To further analyze the function of Jumu in larval hematopoietic homeostasis, we first detected the expression of the plasmatocyte marker P1, the crystal cell marker Hindsight (Hnt) and the lamellocyte marker L1 in the lymph glands of jumu mutants. Because all jumu homozygous null mutants died during embryogenesis, to further reduce the expression of Jumu, we utilized the double heterozygotes generated by crosses between different jumu heterozygote mutants. Only a few jumu double heterozygotes mutants survived until adulthood, and these were found to exhibit poor viability and female sterility. The jumu heterozygote mutant lymph gland showed obvious reductions in plasmatocyte and crystal cell differentiation (Figure 1A–J). Moreover, the number of crystal cells in the jumu double heterozygotes mutants was clearly less than that found in heterozygotes (Figure 1I,J). However, plasmatocyte differentiation in the jumu double heterozygotes mutants was not clearly reduced compared with that found in w¹¹¹⁸ (Figure 1D,E). In addition, only very few lamellocytes could be detected in less than 10% of the lymph glands (n = 30 lobes) of jumu heterozygotes, whereas more than 90% of the lymph glands in the jumu double heterozygotes mutants (n = 40 lobes) showed large numbers of lamellocytes in the primary and secondary lobes (Figure 1K–N). Moreover, a reduced primary lobe size and an enormous overgrowth of secondary lobes were features of jumu double heterozygotes lymph glands (Figure 2E,L). To further determine whether the change in hematopoietic homeostasis observed in the jumu double heterozygotes mutants depends on the expression levels of jumu, the transcription levels of jumu in heterozygotes and double heterozygotes third-instar larvae were quantified. The jumu transcription levels were reduced by approximately half and 40-fold in the heterozygotes and the double heterozygotes mutants, respectively (Figure 1EE). In summary, these results indicate that a change in jumu expression substantially disturbs blood cell homeostasis and the growth and development of the lymph gland.

Figure 1

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Figure 2

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We subsequently analyzed the expression pattern of the MZ-specific markers domeless-Gal4,UAS-2×EGFP (dome>GFP), E-cadherin (Shg) and Patched (Ptc), which is a receptor in the Hh pathway. In contrast to wild-type larvae, the lymph glands of jumu heterozygotes third-instar larvae strongly expressed prohemocyte markers and presented a greatly enlarged MZ (Figure 1O–Y). However, the lymph glands of jumu double heterozygotes mutants did not show an obviously enlarged MZ compared with the wild-type (Figure 1X,Y). In addition, we also assessed ROS as a specific marker for MZ. ROS appear by the third instar, and increased levels of ROS can prime progenitor cells for differentiation (Owusu-Ansah and Banerjee, 2009). Consistent with the results of a previous study, ROS appeared in the MZ of the lymph gland of w¹¹¹⁸ mid-third-instar larvae and remained increased until late stages (Figure 1Z); however, ROS were rarely observed in jumu mutants during the entire third instar stage (Figure 1AA-DD). These results indicate that the loss of jumu prevents the differentiation of hematopoietic progenitors into plasmatocytes and crystal cells.

Previous studies have indicated that an increased PSC domain size can cause an increase in the MZ and a reduction of hemocyte differentiation (Mandal et al., 2007; Pennetier et al., 2012). To investigate whether an increase in the number of PSC cells could be observed in jumu mutants, we detected the expression of Antp, a specific marker of the PSC in the lymph gland. As predicted, the loss of jumu indeed led to substantial increases in the PSC size and cell number to different degrees (Figure 2A–E,U). In particular, the deletion mutants jumu^Df2.12 and jumu^Df3.4 showed strong increases in PSC cells and ectopic expansion of Antp; these ectopic PSC cells were scattered and often formed two or more cell clusters with higher accumulation in the MZ and CZ (Figure 2C,D). Moreover, the jumu double heterozygotes mutants showed more severe defects in PSC morphology, specifically the existence of PSC cells mostly in the secondary lobes (Figure 2E). In addition, the Hh signal from the PSC is required for the maintenance of prohemocytes (Mandal et al., 2007); thus, we subsequently detected Hh expression using the hhF4-GFP reporter gene. Similarly to the expression of Antp, jumu mutants also showed increased numbers of Hh⁺ cells, and these were highly co-localized with Antp⁺ PSC cells (Figure 2F–G’). It has been indicated that Col is also a PSC-specific marker and regulates Antp and Hh expression in the PSC in third-instar larvae (Mandal et al., 2007; Pennetier et al., 2012). Thus, we tested whether jumu affects the PSC by controlling the level of Col. Similar to Antp and Hh, increased Col was also observed in jumu mutant lymph glands, and increased numbers of Col>GFP⁺ PSC cells were colocalized with Antp⁺ cells (Figure 2H–L). Moreover, a reduced expression of Col can inhibit the phenotypes associated with jumu mutants: specifically clearly reduced numbers of both Col⁺ and Antp⁺ PSC cells accompanied by a reduction in the MZ size and an increased differentiation of plasmatocytes (Figure 2M–U).

