1—Identification and Characterisation of the hemotin Gene

We originally identified hemo in a bioinformatics search for putative functional smORFs in the Drosophila melanogaster genome [23]. hemo encodes a putative transmembrane peptide of 88 amino acids that matches a putative peptide fragment from a proteomics study of Drosophila S2 cell membrane extracts (S1A and S1B Fig) [24]. hemo originally mapped to a region of the genome devoid of any annotated gene, but subsequently, this locus has been annotated by the Drosophila Genome Project, and its structure is shown in Fig 1A. A potential polycistronic RNA is expressed by this locus and contains the 88aa hemo-ORF and a second short ORF (Open Reading Frame) of 59 codons (ORF2), currently annotated in Flybase as CG43194 and CG43210, respectively. Poly-Ribo-Seq data from Drosophila S2 cells shows that hemo transcripts are actively translated (Fig 1B) [3].

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Fig 1. Identification and phenotypical characterisation of the hemotin gene.

(A) hemo genomic locus including the hemo, CG7691, fray, and fruitless genes (blue arrows). The hemo^A4 deletion (red bar) was generated by FRT-mediated recombination using the P{RS3}fray^CB-0706-3 and the P-Bac{WH}fru^f02684 transposable elements (blue triangles). Transcript models are represented under their respective genes, orange boxes represent coding exons, whereas gray boxes indicate noncoding exons (untranslated regions, UTRs). hemo^A4 completely removes hemo and CG7691 plus the first noncoding exons of fray and fruitless. The P{PZ}fray⁰⁷⁵⁵¹ insertion is a lethal fray allele [25] and was used for genetic complementation experiments between hemo^A4 and fray. (B) Top: Ribosomal profiling reads obtained from polyribosomes from S2 cells (Poly-Riboseq;(3)) mapped to the hemo full-length transcript (hemoFL). hemo-ORF is translated more efficiently than ORF2 (hemo-ORF RPKM: 29.4, coverage: 0.9 ORF; ORF2 RPKM: 6.6, coverage: 0.7. Note that the reads per kilobase of transcript per million mapped reads [RPKM] value of ORF2 is below the 11.8 cut-off to be considered translated [3]). Bottom: schematic representation of other constructs used in this manuscript. hemo-ORF is a minigene consisting of an mRNA fragment truncated immediately after the hemo-ORF stop codon, ORF2 consists of a mini-gene construct carrying the ORF2 sequence only, including 6 nt upstream of its start codon (to conserve its endogenous Kozak sequence). hemo-GFP (green fluorescent protein) is a hemo-ORF-GFP fusion construct in which the GFP sequence (devoid of a start codon) was cloned into the hemoFL construct, immediately downstream and in frame with hemo-ORF (devoid of a stop codon) (see Materials and Methods). (C) Pattern of expression of hemo in germ band-retracted embryos revealed by in situ hybridisation. hemo is specifically expressed in embryonic hemocytes (arrows; compare with D) in the head, amnioserosa, and dispersed along the body. (D) Spatial distribution of embryonic hemocytes at germ band retraction stage revealed by in situ hybridisation of hemocyte-specific croquemort (crq) gene, showing similar distribution in the head, amnioserosa, and along the body (arrows). (E) Cluster of early embryonic hemocytes of the cephalic region expressing the hemo transcript revealed by FISH (fluorescent in situ hybridisation). Some hemocytes show drop-shape morphologies (asterisk) and membrane projections such as filopodia (arrows). (F) Embryonic hemocytes labelled with crq-Gal4;UAS-GFP expression from the head region displaying similar cellular morphologies (arrows and asterisk) as those in E. (G–H) White prepupal thoracic hemocytes revealed by crq-Gal4;UAS-GFP expression in wild-type (G) and hemo^A4 mutants (H). In hemo^A4 mutants, hemocytes display enlarged vacuoles within the cytoplasm (arrowheads), with larger occupied area index (OAI). Scale bar (50 μm). (I–N) hemocytes observed ex vivo [15] showing Tubulin (green) and Actin (red) cytoskeletons and nuclei (2-(4-Amidinophenyl)-6-indolecarbamidine dihydrochloride, DAPI) with its corresponding orthogonal projection of confocal microscopy z-stacks (above inset) showing only tubulin cytoskeleton (green) and DAPI (blue) staining in the nucleus (n). Scale bar (5 μm). (I) Wild-type hemocyte. (J) hemo^A4 mutant hemocyte shows large disruptions of the tubulin cytoskeleton that appear as rounded vacuoles (arrows; arrowhead in inset). (K) Knocking down the expression of hemo with a UAS-hemo-RNAi construct phenocopies the vacuolation phenotype (arrows and arrowhead in inset). (L) Expression of hemo full length transcript (UAS-hemoFL) rescues the vacuolated hemo^A4 phenotype. Expression of hemo-ORF only (M) also rescues the hemo^A4 mutant vacuolation. (N) Expression of ORF2 does not rescue the hemo^A4 mutant vacuolated phenotype (arrows and arrowhead in inset). (O) Vacuolation measurements in ex vivo primary pre pupal hemocytes. hemo^A4 mutant hemocytes show significantly higher occupied volume index (OVI) (see Materials and Methods) than wild-type. Rescue experiments show that the vacuolation phenotype is specific to the peptide encoded by hemo-ORF. All upstream activating sequence (UAS) constructs were driven by crq-Gal4. Error bars represent standard error of the mean (SEM). Statistical analysis was performed using one-way ANOVA test indicating that samples were significantly different [F(9,486) = 9.5, p < 0.0001]. A post hoc Bonferroni multicomparison test showed that hemo^A4, UAS-hemo-RNAi, UAS-ORF2-hemo^A4, UAS-hemoFS (expressing a hemo full-length transcript containing frameshifts in hemo-ORF and ORF2)-hemo^A4 and CG7691 genomic fragment (GF)-hemo^A4 were significantly different than wild-type. The UAS-hemoFL-hemo^A4,UAS-hemo-ORF-hemo^A4,UAS-hemoGFP-hemo^A4 and fray^PZ/hemo^A4 were not significant to wild-type (n ≥ 24, p < 0.05). Supplemental data are shown in S1 Fig and S1 Data.

https://doi.org/10.1371/journal.pbio.1002395.g001

We assayed expression of hemo throughout development by reverse transcription polymerase chain reaction (RT-PCR) and found that transcription of hemo is temporally regulated, showing higher expression from prepupal to adult stages, in agreement with modEncode RNAseq data (S1 File;Flybase). However, in situ hybridisation also showed that hemo is specifically expressed in Drosophila embryos in a pattern similar to crq, a hemocyte-specific marker (Fig 1C–1F) [26–28]. Accordingly, hemo was detected by RT-PCR from prepupal hemocyte RNA extractions (S1C Fig). Hemocytes are fly macrophages involved in the engulfment of cellular corpses throughout development, and they are the cellular branch of the fly innate immune system [12].

To elucidate the function of hemo, we generated a deletion of the locus by FRT-mediated recombination using the P(RS3)fray CB-0706-3 and PBac(WH)fru f02684 P-element insertions [29]. This deletion (hemo^A4) removes the entire hemo locus, CG7691, and also one 5’ noncoding exon of each of the frayed (fray) and fruitless (fru) genes (Fig 1A and S1C Fig). Homozygous hemo^A4 mutants reached adulthood and did not show overt morphological defects; however, their hemocytes exhibited a more vacuolated morphology in comparison to wild-type controls (Fig 1G and 1H and S1D and S1E Fig). The vacuolated phenotype in hemo^A4 mutant hemocytes was reflected in the microtubule cytoskeleton, and the volume of the vacuoles was significantly larger than those present in wild-type hemocytes (Fig 1I, 1J and 1O).

Further genetic analysis corroborates that this hemocyte phenotype in hemo^A4 is specific to loss of the Hemotin peptide. The fru and fray genes have functions in dimorphic sexual behaviour and nerve fasciculation, respectively [25,30]. We discarded them as candidates for providing the requirement for hemocyte vacuolation since fru was not expressed in hemocytes (S1 File), and the function of fray is provided by a shorter transcript containing the coding exons (Fig 1A), whose expression was not affected in the hemo^A4 deletion (S1 File). In agreement with this, the hemo^A4 and fray^PZ07551 mutations (Fig 1A) [25] complemented each other giving rise to hemocytes with normal vacuoles (Fig 1O) and larvae with normally fasciculated nerves (S2 File). For the CG7691 gene, we generated a 12 Kb genomic construct containing its coding and upstream regulatory regions. Hemocytes carrying this CG7691 genomic fragment (GF) in a hemo^A4 mutant background expressed CG7691 at wild-type levels (S1 File) but still possessed large vacuoles (Fig 1O), indicating that deletion of CG7691 in hemo^A4 is not the cause of hemocyte vacuolation. In contrast, loss of function and rescue experiments pinpoint the hemo transcript as responsible for these vacuolation phenotypes. First, reduction of hemo RNA expression with a ds-RNA construct (UAS-hemo-RNAi) in hemocytes produced enlarged vacuoles mimicking the hemo^A4 phenotype (Fig 1K and 1O). Second, expression of the hemo full-length transcript rescued hemo^A4 phenotypes (Fig 1L and 1O), whereas expression of the same construct carrying a frame-shift in each ORF, which produces scrambled peptides, did not (Fig 1O).

