD. Experiments were repeated at least three times and sets of experiments at least twice. Data were compared using a two-tailed Student's t-test (see figure legends for further details).

Apoptosis assays
Apoptosis was detected by TUNEL and annexin V binding assays. In TUNEL, DNA fragmentation was detected in hemocytes in a hanging drop preparation of hemolymph (see above) using the Roche Diagnostics in situ cell death detection kit, with fluorescein, following the manufacturer's instruction. Annexin V binding was performed using the CLONTECH ApoAlert Annexin V-FITC Apoptosis kit. Hemolymph from five third instar larvae was bled into 100 µl drop of Drosophila Ringer's solution on a well of a ten-well multitest slide. After incubation for 7 minutes, attached hemocytes were rinsed three times with annexin V binding buffer and continued as recommended by the supplier with the slight modification that 1.5 times more annexin V and four times less propidium iodide were used.

beta-Galactosidase activity
Three pucA251-lacZ larvae were bled into 100 µl anticoagulant Ringer's buffer and incubated at room temperature for 90 minutes in a humid chamber followed by a 10-minute fixation in 4% paraformaldehyde. Hemocytes were then permeabilized in 0.5% Triton-X100 for 5 minutes and stained with 5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside (X-Gal) for 18 hours.


   Acknowledgments
 
This work was supported by research grants from the Swedish Research Council, (U.T. and M.D.), Stockholm University (U.T.) and Södertörns Högskola (M.D.). We thank Ylva Engström, Mathias Hornef and Patrick Young for critically reading the manuscript and an anonymous reviewer for useful comments.


   Footnotes
 
Supplementary material available online at http://jcs.biologists.org/cgi/content/full/120/7/1209/DC1


   References
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 Summary
 Introduction
 Results
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 Materials and Methods
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