To determine whether the ectopic PSC cells in jumu mutant lymph glands are migrated from the PSC or are de novo arising in the MZ and CZ, we first detected the morphology of the PSC at different larval stages. Expanded PSC cells were observed in jumu mutant L1 larvae and were amplified in L2 and L3 larval stages (Fgure 3A-L’). In addition, we observed that the ectopically expanded Antp⁺ PSC cells did not co-localize with any dome-GFP⁺ cells or Hml-GFP⁺ cells in the MZ and CZ domains, particularly among the ectopic PSC cells adjacent to MZ or CZ cells during the early larval stage; in fact, we did not detect any intermediate cells co-expressing both Antp and GFP appeared (Figure 3E–L’). In summary, these results indicate that the ectopic PSC cells in jumu mutants are derived from the PSC domain, and Jumu controls hematopoietic homeostasis by serving as a crucial negative regulator of Col⁺ PSC cell proliferation and ectopic expansion.

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To further investigate where and how Jumu regulates the number and organization of PSC cells, we first identified the expression pattern of Jumu protein in the lymph gland at different larval stages through anti-Jumu antibody staining. In wild-type larvae, Jumu is expressed in the entire lymph gland and mainly located in the nuclei and presents markedly higher expression in the PSC than in the other domains during the L1 to L3 larval stage (Figure 4A–F”); however, the signal was nearly abolished in jumu double heterozygotes (Figure 4E). Moreover, we found that Jumu is also expressed in cardiomyocytes and is colocalized with Antp in some cells (Figure 4A–A’’). A recent study suggested that Slit/Robo signaling from the cardiac tube is required to preserve the morphology and function of the PSC (Morin-Poulard et al., 2016). To determine whether the expansion of the PSC cells in jumu mutants is caused by the loss of jumu in cardiomyocytes, we used a heart-specific Gal4 driver (Hand-Gal4) to express jumu in jumu^Df2.12. However, the PSC morphology defects detected in jumu^Df2.12 were not rescued by the overexpression of jumu (Figure 4G,H and S). We then used tissue-specific Gal4 lines to achieve a forced expression of jumu in the jumu^Df2.12 PSC niche (Antp-Gal4), MZ (dome-Gal4, TepIV-Gal4) and CZ (Hml-Gal4) and found that the overexpression of jumu in the PSC or CZ domain cannot rescue the ectopic expansion of PSC cells in jumu^Df2.12 (Figure 4I,J,M,N and S). However, the overexpression of jumu using dome-Gal4 and TepIV-Gal4 effectively inhibited the generation of expanded PSC cells in the MZ and CZ (Figure 4K,L and S; Figure 5A,B and R). Moreover, the overexpression of jumu in the entire lymph gland using the pan-lymph gland specific driver Srp-Gal4 or the ubiquitous driver Tub-Gal4^ts also rescued the PSC morphology defects detected in jumu^Df2.12 (Figure 4O–S). Thus, these results indicate that Jumu protein in prohemocytes might prevent the ectopic expansion of PSC cells through a non-cell-autonomous mechanism.

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To further determine whether jumu expression in the MZ non-cell-autonomously prevents the expansion of PSC cells, we knocked down or overexpressed jumu using the MZ-specific drivers TepIV-Gal4 and dome-Gal4 and assessed the resulting PSC morphology. Similar to jumu mutants, the TepIV>jumu RNAi and dome>jumu RNAi mutants also showed scattered and increased numbers of PSC cells; however, the overexpression of jumu in MZ did not cause any defects in PSC morphology (Figure 5A–D,R; Figure 6A–C,E). We then used the pan-lymph gland specific driver Srp-Gal4 and e33C-Gal4 to knock down or overexpress jumu in the entire lymph gland and both Srp-Gal4>jumu RNAi and e33C>jumu RNAi displayed the scattered and increased numbers of PSC cells (Figure 5E–I,R). Because e33C>UAS-jumu was lethal during embryogenesis, we only detected the PSC of Srp>UAS-jumu and did not observe any changes in the morphology and cell number of the PSC domain (Figure 5G,R). These results strengthen the notion that Jumu protein located in the MZ mainly prevents the expansion and scattering of PSC cells toward the MZ and indirectly maintains the normal MZ development, but is not sufficient to initiate the inhibition of the proliferation of PSC.