Next, we assessed the contribution of each ORF to hemo function. Expression of UAS-minigenes containing hemo-ORF alone (Fig 1M and 1O) or hemo-ORF-GFP (green fluorescent protein-tagged peptides (hemo-GFP) (Fig 1O) rescued the vacuolated hemo^A4 phenotype, whereas UAS-ORF2 did not (Fig 1B, 1N and 1O), indicating that the peptide encoded by hemo-ORF represents the functional unit of the gene in this context. Poly-Ribo-Seq data from Drosophila S2 cells [3] indicated high translation of hemo-ORF but a lower, or non-productive, rate of ORF2 translation (Fig 1B). Finally, carboxyl terminal-tagged GFP fusions of each peptide in a full-length transcript only showed expression of Hemo-GFP, which localized to intracellular membrane structures in S2 cells (S2 File) and Kc167 cells (S1F–S1F” Fig). We could only detect ORF2-GFP peptides from an ORF2-GFP fusion in a short transcript minigene that excludes hemo-ORF (Fig 1B; S2 File). Thus, although ORF2 does show potential for translation, altogether our results support that the 88aa transmembrane peptide encoded by hemo-ORF alone provides the requirement for normal vacuolation in Drosophila hemocytes.

2—Hemotin Peptides Are Required for Endolysosomal Maturation and Resistance to Infection

We next investigated the origin of the mutant vacuoles. hemo^A4 hemocytes showed an increased accumulation of acidic compartments as revealed by pH-sensitive Lysotracker (S1G, S1H and S1K Fig), and vacuoles disrupting the microtubule cytoskeleton contained the lysosomal marker LAMP1 (Figs 2A–2A” and S2A–S2A”). Expression of hemo full-length or hemo-ORF constructs in hemo^A4 mutants rescued the enlarged acidic compartment phenotype (S1I–S1K Fig). These pieces of evidence suggest that the enlarged vacuoles are some type of abnormal degradation compartment and that hemo is necessary for the processing of phagocytic or recycled materials.

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Fig 2. The Hemotin peptide is required for proper endosomal maturation in hemocytes.

(A–A”) Distribution of acidic organelles in hemo^A4 mutant ex vivo hemocytes revealed by the expression of LAMP1-GFP lysosomal marker. The intracellular vacuoles that disrupt the beta-tubulin cytoskeleton (A, A”; red) accumulate LAMP1-GFP positive compartments (A, A’; green). Compare with wild-type in S2A–S2A” Fig. Scale bar (5 μm). (B–B”) Distribution of the endosomal marker FYVE (PI(3)P binding zinc finger domain, early endosomal marker, named after being found in Fab1, YOTP, Vac1, EEA1) (green) (B, B’) and Lysotracker (red) (B, B”) organelles in a hemo^A4 mutant ex vivo hemocyte showing enlarged intracellular compartments coexpressing FYVE and Lysotracker. Scale bar (5 μm). (C–C”) Wild-type ex vivo hemocyte labelled as in (B), showing little overlap between early endosome-FYVE positive (green) (C,C’) and lysosomal (red) (C,C”) compartments. (D) Quantification of the FYVE OAI in ex vivo hemocytes (see Materials and Methods). hemo^A4 mutants display a significantly larger FYVE area than wild-type. This phenotype is rescued by the expression of the hemo full length transcript (UAS-hemoFL) and is specific to hemo-ORF function, as it is also rescued by the expression of the hemo-ORF mini gene (UAS-hemo-ORF) or C-terminal-tagged hemo-GFP (UAS-hemo-GFP). No rescue was observed by a CG7691 genomic fragment (CG7691-GF), or with a hemo full-length transcript containing a frameshift in the hemo-ORF (UAS-hemoFS), or with the ORF2 mini gene (UAS-ORF2). All UAS constructs were driven with He-Gal4. Error bars represent SEM. One-way ANOVA analysis shows that there is a statistically significant difference between these groups [F(7,286) = 27.12, p < 0.0001]. Post hoc comparisons using Bonferroni test indicated that the mean score of hemo^A4, UAS-hemo-ORF2-hemo^A4, UAS-hemoFS-hemo^A4, and CG7691-GF-hemo^A4 did significantly differ from wild-type (p < 0.05), whereas UAS-hemoFL-hemo^A4, UAS-hemoORF-hemo^A4, and UAS-hemoGFP-hemo^A4 did not. (E) Analysis of the overlap between FYVE-positive early endosomal and Lysotracker-positive compartments using Pearson’s correlation coefficient in wild-type and hemo^A4 mutant hemocytes. In hemo^A4 hemocytes, there exist significantly more intracellular compartments displaying FYVE and Lysotracker colocalisation than in the wild-type as shown by a Two-tailed Mann-Whitney test (n ≥ 17; p < 0.05). Error bars represent SEM. (F) Statistical analysis of the Pearson’s coefficient measurements of Hemotin-GFP (Hemo-GFP) with early endosomal (FYVE-cherry) and lysosomal (Lysotracker) markers. Tagged-Hemotin peptides are significantly enriched in early endosomal compartments in comparison with lysosomes as shown by a two tailed Mann-Whitney test (n ≥ 20, p < 0.05). Error bars represent SEM. (G–G”) Localisation of Hemo-GFP peptides (green)(G’) and the endosome FYVE marker (red)(G”) in hemocytes (He-Gal4;UAS-hemo-GFP). A substantial part of Hemo-GFP pattern colocalizes with FYVE-positive compartments (G) (arrows). Scale bar (5 μm). (H–H”) Distribution of Hemo-GFP peptides (green) (H’) and the lysosomal marker (lysotracker; red) (H”) in hemocytes. Only a small overlap exists between Hemo-GFP compartments and lysosomes (H) (arrow). Supplemental data are shown in S2 Fig, S6 Fig, and S1 Data.

https://doi.org/10.1371/journal.pbio.1002395.g002

We have used further endocytic markers to ascertain the nature and integrity of the acidic compartments in hemo^A4 null hemocytes. The hemo^A4 enlarged acidic intracellular organelles show an extensive overlap of early and late endocytic markers: FYVE (a PI(3)P-binding zinc finger domain, early endosomal marker, named after being found in Fab1, YOTP, Vac1, EEA1 [31]) (Fig 2B and 2B’ and 2E and S2E and S2E’ Fig), Rab7 (late endosomal marker) (S2C–S2C’, S2E–S2E’ Fig), and Lysotracker (acidic organelle marker) (Fig 2B, 2B” and 2E and S2C and S2C” Fig) in comparison with wild-type hemocytes (Fig 2C–2C” and 2E and S2B–S2B” and S2D–S2D” Fig). This combination of early and late markers identifies the enlarged compartments as aberrant endolysosomes [21]. We quantified the occupied FYVE area index (see Materials and Methods; Fig 2D) and the average diameter of FYVE vesicles (see Materials and Methods; S6 Fig). hemo^A4 mutant hemocytes showed larger FYVE compartments containing larger vesicles than wild-type. Expression of UAS-hemo full-length transcript, UAS-hemo-ORF, and UAS-hemo-GFP constructs rescued these phenotypes, whereas expression of a UAS-hemo frame-shift, UAS-ORF2 and a CG7691 GF constructs failed to do so (Fig 2D). The simplest interpretation of our results is that Hemotin peptides are required for completing the maturation of at least some early-to-late endosomes, and hence for their subsequent endolysosomal degradation.

Other disruptions of this traffic could also perturb endosomal maturation, since endolysosomal homeostasis depends on the balance between trafficking of membrane from the endocytic pathway (influx) and exit of membrane (efflux) into early recycling compartments [18]. We explored whether the membrane efflux to early recycling compartments was disrupted in hemo^A4 mutants by visualizing Rab11, a marker of recycling endosomes. In hemo^A4 mutant hemocytes, the size of Rab11-positive compartments seemed normal in comparison with wild-type (S2F–S2F’ and S2G–S2G’ Fig), suggesting that membrane efflux to early recycling compartments is not grossly affected. Thus, the role of Hemotin peptides seems to focus at promoting early to late endosomal maturation.