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Slit/Robo signaling controls the clustering and number of PSC cells by regulating DE-cadherin (Shg) and Cdc42 expression and dMyc activity, respectively (Morin-Poulard et al., 2016). Notably, a deficiency in Slit/Robo signaling resulted in defects in the PSC similar to those obtained in jumu mutants, and both Slit, which is the canonical ligand of Drosophila Robo, and Jumu are expressed in the dorsal vessel (Morin-Poulard et al., 2016). Thus, we first asked whether Jumu protein in cardiomyocytes controls Slit/Robo signaling and, in turn, regulates both proliferation and clustering of the PSC. However, the knockdown or overexpression of jumu under the control of hand-Gal4 did not cause the abnormal PSC morphology (Figure 6—figure supplement 1A–D). This result suggests that Jumu does not directly regulate Slit/Robo signaling in the dorsal vessel. Moreover, the loss of Shg or the activation of Cdc42 in the PSC can induce the clustering of PSC cells (Morin-Poulard et al., 2016). Thus, we questioned whether Jumu protein in the MZ controls the PSC morphology by regulating the expression of Shg or the activation of Cdc42. Therefore, we first analyzed Shg expression in the PSC and found that the expression levels of Shg were not reduced in the PSC of jumu^Df2.12 compared with the wild-type (Figure 5—figure supplement 1A–B’’), suggesting that the role of Jumu in the PSC morphology is independent of Shg. Moreover, the overexpression of the dominant negative form of Cdc42 (cdc42-DN) in the PSC of jumu^Df2.12 did not rescue the increased number and clustering of the PSC cells (Figure 5—figure supplement 1C–E). Slit/Robo signaling controls PSC cell proliferation via dmyc, and promotes dmyc transcription through the negative regulation of the HSPG (heparin sulfate proteoglycan) Dally-like (Dlp) and BMP/Dpp signaling pathways (Morin-Poulard et al., 2016). We subsequently asked whether Jumu also controls the PSC cell number by regulating dMyc expression. The use of TepIV-Gal4 to knock down or overexpress jumu did not change the expression levels of dMyc in the MZ and CZ; however, we observed increased dMyc expression in the PSC cells of TepIV>jumu RNAi mutants (Figure 5J–L’, S). In addition, we detected dMyc in the lymph gland of e33C>jumu RNAi and jumu mutant larvae. The loss of jumu throughout the lymph gland did not cause an obvious reduction of dMyc in the MZ and CZ but caused an observable increase in the dMyc levels in PSC cells (Figure 5M–Q’, S). Moreover, the knockdown of dmyc in the PSC of jumu^Df2.12 effectively suppressed the scattering of and the increase in the number of PSC cells (Figure 5T–V). These results indicate that Jumu protein in prohemocytes might be required to prevent the overexpression of dMyc in the neighboring PSC cells in a non-cell-autonomous manner and that it accordingly inhibits the overproliferation and ectopic presence of PSC cells in an indirect manner. The loss of BMP/Dpp activity leads to increased numbers of PSC cells through the up-regulation of the expression of dMyc, and Dpp signaling is regulated directly by Dlp (Pennetier et al., 2012; Morin-Poulard et al., 2016). Thus, we subsequently detected whether Jumu controls dMyc expression by regulating Dlp and Dpp signaling. However, the normal expression of Dlp and phosphorylated Mad (P-Mad), which targets the activity of the BMP/Dpp signaling pathway, were detected in the PSC cells of jumu^Df2.12 (Figure 5—figure supplement 1F–I’’). Taken together, these results suggest that Jumu in MZ might control the morphology of the PSC by indirectly regulating dMyc expression in PSC cells through a Slit/Robo and BMP/Dpp-independent mechanism.

Because Jumu is expressed throughout the lymph gland, we investigated the role of jumu in the various domains of the lymph gland. We knocked down or over expressed jumu under the control of a driver specific to differentiating cells (Hml-Gal4), but this had no effect on the size of the PSC niche or on prohemocyte differentiation (Figure 6—figure supplement 1E–L). To determine whether Jumu is involved in prohemocyte differentiation in a cell-autonomous manner in addition to its non-cell-autonomous role in the regulation of PSC morphology, we used the dome-Gal4 driver to knock down or overexpress jumu in the MZ. Although an expansion of the neighboring PSC population was observed (Figure 6A,B and E), the knockdown of jumu in prohemocytes did not obviously reduce plasmatocyte or crystal cell differentiation (Figure 6F,G,J,K,L and O). However, the overexpression of jumu in the MZ resulted in massive differentiation of plasmatocytes and crystal cells and a reduction in hematopoietic progenitors (Figure 6H,J,M and O). Because dome-Gal4 is also expressed in the brain, and some signaling pathways derived from the brain also control the differentiation of prohemocytes (Shim et al., 2013), we examined the possible involvement of the brain by overexpressing or knocking down jumu using the brain-specific driver elav-Gal4, but this did not result in an abnormal differentiation of prohemocytes (Figure 6—figure supplement 1M–P). Because the forced expression of jumu in the MZ was not sufficient to reduce the number of PSC cells (Figure 6C,E), the overdifferentiation of prohemocytes could not be indirectly caused by the reduction in the number of neighboring PSC cells. Hh signaling from the PSC is required for the maintenance of hematopoietic progenitors (Mandal et al., 2007). Thus, we subsequently detected Hh expression in the PSC using the HhF4-GFP reporter gene. However, the overexpression of jumu in the MZ did not reduce the HhF4-GFP expression levels in the PSC, and the knockdown of jumu in the MZ also did not increase the HhF4-GFP expression levels in the PSC (Figure 6—figure supplement 2). Based on these results, jumu might control the differentiation of prohemocytes through other cell-autonomous mechanisms.

To further confirm whether Jumu cell-autonomously affects prohemocyte differentiation, we knocked down or overexpressed jumu in lymph gland-specific clonal populations of hemocytes using HHLT-Gal4 (Hand-Hml lineage-traced Gal4). A proportion of P1⁺ differentiated plasmatocytes were colocalized with the wild-type clones (GFP⁺) generated by HHLT-Gal4 (Figure 6—figure supplement 3A–A’’). However, most P1⁺ differentiated plasmatocytes did not overlap with clonal cells lacking jumu (GFP⁺) (Figure 6—figure supplement 3B–B’’), suggesting that the loss of jumu in clonal cells can inhibit hemocyte differentiation cell-autonomously. Conversely, most P1⁺ cells were colocalized with clonal cells overexpressing jumu (GFP⁺) (Figure 6—figure supplement 3C–C’’), suggesting that the overexpression of jumu in clonal cells can promote hemocyte differentiation in a cell-autonomous manner. Altogether, the HHLT clonal analyses indicate the cell-autonomous requirement of Jumu in controlling lymph gland hemocyte differentiation.

A recent study suggested that in addition to its expression in the PSC, Col is also expressed at low levels in prohemocytes and is required for preventing the differentiation of prohemocytes (Benmimoun et al., 2015). The above-described results show that the loss of col in the lymph gland of the jumu mutants also rescued the increased MZ size and the reduced differentiation of plasmatocyte phenotypes in addition to inhibiting the expansion of PSC cells (Figure 2M–T). Thus, we evaluated whether Jumu could cell-autonomously inhibit the expression of Col in prohemocytes in addition to preventing the ectopic expansion of Col⁺ PSC cells. The intensity of the Col signal in the MZ was quantified. As predicted, the expression of Col was clearly increased in the dome>jumu RNAi lymph gland and significantly reduced in the dome>UAS-jumu lymph gland (Figure 6P–R’, V). Similarly, jumu mutants also displayed an increased Col expression signal in the MZ (Figure 6S–V). A previous study suggested that Col regulates the expression of (Dlp) in the PSC (Pennetier et al., 2012). We found that Dlp is also expressed in the MZ (Figure 6W,W’), and the overexpression of col in the MZ enhanced the production of Dlp protein, an effect that was accompanied by the expansion of the MZ (Figure 6X,X ’and AA). Consistent with this phenotype, we found increased and reduced levels of Dlp expression in the MZ of dome>jumu RNAi and dome>UAS-jumu mutants, respectively (Figure 6Y–AA). In addition, the dome-driven expression of col was sufficient to suppress the plasmatocyte and crystal cell differentiation phenotypes associated with jumu overexpression in prohemocytes (Figure 6I,J,N and O). These results demonstrate that Jumu cell-autonomously promotes the differentiation of prohemocytes by inhibiting the expression of Col in the MZ. Moreover, the overexpression of Col in prohemocytes did not induce the ectopic expansion of PSC cells (Figure 6D,E); thus, the expanded PSC cell phenotype caused by jumu loss-of-function is independent of the increased expression of Col in the MZ.