To corroborate this hypothesis, we ascertained the precise subcellular localisation of Hemotin peptides in hemocytes. The Hemo-GFP peptides, which are functional and able to rescue hemo^A4 mutants (Fig 1B and 1O and Fig 2D), localised to early endosomes, as shown by their colocalization with the FYVE marker (Fig 2F and 2G–2G”) and HRS proteins [32] (S2H–S2H” Fig) but only limited colocalization with the late endocytic marker Lysotracker (Fig 2F and 2H–2H”). Thus, the in vivo Hemotin peptide localization correlates with their inferred role in promoting endosomal maturation.

Hemotin’s requirement in endolysosomal maturation could be related to the digestion of phagocytic materials and hence have an effect on immune defence and individual survival. We analysed ex vivo the ability of hemocytes to phagocytise and digest bacteria by monitoring uptake of pHrodo-E. coli bacterial particles, which fluoresce in acidic compartments such as endolysosomes, while simultaneously revealing early endosomes with the FYVE marker (Fig 3A–3E; S1 and S2 Videos). hemo^A4 mutant hemocytes internalised pHrodo particles at similar rate to wild-type hemocytes (S3A Fig), indicating that the initial phagocytic uptake is not affected. However, in the hemo mutants, the internalised particles acidified at a slower rate and to a lesser degree, and remained for longer in FYVE-positive vesicles (Fig 3A–3E), suggesting a reduced ability of hemo^A4 mutant hemocytes to digest phagocytosed material. Such impairment could compromise the mutant fly’s ability to deal with invading and commensal microorganisms, especially for those usually cleared through phagocytosis by hemocytes [33,34]. Indeed, hemo^A4 mutant hemocytes also showed in vivo a similar normal uptake but slower processing of intact E. coli bacteria expressing mCherry (Fig 3F–3J), and homozygous hemo^A4 mutants carried a higher bacterial load than wild-type flies raised simultaneously in the same vial (Fig 3K and 3L). This increased bacterial load seems to affect lifespan, since hemo mutants had a median life span that is only 47% of the wild-type (Fig 3M) in normal cultures, but this increases to 77% when raised in germ-reduced media supplemented with antibiotics (Fig 3M). Furthermore, hemo^A4 mutant flies show reduced resistance when infected with normally nonpathogenic E. coli bacteria, comparable to other mutants that have a reduced capacity for phagocytosis [33,34] (S3B and S3D Fig).

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Fig 3. hemotin is involved in phagocytic processing and is necessary for optimal bacterial clearance and life span.

(A–B) Time-lapse imaging showing the phagocytic trafficking of pHrodo-labelled bacterial particles (arrowhead, red) in ex vivo hemocytes expressing the early endosomal marker FYVE-GFP (green) with the He-Gal4 driver in wild-type (A) or hemo^A4 mutants (B) (see S1 and S2 Videos). (t0) represents the time when the particle docks into the cell membrane, displaying a relatively faint intensity. By t = 6 minutes (min), the pHrodo particles are in FYVE-positive early endocytic vesicles in both wild-type (A) and hemo^A4 mutant hemocytes (B). By t = 22 min in the wild-type hemocyte (A), the FYVE signal around the particle is dramatically reduced, while the intensity of the pHrodo signal increases, suggesting that the vesicle has progressed into a PI3P-depeleted and acidified endolysosome. In the hemo^A4 mutant hemocyte (B), the FYVE signal around the particle is still visible at t = 44 min, showing an increased prevalence of PI(3)P in this vesicle, and therefore an extended early endocytic phase. However, the intensity of the pHrodo signal is lower than in wild-type, indicating a delay in the acidification of the endocytic vesicles (See also C and D). The insets show a magnification of the specific particles. Scale bars = 5 μm. (C) Magnified raw images from the particles shown in (A) and (B), showing the region of interest (ROI) used to quantify their fluorescent intensity. The integrated intensity read-out for each time point is indicated in red. (D) Quantification of fluorescent intensity of pH-sensitive pHrodo particles undergoing phagocytosis in wild-type (blue) or hemo^A4 mutant (red) hemocytes. Average integrated intensity per pixel is represented for each time point. Error bars represent SEM. A two-way ANOVA analysis with Bonferroni post test indicates that the difference between these curves is significantly different from t = 16.5 min (n = 8, p < 0.001). (E) Quantification of FYVE prevalence on pHrodo particles undergoing phagocytosis in wild-type (blue) or hemo^A4 mutant (red) hemocytes. The FYVE signal remains significantly longer in hemo^A4 mutants (mean = 46.06 +/− 5.3 min) than wild-type (mean = 19.87 +/− 2.1 min) as indicated by a one-tailed unpaired t test (n = 11, p < 0.0003). Error bars represent SEM. (F–I) Visualisation of bacterial up-take and processing in vivo. Dorsal vessel-associated adult hemocytes (see Materials and Methods) expressing FYVE-GFP (green) driven by He-Gal4 from wild-type (F, G) or hemo^A4 mutant flies (H, I) infected with mCherry-expressing E. coli bacteria (red) (OD600 = 0.05) and dissected 20 min (F, H) or 120 min (G, I) postinjection. Wild-type and mutant flies exhibit a similar number of bacterial cells per hemocyte after 20 min; however, this number increases in mutant flies after 120 min, whereas it remains constant in wild-type flies, suggesting that mutant hemocytes accumulate undigested bacterial cells. Yellow dashed lines represent the cell body outline. (J) Quantification of bacterial uptake. hemo^A4 mutant hemocytes contain a similar number of bacterial cells as wild-type after 20 min but significantly more bacterial cells after 120 min. Average number of bacterial cells per hemocyte is represented in the y-axis. (n = 30, p < 0.0001). (K) Visualisation of bacterial load from either wild-type or hemo^A4 mutant 30days (d)old adult fly homogenates. Each spot represents the bacterial colonies grown from an individual male fly. Each homogenate was plated in decreasing dilution (1:1 or 1:10). Note the higher density of bacterial colonies in the spots from hemo^A4 mutants compared to wild-type. (L) Quantification of bacterial colonies grown from single adult fly homogenates. hemo^A4 mutants contain significantly more bacteria than wild-type, both at 10 (n = 12, p < 0.0054) or 30 d old (n = 8, p < 0.0078). Error bars represent SEM. (M) hemo^A4 mutants (red) have a reduced viability over time compared to wild-type (blue), with a median life-span of 23 +/− 6 d, compared to 49 +/− 3 d for wild-type. The addition of antibiotics (penicillin-streptomycin) to the food media significantly increases the median life span of hemo^A4 mutants to 38 +/− 6 d, as determined by a paired t test (p < 0.05). For each condition, five different replicates were analysed for a total of 50 flies. Error bars represent SEM. Supplemental data are shown in S3 Fig, S1 Video, S2 Video, and S1 Data.

https://doi.org/10.1371/journal.pbio.1002395.g003

These results suggest that hemo mutants have a reduced cellular immunity. However, their humoral immunity (driven by antibacterial peptides [13]) does not seem affected. hemo^A4 median life span after infection with pathogenic bacteria such as Micrococcus luteus and Enterobacter cloacae, which overcome hemocytes but fully engage the humoral production of antibacterial peptides [13,33,34], was not significantly affected (S3B, S3C and S3D Fig), and accordingly the production of antibacterial peptides in hemo^A4 mutants was not impaired (S1 File).

Thus, our results suggest that Hemotin peptides in the early endosomes of fly macrophages are required for normal phagocytic processing and that absence of these peptides results in abnormal and slower maturation and degradation of phagocytic materials. In turn, this abnormal phagocytic processing reduces the ability of the organism to fight off bacteria and has a direct impact on fly immunity and viability.

3—Hemotin and Vertebrate Stannin Belong to a Conserved Family

To elucidate the molecular function of the Hemotin peptide, we searched for similarity to known structural domains using the Phyre2 engine [35]. The human Stannin peptide is encoded by an 88aa smORF-like Hemotin and appeared as a possible match (Fig 4A). Structural analyses of the Stannin peptide suggest a transmembrane peptide containing two alpha helices, an N-terminal one that spans the lipid bilayer and a C-terminal helix at the cytosolic side (Fig 4A) [22,36], and our independent analysis of the Hemotin sequence using a transmembrane topology prediction program [37] also revealed a very similar potential transmembrane alpha-helical domain (Fig 4A, S1B Fig). Threading the Hemotin sequence onto the predicted human Stannin tertiary structure (see Materials and Methods) confirms a structural compatibility similar to vertebrate members of the Stannin family (Fig 4C).

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Fig 4. Conserved sequence and structure in Stannin, the Hemotin vertebrate homologue.