To test whether jumu is also cell-autonomously required in the PSC, we used a col-Gal4 driver to knock down and overexpress jumu in the PSC. The expression of dome-MESO, a reporter of JAK-STAT signaling, was used to identify prohemocytes. In contrast to knockdown in the MZ or the entire lymph gland, the knockdown of jumu in the PSC (col>jumu RNAi) led to a reduction in the number of PSC cells and the overdifferentiation of plasmatocytes at the expense of the MZ; consistently, the overexpression of jumu in the PSC (col>UAS-jumu, Antp>UAS-jumu) substantially increased the number of PSC cells and resulted in an expansion of the MZ at the expense of hemocyte differentiation (Figure 7A–L; Figure 7—figure supplement 1A–C). Moreover, we also observed the similar phenotypes by clonally knocking down or overexpressing jumu in the PSC using HHLT-Gal4 (Figure 6—figure supplement 3D–F). These results suggest that Jumu cell-autonomously controlsthe size and cell number of the PSC. A previous study suggested that a reduction in the Col levels in the PSC can cause an increase in the number of PSC cells by down-regulating Dlp and Hh expression (Pennetier et al., 2012). The above-described results indicate that jumu negatively regulates the expression of Col in prohemocytes. Thus, we first asked whether the overexpression of jumu also reduces the expression of Col in the PSC, thereby indirectly increasing the number of PSC cells. However, although the number of Col⁺ PSC cells was significantly increased, the average pixel intensity of the Col signal per PSC cell was not reduced in the Antp>UAS-jumu mutant (Figure 7—figure supplement 1D–F). In addition, the overexpression of jumu in the PSC did not result in a loss of Hh and Dlp expression, which are directly regulated by Col (Figure 7—figure supplement 1G–K’). Dpp signaling is regulated directly by Dlp and indirectly by Col (Pennetier et al., 2012). We subsequently detected whether Jumu protein in the PSC controls the proliferation of PSC cells by suppressing the activity of the Dpp signaling pathway. The normal expression level of P-Mad could be detected in the Antp>UAS-jumu mutant (Figure 7—figure supplement 1L–M”). These results indicate that jumu cell-autonomously controls the size of the PSC niche through a Col-independent and Dpp-independent mechanism.

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Recent studies have suggested that some genes and signaling pathways control the PSC cell number and size by regulating dMyc expression in the PSC (Pennetier et al., 2012; Sinenko et al., 2009; Benmimoun et al., 2012; Tokusumi et al., 2015). Moreover, the above results suggest that Jumu protein in the MZ non-cell-autonomously prevents an increase in dMyc expression in the PSC. In addition, the knockdown of dmyc or jumu in the PSC resulted in a similar phenotype of fewer PSC cells, and correspondingly, the overexpression of dmyc or jumu in the PSC also caused increased numbers of PSC cells. Thus, we hypothesized that the existence of a regulatory relationship between Jumu and dMyc. Therefore, we first detected the expression of dMyc in jumu loss- and gain-of-function lymph glands. Surprisingly, a reduced level of dMyc was observed in the PSC of col>jumu RNAi lymph glands, whereas an increased level of dMyc was observed in col>UAS-jumu lymph glands (Figure 7M–O’, R). However, the knockdown of dmyc in the PSC did not result in an obvious absence of Jumu expression in the PSC (Figure 7P–Q”, S). Additionally, the knockdown of dmyc in PSC cells can suppress the increase in PSC cells caused by the overexpression of jumu (Figure 7T–V). These results indicate that Jumu is a direct or indirect positive regulator of dMyc expression in the PSC and promotes the overproliferation of PSC cells.

The above results show that Jumu in the MZ does not cell-autonomously regulate dMyc expression in the MZ (Figure 5J–L’). Similarly, the loss or overexpression of jumu in the CZ did not cause obvious changes in the dMyc levels (data not shown). These results suggest that the role of Jumu in the regulation of dMyc expression is specifically restricted to the PSC of the lymph gland. To further address whether Jumu can directly regulate dmyc transcription, we subsequently detected the protein and transcription levels of the dmyc gene in circulating hemocytes and found that the overexpression of jumu in circulating hemocytes led to increases in the dMyc protein level and the transcription of the dmyc gene (Figure 8A,B). We then analyzed the 2000 bp promoter sequence of the dmyc gene using Ensembl, a web system for predicting promoters, and found 23 putative FKH transcription factor- binding sites on the promoter sequence using JASPAR, a database of transcription factor binding profiles (Figure 8C,D). Because Jumu contains a conserved FKH DNA-binding domain, we then examined whether Jumu regulates the expression of dmyc at the transcriptional level by directly binding to its promoter via the FKH domain in vivo through a chromatin immunoprecipitation (ChIP) assay. We designed specific primers for possible FKH domain-binding sites in the dmyc promoter (Figure 8D). Total lysates from S2 cells stably expressing Flag-Jumu were used as an input positive control, along with immunoprecipitation with IgG (as a negative control) or antibodies against Flag, and the immunoprecipitated chromatin fragments were detected using PCR. As shown in Figure 8E, using anti-Flag antibodies compared with IgG control, there was no obvious positive signal at the dmyc promoter region of −1993 bp to −105 bp, but a specific enrichment of the dmyc gene region of −46 bp to +235 bp was observed. This fragment contains three putative FKH-binding sites, one inside the promoter region and two inside the 5’UTR (Figure 8F). To further determine the binding site recognized by Jumu in the dmyc promoter, we mutated each of the three putative FKH-binding sites at two positions crucial for the efficiency of FKH binding (Figure 8G). The Fragment from −643 to +467 of the dmyc promoter was cloned into the luciferase reporter vector pGL3, and a reporter assay showed that the co-transfection of S2 cells with a jumu expression construct and pGL3-dmyc (WT-643/+467) resulted in a nearly two-fold induction of reporter gene activity compared with the control. The disruption of the Site 1 (Mut1) led to an almost three-fold reduction of reporter gene activity compared with WT-643/+467, whereas the mutation of the Site 2 (Mut2) and Site 3 (Mut3) did not affect the reporter gene activity. Taken together, these results suggest that Jumu can be endogenously recruited to the conserved FKH-binding sites located in −27/–17 of the dmyc promoter via its FKH domain, and further strengthen the possibility that Jumu positively up-regulates dmyc expression by directly binding to the dmyc promoter.