(A) Alignment showing Hemotin amino acid peptide sequences (see S4 File) from two insect lineages (Diptera and Hymenoptera) and vertebrate Stannin. The “Conservation” lane at the bottom reflects the conservation of physical–chemical properties in the amino acids, while the “Quality” lane scores the likelihood of observing the mutations displayed in each particular position of the alignment. Note that there exists high conservation in the transmembrane domain (blue) motif among these peptides. The N-terminal transmembrane and C-terminal alpha-helix domains are indicated below the alignment. Sequences studied in C) (see below) are highlighted with stars. (B) Cladistic guide tree showing the relationships between insect Hemotin and vertebrate Stannin peptides shown in (A). The Hemotin and Stannin sequences cluster with their respective counterparts from related species. Insect Hemotin sequences are branched into two main clusters, one represented by flies and the other by hymenopterans, with other dipterans such as mosquitoes being between these two. New Stannin homologues identified by us cluster with annotated Stannin sequences (prefixed by snn_) and show an overall correct phylogenetic position, with agnathans Lamprey and Hagfish basal to other vertebrates and closer to insect sequences (see also S4 Fig). Sequences studied in C) (see below) are highlighted with stars. (C) Threading of the peptide sequences (shown in red) from a basal vertebrate (hagfish), a basal hymenopteran (the wasp Microplitis), and D. melanogaster onto the human Stannin structure (shown in grey) (see Materials and Methods). It has been proposed that the N-terminal α-helix of the Stannin peptide transverses the membrane at an 80^o angle, whereas the C-terminal α-helix lies upon the lipid bilayer at the cytoplasmic side [36]. Note that the three peptides can adopt similar tertiary structures. GDT indicates the value of the High-Accuracy Global Distance Test, which measures the average distance (in Angstroms) between the Human Stannin model and the model for each peptide [38]. As expected, the Hagfish peptide, more closely related to human Stannin, obtains a lower distance, but insect peptides display values in the same range despite their lower sequence similarity (see A) above and S4C Fig). Supplemental data are shown in S4 Fig, S3 File and S4 File.

https://doi.org/10.1371/journal.pbio.1002395.g004

We further searched for homologues of Hemotin and of Stannin following a bioinformatics pipeline used in Magny et al., 2013 [7]. We identified homologues of Hemotin peptides in other dipterans (mosquitoes) and in other insects such as hymenopterans (bees, ants, and wasps), plus new homologues of Stannin in ancestral vertebrates, such as hagfish, lamprey, and cartilaginous fishes (Fig 4A and 4B and S4A and S4B Fig), and these sequences also show structural compatibility with Stannin (Fig 4C and S4C Fig). Despite considerable amino acid sequence variation, comparison of Hemotin and Stannin sequences reveals conservation in the alpha-helices and in the intervening sequence, which have been implicated in Stannin function [22] (Fig 4A). The resulting tree of Stannin and Hemotin sequences shows a good correlation with the animal phylogeny (Fig 4B) and correctly locates the 88aa mitochondrial ribosomal protein S21 as an outgroup (S4B Fig). Altogether, the amino acid sequence analyses suggest that that Stannin and Hemotin are members of the same peptide family, displaying sequence and structural homology.

4—Hemotin and Stannin Are Functional Homologues That Interact with 14-3-3ζ

Stannin is a peptide involved in organometallic toxicity, but its endogenous physiological and cellular functions have remained elusive [22]. In rodent models, snn expression has been detected in hematopoietic organs and immune cells including macrophages [39–41]. Similarly as with Hemotin (see above; unp. obs.), Flag-tagged Snn peptides have been detected in membrane fractions of intracellular compartments, such as endoplasmic reticulum, peroxisomes, mitochondria, and endosomes in mouse NIH-3T3 cell lines [42].

To explore the endogenous cellular function of hemo’s vertebrate relative, snn, in a vertebrate innate immune cell context, we used mouse RAW264.7 (macrophage-like) cells [43]. Firstly, we confirmed expression of snn in this cell line (S5A and S5B Fig). Secondly, we knocked down the expression of snn in RAW264.7 cells using siRNAs (small interfering RNAs) (S5A and S5B Fig) and monitored the formation of acidic compartments with Lysotracker. snn si-RNA-treated RAW 264.7 cells showed abnormally large acidic aggregates (Fig 5A–5C) mimicking the hemo^A4 null phenotype observed in hemocytes (Fig 2A” and 2L” and S1G–S1K Fig). Expressing a GFP-tagged human Stannin (Snn-GFP) peptide in Drosophila Kc167 cells revealed its localisation to cellular membranes, comprising intracellular punctate compartments (S5D–S5D” Fig). In addition, coexpression of tagged Stannin and Hemotin peptides in these cells show their colocalisation in membrane intracellular organelles (Fig 5D–5D”). These results suggest that Stannin and Hemotin peptides localize to similar cellular compartments and are involved in the formation of acidic compartments, suggesting a functional homology. To test this possibility, we measured the ability of Stannin to rescue hemo^A4 mutant phenotypes in hemocytes: expression of the human Stannin peptide rescued the vacuolated phenotype (Fig 5G) and the size of the FYVE-positive vesicles (Fig 5H, 5I, 5J and 5L; S6 Fig) of hemo^A4 mutants almost as effectively as endogenous Hemotin peptides (Fig 5G, 5H, 5I, 5J and 5K; S6 Fig), supporting that hemo and snn are functional homologues.

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Fig 5. Stannin and Hemotin replicate each other’s functions and counteract 14-3-3ζ function during endosomal maturation.

(A) Mouse macrophage-like RAW264.7 cells treated with control-scrambled siRNAs and labelled with the acidic pH-sensitive Lysotracker (Red). Scale bar (5 μm). (B) RAW264.7 cells treated with two Fluorescein isothiocyanate (FITC)-labelled snn siRNAs (green) (see S5 Fig) and stained with Lysotracker. Note the highly enlarged lysosomal compartment in snn siRNA-treated cells (see C). Scale bar (5 μm). (C) Lysosomal OAI, revealed by Lysotracker, in nontransfected or control siRNA and snn siRNA-treated RAW264.7 cells, showing that snn siRNA-treated RAW 264.7 cells have significantly larger lysosomal compartments than control samples (see S1 Data). The graph shows averages of three independent experiments. One-way ANOVA test showed that samples were significantly different [F(2,780) = 185.6, p < 0.0001]. Post hoc Bonferroni multicomparison test showed that the siRNA-snn sample was significantly different to nontransfected and siRNA-control samples (n ≥ 240, p < 0.05). On average, snn siRNA-treated cells show a reduction in snn expression of 60% relative to nontreated cells, whereas cells treated with control siRNA only show a reduction of 0.8% (see S5A Fig). Error bars represent SEM. (D–D”) Hemo-GFP and Snn-FLAG peptides expressed in hemocyte-like Drosophila Kc165 cells (20) using the Act5-Gal4 driver. Hemo-GFP (green) (D’) and Snn-FLAG (red) (D”) peptides colocalize in intracellular vesicles and punctate organelles (arrows). Scale bar (5 μm). (E) Colocalisation of Hemo-GFP peptides and Nt-tagged HA-14-3-3ζ protein in intracellular compartments in ex-vivo hemocytes. (E’) Hemo-GFP. (E”) HA-14-3-3ζ. (E”‘) Merged image. UAS constructs were driven with He-Gal4. Scale bar (10 μm). (F) Pull down of myc-14-3-3ζ with Hemo-GFP in transfected Drosophila Kc167 cells. Myc-14-3-3ζ interacts with Hemo-GFP but not with a GFP-only control. Molecular weight is indicated in kilodaltons Retention of Hemo-GFP and GFP is shown in S5C Fig. (G) Vacuole OVI measurements (see Materials and Methods and S1 Data) in primary hemocytes. Expression of human snn (UAS-snn) rescues the hemo^A4 vacuolation phenotype to a similar extent as the rescue observed by the UAS-hemoFL and hemo-ORF constructs. Reducing the dosage of 14-3-3ζ (in a heterozygous null 14-3-3ζ ^12BL/+ background, labelled 14-3-3ζ −/+) reduces the hemo^A4 vacuolation phenotype. Conversely, overexpression of 14-3-3ζ (UAS-14-3-3ζ) in hemocytes induces the formation of larger vacuoles. The induction of vacuoles by excessive 14-3-3ζ is reversed by simultaneous overexpression of hemo full-length transcript (UAS-hemoFL) or overexpression of the human Stannin peptide (UAS-snn) but not by the expression of the control UAS-GFP construct. One-way ANOVA test showed that the means were significantly different [F(9,365) = 14.26, p < 0.0001]. Post hoc multiple comparison Bonferroni’s test showed that hemo^A4, UAS-14-3-3ζ and UAS-14-3-3ζ;UAS-GFP samples were significantly different than wild-type, whereas the rest were not (n ≥ 20, p < 0.05). Error bars represent SEM. (H) Measurement of the occupied FYVE area index (OAI) in ex vivo prepupal hemocytes (see Materials and Methods and S1 Data). Overexpression of human Snn peptide (UAS-snn) rescues the hemo^A4-enlarged FYVE compartments. Similarly, reducing 14-3-3ζ function by expression 14-3-3ζ-RNAi restores the size of hemo^A4 mutant FYVE-organelles to wild-type. Conversely, overexpression of 14-3-3ζ (UAS-14-3-3ζ) mimics the hemo^A4 mutant FYVE phenotype. The overexpression 14-3-3ζ-phenotype is reversed by coexpression with hemo full-length transcript (UAS-hemoFL). UAS constructs were driven with He-Gal4. One-way ANOVA test showed that the means of samples were significantly different [F(8,346) = 23.15, p < 0.0001]. Multiple comparison post hoc Bonferroni’s test indicated that UAS-14-3-3ζ and hemo^A4 were significantly different than wild-type whereas the rest of the samples were not (n ≥ 20, p < 0.05). Error bars represent SEM. (I–N) Intracellular distribution of FYVE-positive (green) compartments in ex vivo prepupal hemocytes (see also H). Scale bar (5 μm). (I) In wild-type FYVE-positive organelles appear as small rings and punctae. (J) In hemo^A4 mutant hemocytes, FYVE compartments contain larger rings than wild-type (arrowheads). (K) Expression of Hemo-ORF peptide (UAS-hemo-ORF) rescues the enlarged hemo^A4 mutant FYVE compartments. (L) Expression of snn (UAS-snn) reduces the hemo^A4 FYVE phenotype. (M) Overexpression of 14-3-3ζ (UAS-14-3-3ζ) produces enlarged FYVE compartments (arrowheads). (N) Reducing 14-3-3ζ function with RNAi rescues the hemo^A4 mutant FYVE phenotype. UAS constructs were driven with He-Gal4. Yellow dashed lines indicate the cell body. Supplemental data are shown in S5 Fig, S6 Fig, and S1 Data.

https://doi.org/10.1371/journal.pbio.1002395.g005

Stannin peptides bind to the 14-3-3ζ adaptor protein, although the molecular and cellular implications of this binding have not been fully clarified [44]. 14-3-3ζ proteins form dimers that bind to phosphorylated amino acid residues of target proteins and modulate their functions [45]. We investigated whether a similar interaction exists between Hemotin and 14-3-3ζ during endosomal maturation in hemocytes. We observed that Hemo-GFP peptides (which localize to early endosomes, Fig 2G–2G” and S2H–S2H” Fig) colocalised with 14-3-3ζ in hemocytes (Fig 5E–5E”‘). In addition, Hemo-GFP peptides coimmunoprecipitated with Nt-tagged 14-3-3ζ proteins expressed in Drosophila Kc167 culture cells (Fig 5F and S5C Fig) and hemocytes (S1 File). The coimmunoprecipitation appears moderate but reproducible and specific, perhaps reflecting a moderate affinity, or a limited amount of the multifunctional 14-3-3ζ protein [45] being engaged by Hemotin peptides.

Our genetic analysis reveals a negative functional relationship between 14-3-3ζ and hemo/snn during endosomal maturation in hemocytes. Thus, excess of 14-3-3ζ function (in He-Gal4;UAS-14-3-3ζ hemocytes) resembled loss of hemo function (as in hemo^A4) by producing large vacuoles (Fig 5G) and FYVE vesicles (Fig 5H, 5I, 5J and 5M), thus suggesting that 14-3-3ζ and hemo work in opposite directions. An antagonistic, yet closely related, function is further suggested by gene dosage interactions. First, the 14-3-3ζ gain of function phenotypes were corrected by simultaneous gain of either hemo or snn: both UAS-hemo and UAS-snn corrected the extra vacuolation of UAS-14-3-3ζ (Fig 5G), while UAS-hemo corrected the large FYVE vesicles produced by UAS-14-3-3ζ. Second, reducing 14-3-3ζ function rescued the hemo^A4 phenotype to near wild-type, as indicated by vacuolation (after removing a copy of 14-3-3ζ in a hemo^A4 homozygous background, Fig 5G) and the size of FYVE vesicles (by expressing 14-3-3ζ RNAi in a hemo^A4 homozygous background, Fig 5J and 5N). These negative dosage interactions suggest that 14-3-3ζ works in a common pathway, yet antagonistically, with hemo and snn [46,47]. Because 14-3-3ζ dosage is able to modify hemo^A4 null phenotypes, the formal interpretation is that 14-3-3ζ acts downstream of hemo, or in other words, that Hemotin regulates 14-3-3ζ. Altogether, these genetic results suggest that Hemotin function represses or down-regulates 14-3-3ζ activity.

Thus, our results suggest that Hemotin and Stannin are functional homologues that are required at the cellular level for endosomal maturation; and at the molecular level, to bind and antagonize 14-3-3ζ.

5—Hemotin Represses the 14-3-3ζ-Mediated Stimulation of the Class II PI3K68D Kinase

Specific steps in the endosomal maturation process depend on the phosphorylation states of PI [18,20]. In hemocytes, regulation of PI(3)P endocytic pools is key for early endosomal trafficking [21,48]. It has been shown that PI(3)P homeostasis depends on the class II PI(3)Kinase, PI3K68D, which phosphorylates PI at the carbon 3 position to form PI(3)P and the Mtm phosphatase that dephosphorylates this residue to revert to PI [21]. Either mutations in mtm or overexpression of PI3K68D increase PI(3)P in early endosomes and induce the formation of abnormal enlarged endolysosomal compartments retaining larger amounts of the PI(3)P sensor FYVE-GFP (Fig 6E, 6F and 6J) [21], similar to that observed in hemo^A4 mutants (Fig 2B–2B”, S2C–S2C” and S2E–S2E” Fig).

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Fig 6. Hemotin modulates PI(3)P formation by repressing 14-3-3ζ-mediated activation of PI(3)K68D kinase.

(A–I) Distribution of FYVE-positive compartments in ex vivo hemocytes (see also J). Yellow dashed lines indicate cell body area, and arrowheads indicate enlarged FYVE compartments. Scale bar (5 μm). (A) Wild-type hemocytes. (B) hemo^A4 mutants show enlarged FYVE compartments (arrowheads), (C) this phenotype is rescued by mtm gain of function (UAS-mtm). (D) Hemocytes overexpressing 14-3-3ζ (UAS-14-3-3ζ) or (E) the PI3K68D Kinase (UAS-PI3K68D) also show enlarged FYVE compartments (arrowheads), similar to those observed in hemo^A4 mutants. (F) Enlarged FYVE compartments are also observed by the reduction of mtm (mtm-RNAi) in hemocytes. (G) Reduction of PI3K68D kinase function by RNAi rescues the enlarged-FYVE phenotype produced by overexpression of 14-3-3ζ. (H) Reducing the function of 14-3-3ζ (14-3-3ζ-RNAi) rescues the enlarged FYVE compartment produced by overexpression of PI3k68D or (I) by reduction of mtm. (J) Quantification of the FYVE OAI in primary hemocytes (see Fig 6, S1 Data). Knocking down the mtm function (mtm-RNAi) produces enlarged FYVE compartments as shown with hemo^A4 mutants. Both overexpression of mtm (UAS-mtm) and haploinsufficiency of PI3K68D (Df(3)PI3K68D/+), rescue the hemo^A4 mutant FYVE phenotype. Overexpression of PI3K68D (UAS-PI3K68D) mimics the hemo^A4 mutant FYVE phenotype, and this is rescued by overexpression of hemo full-length transcript (UAS-hemoFL). Reduction of 14-3-3ζ function (14-3-3ζ-RNAi) corrects the mtm loss of function and PI3K68D gain of function FYVE phenotypes. The enlarged FYVE compartments produced by over-expression of 14-3-3ζ (UAS-14-3-3ζ) are corrected by knocking down PI3K68D function (PI3K68D-RNAi). One-way ANOVA test indicated that means of samples are significantly different [F(12,508) = 14.01, p < 0.0001]. Post hoc Bonferroni’s multiple comparison test showed that hemo^A4, UAS-14-3-3ζ, UAS-mtm-RNAi, UAS-PI3K68D were significantly different to wild-type, whereas the rest of genotypes were not (n ≥ 19, p < 0.005). Error bars represent SEM. (K) In hemocytes, Hemo-GFP peptides colocalize with PI3K68D Kinase in intracellular compartments (arrowheads), presumably early endosomes. (K) PI3K68D-Cherry expression. (K’) Hemo-GFP, (K”) merge image. Scale bar (10 μm). (L) Western Blot of a Pull down experiment from hemocytes expressing PI3K68D-GFP and HA-14-3-3ζ revealing a protein interaction between PI3K68D and 14-3-3ζ. Supplemental data are shown in S6 Fig and S1 Data.