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The above results show that the knockdown of jumu in the MZ and PSC has oppositing effects on the expression of dMyc and PSC morphology. We subsequently analyzed the spatial-temporal regulation of Jumu on PSC development during larval development. First, the expression pattern of dMyc during larval development was detected. dMyc is expressed at high levels in MZ and CZ cells but at low levels in PSC cells, and its expression pattern remains the same at all stages of larval development (Figure 7—figure supplement 2). Similarly, the expression pattern of Jumu does not change during larval development (Figure 4A–F’’), suggesting that Jumu protein in the PSC might be responsible for the maintenance of dMyc expression. We then used a temporal and regional gene expression targeting system (Gal^80ts/Gal4 expression system) (McGuire et al., 2003) to knock down jumu in the PSC at different larval stages (Figure 7—figure supplement 3M). Similar to the phenotype observed after the knockdown of jumu in the PSC from embryogenesis, the knockdown of jumu in the PSC from L1 larval stage or L2 stage also caused the reductions in the PSC cell number to different degrees; in fact, the earlier knock down of jumu resulted in a greater reduction in the PSC cell number (Figure 7—figure supplement 3A–F,N). This result indicates a continuous role for Jumu in PSC cell proliferation during larval development. The above results show that the loss of jumu in the MZ is responsible for the ectopic PSC cells detected in the early larval stages (Figure 3A–D). We then used the Gal80ts/Gal4 expression system to knock down jumu in the MZ at different larval stages (Figure 7—figure supplement 3M). The loss of jumu in the MZ from L1 larval stage or L2 stage also caused the ectopic expansion of PSC cells, and an earlier knockdown of jumu resulted in an increased ectopic expansion of PSC cells (Figure 7—figure supplement 3G–L,O). This result further confirms that the expression of Jumu in the MZ prevents the expansion of PSC cell toward the MZ throughout larval development. In conclusion, the expression of Jumu in the MZ and PSC simultaneously maintains the normal development of the PSC. Moreover, changes in the jumu expression levels in the entire lymph gland result in defects in PSC morphology similar to those detected in response to changes in the jumu levels in the MZ but opposite to those caused by changes in the jumu levels in the PSC, indicating that the expression of Jumu in the MZ might play a more prominent role in regulating PSC morphology when jumu is lost or overexpressed in both the PSC and the MZ.

The above-described results suggest that the severe loss of jumu can induce an increase in lamellocytes (Figure 1K–N). We subsequently investigated where and how Jumu controls the generation of lamellocytes. The knockdown of jumu in different zones of the lymph gland did not cause obvious increases in the lamellocytes; in fact, only less than 20% of the lymph gland showed a few lamellocytes (n > 30) (Figure 9—figure supplement 1A–C). However, using e33C-Gal4 to knock down jumu in the entire lymph gland led to the generation of numerous lamellocytes in more than 50% of the lymph gland (n = 32), and similar to jumu mutant, e33C>jumu RNAi also showed reduced numbers of crystal cells (Figure 9A–E). Similarly, more than 60% of lymph gland of the Srp> jumu RNAi lymph gland also showed obvious increased numbers of lamellocytes (n = 36) (Figure 9F,G). Our previous study demonstrated that the overexpression of jumu in both hemocytes and the fat body induces the deposition of hemocytes and melanotic nodules through the activation of Toll signaling (Zhang et al., 2016). The constitutive activation of Toll signaling in hemocytes or the fat body can lead to the differentiation of lamellocytes (Schmid et al., 2014). Moreover, a recent study show that the Toll signaling transcription factors Dorsal and Dorsal-related immunity factor (Dif) are expressed in the lymph gland and that the ectopic expression of both factors in the PSC can cause the generation of lamellocytes in the lymph gland (Gueguen et al., 2013). Thus, we asked whether the lamellocytes caused by the loss of jumu in the lymph gland are linked to Toll signaling. The expression and localization of Dorsal and Dif were detected via antibody staining, and interestingly, we found that Dorsal was enriched in nuclei throughout the lymph glands in the control animals (Srp-Gal4), whereas Dif was located in the cytoplasm and showed high expression levels in the PSC and lower signals in the MZ and CZ (Figure 9L,R). To rule out the effect of the transgenes background of the Srp-Gal4 line on the localization of Dorsal and Dif, we also detected the expression of Dorsal and Dif in the wild-type (w¹¹¹⁸) lymph gland and observed the same results (data not shown). The knockdown of jumu in the entire lymph gland did not affect the expression and localization of Dorsal but caused an activation of Dif, mainly in the CZ and partly in the MZ but not in the PSC; in fact, most cells in the CZ showed obvious nuclei enrichment of Dif (Figure 9M,S; Figure 9—figure supplement 2E–F’). In addition, Dorsal and Dif were not enriched in the nuclei of circulating cells of the Srp>jumu RNAi mutant (Figure 9—figure supplement 2A–D), suggesting that the knockdown of jumu might activate the Toll signaling specifically in the lymph gland. We found that the activation of the Toll pathway throughout the through the overexpression of Toll^10b under the control of Srp-Gal4 also caused a robust increase in the lamellocyte in more than 80% of the lymph glands (n = 30), and also led to enrichment of Dif in the nuclear in the MZ and CZ without affecting the Dorsal expression and localization (Figure 9N,T; Figure 9—figure supplement 2G,G’). To further confirm whether Toll signaling is involved in the generation of lamellocytes in the Srp>jumu RNAi mutant, we performed rescue experiment with a dif mutant. The loss of dif effectively reduced the lamellocytes in the Srp>jumu RNAi mutant; specifically, only less than 10% of the lymph gland generated a few lamellocytes, and the nuclear enrichment of Dif was abolished (n = 36) (Figure 9J,V). Taken together, these results suggest that the activation of Dif in the lymph gland can spontaneously induce lamellocyte differentiation and that Jumu can prevent the activation of Dif and consequently maintain the normal hemocyte differentiation in the lymph gland.

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Similar to the phenotype caused by the loss of jumu in the lymph gland, a previous study indicated that the ectopic expression of col in the entire lymph gland also spontaneously triggers lamellocyte differentiation at the expense of crystal cells spontaneously (Figure 9I) (Crozatier et al., 2004). Moreover, we found that the forced expression of col in different zones of the lymph gland also did not cause an obvious increase in the lamellocytes, with less than 10% of the lymph gland showing a few lamellocytes (n > 30) (Figure 9—figure supplement 1D–F). The above results suggest that Jumu negatively regulates Col expression; thus, we asked whether Col is also involved in the activation of Toll signaling. As expected, col overexpression in the entire lymph gland also resulted in Dif enrichment, mainly in the nuclei of CZ cells, but did not affect Dorsal expression (Figure 9O,U; Figure 9—figure supplement 2H,H’). Moreover, the knockdown of col in Srp>jumu RNAi mutants also inhibited lamellocyte differentiation and nuclear Dif enrichment (Figure 9K,Q and W). Altogether, these results demonstrate that the loss of jumu in the entire lymph gland might indirectly activate Toll signaling by up-regulating the expression of Col, consequently inducing lamellocyte differentiation.

The following mutant alleles, deficiencies, and transgenes were used: jumu^GE27806 and shg^GE13814were purchased from GenExel (Daejeon, South Korea); jumu^Df2.12, jumu^Df3.4 and UAS-jumu were gifts from Alan M. Michelson (Ahmad et al., 2012; Zhu et al., 2012); Hml-Gal4 UAS-2xEGFP, domeless-Gal4 UAS-2xEGFP, col¹/CyO;P(col5-cDNA)/TM6B, Tb, Antp-Gal4/TM3, Sb and Srp-Gal4 were gifts from Utpal Banerjee (Mandal et al., 2007); col-Gal4 UAS-mCD8GFP, domeMESO, hhF4-GFP, UAS-col and UAS-col RNAi were gifts from Lucas Waltzera (Benmimoun et al., 2015); P{en2.4-GAL4}e33C (e33C-Gal4) was a gift from Dominique Ferrandon; Tep-IV-Gal4 was a gift from the Kyoto Stock Center (DGRC); UAS-jumu RNAi (jumu^GD4099) was obtained from the Vienna Drosophila RNAi Center (VDRC); Tub-Gal4, Tub-Gal80^ts (Tub-Gal4^ts), UAS-dmyc RNAi (JF01761) and elav-Gal4 were obtained from the Tsinghua Fly Center; Hand-Gal4 was a gift from Wuzhou Yuan; cdc42-DN was obtained from Bloomington; HHLT-Gal4 was a gift from Jiwon Shim; and UAS-Toll^10b and dif were gifts from Bruno Lemaitre. The w¹¹¹⁸ line or the respective UAS or Gal4 parent lines were used as controls when required. The crosses involving RNAi lines or Tub-Gal4^ts were reared at 29°C, and the other strains and crosses were reared at 25°C. All strains and crosses were cultured on standard cornmeal-yeast medium under a 12 hr light/12 hr dark cycle.