https://doi.org/10.1371/journal.pbio.1002395.g006

The similarities of the cellular phenotypes between hemo^A4 mutants and overproduction of PI(3)P by PI(3)P enzymes prompted us to carry out a genetic analysis of their functional interactions during endosomal maturation in hemocytes. The PI3K68D gain of function phenotype of enlarged FYVE organelles and vesicles was suppressed by coexpression of either a hemo full-length transcript (Fig 6J and S6 Fig) or a 14-3-3ζ-RNAi construct (Fig 6H and 6J and S6 Fig). Similarly, reduction of mtm function by expressing an mtm-RNAi construct produced aberrant enlarged FYVE compartments with larger-sized vesicles (Fig 6F and 6J; S6 Fig) [21]. Codepletion of mtm and 14-3-3ζ with RNAi constructs corrected the size of these abnormal FYVE compartments and vesicles (Fig 6I and 6J and S6 Fig). Formally, these results indicate that 14-3-3ζ cooperates with PI3K68D, whereas Hemotin and Mtm antagonize their action.

Three further results suggest that mtm and PI3K68D act downstream of Hemotin and 14-3-3ζ. Either overexpression of mtm or removing a copy of the PI3K68D corrected the abnormally large FYVE vesicles of hemo^A4 null hemocytes (Fig 6A–6C and 6J and S6 Fig). Similarly, reducing PI3K68D function with a RNAi construct produces little or no phenotype on its own (Fig 6J) [21] but is also able to rescue the enlarged size of FYVE vesicles produced by overexpression of 14-3-3ζ (Fig 6G and S6 Fig). Thus, in three different genetic conditions, a reduction of PI(3)P synthesis was able to suppress the enlarged and abnormal early-endosome-like vesicles produced by total loss of hemo or gain of function of 14-3-3ζ. These epistatic results strongly suggest that mtm and PI3K68D act downstream of Hemotin and 14-3-3ζ during endosomal maturation; in other words, that Hemotin and 14-3-3ζ fulfil their roles through regulation of the PI(3)P enzymes.

Two independent lines of evidence corroborate this hypothesis. First, Hemo-GFP peptides colocalize with PI3K68D kinase at early endosomes (Fig 6K–6K”‘). Second, PI3K68D is able to pull down 14-3-3ζ from hemocyte protein extracts (Fig 6L), suggesting that 14-3-3ζ directly binds PI3K68D. Altogether, the genetic, cellular, and biochemical results support a model where Hemotin peptides indirectly affect the PI(3)P labelling of early endosomes by binding 14-3-3ζ, and hence repressing the positive effect of 14-3-3ζ on PI3K68D kinase (Fig 7).

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Fig 7. Model for the role of Hemotin in phagocytic processing.

Simplified models of endosomal maturation, modified from [18] depicting the role of the proteins and markers analysed in this work. A) Wild-type endosomal trafficking is regulated by different phosphorylation states of PI. Phosphorylation of PI into PI(3)P is achieved in early endosomes by the class II or class III PI3 kinases PI3k68D and Vps34, respectively. In late endosomes PI(3)P is again phosphorylated to produce PI(3,5)P2. This phosphorylation step allows late endosomes to progress into degradation allowing lysosomes to fuse to late endosomes to produce multivesicular bodies. This trafficking progression can be reversed by dephosphorylation of PI(3)P or PI(3,5)P2 by myotubulurin phosphatases. Hemotin and Stannin are functional homologues that localise to early endosomes, where they bind and repress 14-3-3ζ. Our genetic and biochemical data indicates that 14-3-3ζ binds the PI3K68D kinase and promotes its function, perhaps by directly increasing its enzymatic activity, or indirectly by promoting its correct localisation in early endosomes. Since Hemotin antagonises 14-3-3ζ, it indirectly reduces the development of early endosomes through PI3K68D. B) The absence of Hemotin produces an excess of 14-3-3ζ function, which results in an excess of PI3K68D function and leads to an increase in endocytic vesicles containing PI(3)P, as detected by expansion of the area occupied by the FYVE marker. These abnormal vesicles display an abnormal maturation during phagocytosis, with excessive co-expression of early lysosomal markers (such as FYVE) and late ones (Lysotracker and Rab7), and a slower and less intense acidification of their contents, as revealed by the pHrodo pH marker.

https://doi.org/10.1371/journal.pbio.1002395.g007

Fly Genetics and Generation of Hemotin A4 Mutant Strain

Fly stocks and crosses were cultured at 25°C. The Oregon-Red line was used as our wild-type strain. The following lines were obtained from the Bloomington Stock Centre: fray^PZ07551, Df(3)BSC626 (PI3K68D), fru¹, 14-3-3ζ^BL12, UAS-Rab7-YFP, UAS-2XFYVE-GFP, UAS-LAMP1-GFP, UAS-PI3K68DIR(GL00159).

For ectopic and rescue experiments, we used Hemese-Gal4, and crq-Gal4 lines as described in Sampson et al., 2013 [15].

Fly strains used in this study: UAS-HA-14-3-3ζ and UAS-14-3-3ζIR [69], UAS-14-3-3ζIR (VDRC#48725). UAS-mtmIR, UAS-mtm-Cherry, UAS 2XFYVE-Cherry, UAS-PI3K68D-Cherry, and UAS-PI3K68D-GFP [21]. fray^R1 [25].

The P{RS3}fray (CB-0706-3) and the P-Bac{WH}fru(f02684) from DGRC and Exelixis, respectively, were used to generate a 34-Kb deficiency by FRT-mediated recombination [29] depleting the first 5’exon of fru and fray genes and the whole gene locus of CG7691 and CG43210 (hemo) genes, which was confirmed by Taq-polymerase PCR (Qiagen, Venio, Netherlands) from genomic DNA extractions. In addition, detection of mRNA levels of these genes were conducted by RT-PCR from mRNA extraction using Trizol (Ambion/Life Technologies, Carlsbad, CA) of hemoA4 larval haemolymph.

Primary Hemocyte Culture, Live Cell Stainings, Immunocytochemisty and Microscopy

Primary Drosophila hemocytes were isolated from postembryonic life stages as indicated in Sampson et al., 2013 [15]. Extraction of free-flowing hemocytes was achieved by bleeding individual specimens using a 25-gauge dissecting needle in a culture medium (80% Schneider’s Drosophila medium [Pan-Biotech, Dorset, UK] and 20% fetal bovine serum [Invitrogen/Life Technologies] with no antibiotics).

For measuring vacuolation primary hemocytes from white prepupae (>100 h AEL) were cultured into a glass-bottom dish containing collagen extracellular matrix (MatTek Corp., MA, US) and stained with anti-beta tubulin and phalloidin (Invitrogen/Life Technologies) following the procedures described in Sampson et al., 2013. Similar procedures were followed to identify lysosomes using lysotracker (Invitrogen/Life Technologies) or by detecting the LAMP1-GFP marker.

Detection of GFP and Cherry–2XFYVE, Hrs, Cherry-PI3K68D, Cherry-Mtm, HA-14-3-3ζ, and hemo-ORF1-GFP was achieved by culturing primary hemocytes from wandering late third instar larvae and white prepupa stages in glass multisport microscope slides (Hendley-Essex, Essex, UK) and immunostaining was performed as described in Sampson et al., 2013.

Antibodies used in this study are: mouse anti-GFP (Roche) 1:500; rabbit anti-cherry (Invitrogen) 1:5000; mouse anti-HA (Roche) 1:500; anti-Hrs [70], mouse anti-FLAG M2 (SIGMA) 1:1000. Actin cytoskeleton was labelled with fluorophore-conjugated phalloidin (Molecular-Probes. Invitrogen) 1:50.

For measuring lysosomal aggregation, live primary hemocytes were stained with 50 nM Red-lysotracker (Invitrogen) in culture medium for 15 min and then washed several times and finally replaced with normal culture medium for live imaging for 20–30 min.

For embryonic hemocyte in vivo imaging, adult flies were allowed to lay eggs on apple juice agar plate. Embryos collected at stage 15, dechorionated and mounted on double-sided tape stuck to a standard glass slide. Voltalef oil was applied to the embryos to prevent dehydration, and a coverslip fixed with nail polish applied on top. For pupal hemocyte in vivo imaging, pupae at stage P6 (50 h APF) were mounted ventral side down on double-sided tape applied to a glass slide and dissected to remove a window in the pupal case over the thorax as described in Sampson et al., 2013. After application of 10S Voltalef Oil on the ROI, a ring of petroleum jelly was made around the samples, a coverslip was rested on this ring, and then it was pressed down on the sample.

Confocal laser microscopy was used to image fluorescence in live and fixed cells. An inverted Zeiss Axiovert 200M series microscope with a LSM-510 confocal laser attachment was used. Images were captured using a Hamamatsu ORCA-ER C4742-95 camera. Most cells were observed at 63x using the Zeiss Apochromat 63x oil objective NA = 1.4, but in some cases 100x magnification was required, using a Zeiss Epiplan—NeoFluar 100x oil objective with NA = 1.3. Fluorescent imaging utilised the automated prior stage for Z-stacking throughout the cells imaged. Live cells were imaged at 0.5–1 μm path length between each imaging slice. Fixed cells were also imaged within the same range of path length for nondetailed images, whilst detailed images, particularly for 3-D reconstruction, were imaged at a path length of 0.2–03 μm between each imaging slice. LSM image browser and imageJ v.1.46 were used to analyse acquired images and Photoshop (Adobe) was used for editing.

Bacterial Phagocytosis Assay in Primary Hemocytes

Phagocytosis assays were performed on wild-type and hemo^A4 mutant live primary hemocytes expressing FYVE-GFP endocytic marker supported in ex vivo cell culture without ECM coating and using HEPES+HBSS pH 7.4. The phagocytic stimulation used was in the form of pHrodo pH-sensitive conjugated heat-killed bacterial particles. E. coli (Gram -) bio-particles were used (Life Technologies).

A 1:1 dilution of pHrodo-conjugated bacterial particles (0.5 mg/mL) with HEPES+HBSS pH 7.4 buffer was sonicated for 5 min to break apart large conglomerates. Further dilution was carried out by using 16 μl (1:1 dilution) into a final volume of 200 μl HEPES+HBSS pH 7.4 which was added to the ex vivo hemocyte culture. Time-lapse Imaging was conducted using a SP8 Leica confocal microscope with a short Z-stack throughout the cell at 0.5 μm path length between each image slice. Bright Field Phase, 488 and 543 nm laser illuminations were used to observe the cell outline and the fluorescent FYVE and pHrodo particles. Live cells were imaged every 90 to 120 seconds with a Z-stack for a period lasting for 90 min.

The integrated pHrodo intensity over time was measured for each particle from the moment the particles docked into the cell membrane and for each time point, using an identical ROI of 0.9 μm² for every series on single confocal z-stack plane across on the middle of the pHrodo particle. To quantify FYVE prevalence, the number of frames for which individual pHrodo particles were surrounded by FYVE, using a single z-stack plane across on the middle of the pHrodo particle, were counted and converted into time according to the frame rate of each time lapse.

Viability Assays

For viability assays, virgin male flies were collected from each genotype and kept at 18°C until sufficient numbers were reached (maximum 2 d, at least 50 flies per genotype or condition). Flies (2-d-old) were then kept at a density of 10–25 flies/tube, reared at 25°C and scored daily for number of surviving flies.

To assess the viability of flies in antibiotic media, 100 μL of Penicillin-Streptomycin solution (10,000 units penicillin and 10 mg streptomycin/mL, Sigma-Aldrich) was added to the surface of standard corn-meal food media vials (containing approximately 10ml of media), and left over night at 25°C, until fully absorbed.

To assess the effect of bacterial infection on viability, nonpathogenic Escherichia coli (DH5α), or pathogenic Micrococcus luteus (gram+) and Enterobacter cloacae (gram−) bacterial strains were used. Flies were infected by wounding at the top of the abdominal segment, under the haltere, using a tungsten dissection needle dipped in bacterial culture grown to a cell density of (OD600 = 0.8).

In Vivo mCherry E. coli Uptake Assay

To quantify the number of mCherry E. coli bacteria (K12 E. coli expressing PDSpRSETD-cherry plasmid from Dr. Stephan Mesnage) in hemocytes in vivo, He-Gal4; UAS-FYVE-GFP or he-Gal4; UAS-FYVE-GFP, hemo^A4 flies were injected with 0.2 μL of bacterial culture grown to OD600 = 0.05, using a glass capillary microinjection needle. Flies were then dissected as described in Magny et. al 2013 [7] to expose the dorsal abdominal cuticle in order to image the hemocytes associated to the dorsal vessel, as described in Horn et al. 2014 [71]. Flies were dissected after 20 or 120 min postinjection, in PBS, and the cuticles fixed in 4% PFA in PBS for 20 min. The preparations were then washed in, PBTX, incubated 30 min in PBS with phalloidin-Cy5 (Sigma-Aldrich) (1:10), washed in PBS, and mounted in Vectashield (vector). Hemocytes were imaged with a Zeiss laser scanning microscope LSM 5.10 on a Zeiss Axioskop 2 stage, with a 40X Achroplan objective, and bacterial cells within hemocytes, as determined with the phalloidin and FYVE counter-stains, were counted in Z-stack reconstructions.

Endogenous Bacterial Content Assay

To quantify endogenous bacterial contents, we followed the method described in Khalil et al. 2015 [72]. Briefly, hemo^A4 and wild-type Oregon Red L3 larvae were collected, washed in PBTX and PBS, and reared together in the same vial at a density of 20 flies per vial, in order to minimize the effect of external bacteria present in the media. Wild type and mutant flies were collected at either 10 d or 30 d after eclosion, and homogenised individually in 100 μL of autoclaved and sterile filtered PBS. The homogenates were diluted serially, and plated in antibiotic-free LB-agar plates. The plates were incubated for 48 h at 29°C, and the bacterial colonies from each homogenate spot counted.

Generation of DNA Constructs

The full-length hemo cDNA was obtained by RT-PCR (RACE) (PCR from overlapping EST (Flybase) using specific primers and cloned in TOPO vector (Invitrogen). pUAST-hemo full length vector was constructed by inserting flanking restriction sites to the 5’ and 3‘ end of the hemo cDNA respectively by PCR amplification. hemo cDNA was inserted in the pUASt (AttB) vector using these restriction sites. A similar strategy was used to clone ORF1 and ORF2 into the pUASt (AttB) vector. Generation of the pUASt-hemo-ORF carboxyl-tagged with GFP was carried out by generating a single PstI restriction site at the end of the ORF removing the stop codon in the hemo cDNA in TOPO by PCR and subsequently inserting a PstI flanking GFP cds in frame lacking its kozak and methionine sequences. Then the hemo-GFP was cloned in the pUASt-vector. The pUASt (AttB)-hemo-GFP was constructed for site-directed transgenesis by removing the hemo-GFP from pUASt and cloned in the pUASp(ATTB) vector (DGRC). The pUASt-ORF2 carboxyl terminal GFP tagged construct was made by amplification of ORF2 sequence and a short 5’UTR according to the short hemo cDNA described in flybase by PCR. The PCR product was cloned in pENTR vector (Invitrogen). LR recombination (Invitrogen) was used to clone the ORF2 into the pUASt-ctGFP vector (Murphy collection, Carnegie Institute).

The hemo frameshift construct was engineered by introducing a double nucleotide insertion in each ORF (in ORF1 a CG insertion three codons after the ATG and in ORF2 a GC insertion just after the ATG) by site directed mutagenesis (Stratagene, CA, USA) using the hemo full length cDNA in TOPO as a template. Subsequently, the hemo frameshift cDNA was cloned in pUASt (ATTB) vector.

A CG7691 Genomic Fragment spanning 11.8 Kb was constructed by amplifying independently four sequential genomic fragments by Long Expand Range PCR (Roche, Basel, Switzerland) each containing the EcoRI-Kpn1, KpnI-NotI, NotI-XbaI and XbaI-AscI restriction sites respectively and then cloned in TOPO vector (Invitrogen). Each fragment was sequentially cloned in the Casper 5 vector using the restriction sites. Proof of the presence of the wild-type the CG7691 gene was obtained by sequencing. CG7691 mRNA expression from the construct was detected in adult transgenic flies over an hemo^A4 deletion background by RT-PCR as described in [7].

To generate an amino terminal tagged version of 14-3-3ζ the ORF and 3’UTR from RH61958 cDNA (DGRC) were amplified by PCR and subsequently the PCR product was cloned in the pENTR vector (Invitrogen). The pUASt-Ntmyc-14-3-3ζ construct was generated by LR recombination into the acceptor vector (Murphy’s collection; Carnegie Institute).

A snn ORF clone in an entry vector (GeneCopoeia, MD, US) was inserted in the pUASp vector (Murphy’s collection, Carnegie Institute) by LR recombination (Invitrogen). Similarly, a pUASt- amino-terminal GFP- or FLAG-tagged SNN constructs were generated using the former snn ORF entry vector and destination vectors (Murphy’s collection, Carnegie Institute).

To generate the hemo-RNAi construct used the method described by Kondo et al. (2006) [73]. Briefly, we used the following primers (hemo RNAi Fw GTTCCACAGAGATATCGTCT hemo RNAi Rv ACCACGAAGCTAACGCACAGC) to amplify a 354 bp DNA fragment from genomic DNA, corresponding to a region of the hemo locus with no homology to other genomic regions. This fragment was then cloned into the pRISE vector and used to generate transgenic flies by Bestgene.

Drosophila S2 and Kc167 and Mouse RAW264.7 Cell Culture

S2 cell culture, transfections, and immunocytochemistry were performed as described in [7]. Kc167 cells (250X10⁵) were cultured in a 6-well plate dish in M3 insect medium (Sigma, MO, USA) containing 10% FBS and 1% pellicillin/streptomycin (Sigma). After 24 h, pUASt- and Act5-Gal4-constructs were transfected using FuGene (Roche). 3 d later, cells were pelleted for immunoprecipitation.

Mouse RAW264.7 cells were cultured in RPMi-1640 with L-glutamine (Sigma) supplemented with 10% FBS and 1% penicillin-streptomycin (Sigma) at 37°C and 5% CO₂ conditions. 300 x 10⁵ cells/well were grown in a 6-well plate (for lysotracker they were placed on acid-treated coverslips). siRNA treatments were performed as described by Ulvila et al., 2011 [43]. A mixture of two 5’ FITC labelled siRNAs were generated for snn, one targeting the ORF (sense sequence-GGCCAUGUGUGGAAAGAAAUU) and the other the 3’UTR(sense-sequence-GGGAGGAGCUGUAGGGAAGUU), and a control sample of nontargeting siRNAs pool manufactured by Dharmacon (GE Healthcare, NJ, US) were transfected using RNAmax reagent (Invitrogen) into cells at 24h and 48h after seeding the cells. After 24 h, cells were pelleted for Trizol (Ambion) mRNA extraction. Retrotranscription was conducted using random and polydT primers using RNA isolation kit (Promega, CA, US). snn and elongation translation factor 4 primers were used for RT-PCR. For quantification of lysosomes with Lysotracker, the media was removed and cells were washed with PBS1x. Afterwards, 50nM Lysotracker in culture media was added and left for 15 min. Cells were washed with culture media and mounted for acquisition of live cell imaging (30min) using confocal microscopy with a Zeiss LSM 510 NLO AXIOSkop microscope. Three independent siRNA treatment repeats were performed in our analysis.

Immunoprecititation and Western Blot

Late third instar larvae (80) and pelleted Kc167 cells were homogenized in lysis buffer (50 mM Tris pH 7.5;150 mM NaCl;1 mM EDTA;1 mM EGTA; 2.5 mM pyrophosphate; 1 mM Na₃VO₄; 1 mM glycerol phosphate) for 1 h (4°C). Cellular debris was spun at 3,000 rpm for 15 min at 4°C. Supernatant was again spun at 9,500 rpm for 45 min at 4°C. Supernatant was added to equilibrated GFP-beads (Chromo Tek, NY, US) and left rotating overnight at 4°C. Beads were washed several times and then boiled in Laemmi Loading Buffer (Biorad, CA, US). Beads were loaded onto 8% or 12% polyacrylamide gel and proteins were separated by SDS-PAGE (Biorad). Specific proteins were detected by Western Blot using a semidry blotting or tetra cell (Biorad). Antibodies used were: mouse anti-GFP 1:3000 (Roche); anti-HA 1:5000 (Roche); anti-Myc 1:1000 (Upstate).

Statistical Analyses of Cell-Based Assays

To measure the size of the vacuoles of in vivo embryonic and pupal thoracic hemocytes, we calculated the OAI, which is the accumulative area of intracellular vacuoles divided by the cellular area (>30 cells for most of the genotypes randomly selected as described above) in hemocytes in the wild-type and hemo^A4 mutant. Z-stack images were processed in the ImageJ software, and the outlines of the vacuole and cell were highlighted using the draw tool and the areas were measured. Student t test was used for statistical analysis.

For measuring vacuolation in ex vivo hemocytes, we determined the OVI, which corresponds to the addition of the volume of intracellular vacuoles (voids that disrupt the tubulin cytoskeleton)/total cytoplasmic volume of the cell. The Z-stack images were loaded onto the ImageJ software. To calculate the total cytoplasmic volume, we used the oblate ellipsoid volume formula: 4/3πab² (a = radius, b = height). The radius of the cytoplasm was measured at widest Z-slice of the cell and the height at the highest point at the orthogonal projection. The average wild-type cytoplasmic volume of ex vivo hemocytes (>30 cells for most of samples were selected at random across >3 separate imaging foci and 3 experimental repeats) was 311.06 μm³, which diverged only slightly from A4 mutants (375.57 μm³), therefore the wild-type cytoplasmic volume was used for our analysis. To calculate the volume of intracellular vacuoles, we used the sphere volume formula: = 4/3πr³, the radius was calculated by measuring the diameter of the widest point of the vacuole, only the vacuoles with a radius equal to or larger than 1.5 μm were considered for OVI analysis, with cells lacking vacuoles of at least 1.5 μm giving an OVI of 0.

To measure early endosome cellular compartments (FYVE-positive) in ex vivo hemocytes, we calculated the OAI, meaning the addition of FYVE particle area/total cytoplasmic cell area (volume inside of cortical actin ring). ImageJ software was used to calculate the OAI. A maximum projection of the Z-stacks was made and both FYVE and Phalloidin channels were separated. FYVE image was transformed into a binary image and particle analyses (size pixel μ²: 20-infinity and circularity: 0–1) were used to calculate the area of particles. Addition of FYVE particle areas was done in Excel. Measurement of the cytoplasmic area was performed manually by using the free hand selection tool. OAI of endosomes of 20–60 cells selected at random as described above were calculated per genotype. In addition, we measured FYVE diameter ratio consisting of the average of three FYVE vesicle diameters per cell (randomly selected 20–60 cells from >3 imaging foci and 3 experimental repeats per genotype) divided by the averaged FYVE diameter of the He-Gal4 UAS-hemo full-length transcript in an A4 mutant background to diminish the genetic background effects possibly caused by the Gal4, hemo^A4 and UAS-docking chromosomes.

Lysosome aggregates in ex vivo hemocytes were measured by calculating the area of lysosome particles as described above (visualized with lysotracker)/number of lysosomal particles. For each genotype >30 cells were selected randomly as described above and were measured.

We calculated the OAI as described above as a measure of the lysosome occupied area in the siRNAs experiments in mouse RAW264.7 cells. 80–110 cells were measured per sample in each of the three repeats.

Statistical analysis was performed with one-way ANOVA to assess whether the means of the groups were significantly different (p < 0.0001), and a post hoc Bonferroni’s test was carried out to compare multiple groups with p < 0.05 considered as significant using the Prism suite (Graph Pad).

As a measure of the colocalisation of endocytic markers and hemotin-GFP peptides, we used the Pearson’s correlation coefficient (ImageJ WCIF colocalisation plugins) that evaluates the amount of signal intensity from one channel that occurs in the same location in the other colour channel. Pearson’s coefficients range between −1 to 1, with values closer to 1 indicating reliable colocalisation. Pearson’s coefficients of hemocyte cells (n = 17–32 per genotype) were analysed with two-tailed Mann-Whitney test (p < 0.0001).

Search for Sequence and Structure Homologues Using Bioinformatics

The Phyre2 online search engine was used to find structural homologues using the Hemo-ORF peptide sequence as an input. To find Hemotin sequence homologues, we used the pipeline described in Magny et al., 2013 [7] that is based on the identification of closest homologues, followed by expanded searches using consensus sequences weighted by the phylogeny from the alignment of such close homologues. In addition, we introduced an extra corroboration step of reciprocal best Blast hit in doubtful cases.

Alignments and trees of peptide sequences were generated using MAFFT and Clustal programs, with MAFFT parameters set at 5 iterations and global pair alignment. The alignments were visualized with Jalview.

To evaluate the ability of hemo peptides to adopt the tertiary peptide structure of human Stannin, we threaded the sequence of insect Hemotin peptides (D. melanogaster and Microplitis demolitor) and the basal vertebrate Stannin peptide (hagfish) onto the human Stannin peptide structure (1zzA; RCSB protein data bank) using the Phyre2 server [35] on one-to-one threading mode. These preliminary structures were then refined, and Global distances test values were obtained with respect to the original human 1zzA Snn structure using the KobaMIN server [38].