A fusion protein containing a nonconserved portion of the deduced Jumu protein (residues 141–330) was produced in E. coli using the pRSETA (6x-His) vector from Qiagen. The fusion protein was purified using Ni-NTA resin and used to immunize rats.

For antibody staining, the lymph glands were dissected in PBS, fixed with 3.7% formaldehyde in PBS for 20 min, pre-incubated in blocking solution (PBS with 0.1% Tween-20% and 5% goat serum) and then incubated with primary antiserum diluted in blocking solution. The following primary antibodies were used: mouse anti-P1, mouse-anti-L1 (gifts from I. Ando), mouse anti-Hnt (AB_528278), mouse anti-Ptc (AB_528441), Rat anti-Shg (AB_528120), mouse anti-Antp (AB_528082), mouse anti-Dlp (AB_528191) (Developmental Studies Hybridoma Bank), mouse anti-Col (gift from M. Crozatier) (Pennetier et al., 2012), mouse anti-β-galactosidase (Sigma), rat anti-Jumu, rabbit anti-dMyc (Santa Cruz), and mouse anti-P-Mad (gift from Ed Laufer, USA). Alexa Fluor 488-, Alexa Fluor 568- and Alexa Fluor 594-conjugated secondary antibodies (Molecular Probes) were used. ROS detection was performed as previously described (Evans et al., 2014) using dihydroethidium (Invitrogen). Images were obtained using a Zeiss LSM510 confocal microscope or a Zeiss Axioplan 2 microscope equipped with fluorescence optics. All staining was performed in samples from at least three independent experiments.

For quantifications, the lymph glands were scanned at an optimized number of slices (aiming to acquire all staining signals throughout the tissue) using a Zeiss Axioplan 2 microscope, usually less than three optical sections from the Z stack can contain all signal from one tissue. To quantify the crystal cell and ROS indexes and the PSC cell number, the consecutive slices from the same primary lobe were assembled in Photoshop CS6, and the merged images were used for the quantifications. The crystal cell and ROS indexes were calculated as the total number of Hnt⁺ or ROS⁺ cells (quantified using ImageJ, NIH), respectively, in each primary lobe divided by the relative area of the primary lobe. The total PSC cell (Antp⁺ cell) number in each primary lobe was also quantified using ImageJ. At least 30 lymph glands were scored for each genotype. For the quantification of P1⁺ cells and the Shg⁺ and domeMESO⁺ cell indexes, the fluorescent images collected by Zeiss Axioplan 2 microscope were converted to 8-bit images and then recalibrated to obtain an identical threshold using ImageJ. The area with the identical threshold was captured and measured with ImageJ. To measure the total size of the primary lobes, the DAPI-expressing area was captured using freehand selection tool of ImageJ and measured. The plasmatocyte differentiation index was assessed by determining the average area of P1⁺ cells from all optical sections of a primary lobe compared with the area of the primary lobe. The proportion of prohemocytes in the primary lobes (MZ/lobe 1) was defined as the average area of Shg⁺ or domeMESO⁺ cells in all optical sections of a primary lobe compared with the area of the primary lobe. At least 30 lymph glands were analyzed for each genotype. For quantification of the Col, Dlp, dMyc and Jumu fluorescence signal intensities in the MZ or PSC, images were captured with a Zeiss LSM510 confocal microscope at the same exposure times, and at least three confocal slides (4 μm) per stack were analyzed. The total intensity values for Col, Dlp, dMyc and Jumu in each ROI (region of interest) were measured using ImageJ. For the analysis of Col and Dlp staining in the MZ, dome-GFP was used to define the ROIs. For the analysis of dMyc and Jumu staining in the PSC, Antp labeling was used for to define the ROIs. The ROIs in the fluorescent images were captured using the freehand tool and then converted to 8-bit images. The total intensity value of the ROIs with an identical threshold was captured and measured with ImageJ. The average pixel intensity values of dMyc or Jumu in PSC cells (integrated intensity/area of Antp-labeled cells) were calculated. The pixel intensity values of Col or Dlp in the MZ were assessed as the integrated intensity in the ROIs compared with the area of the primary lobe. For each fluorescence quantification experiment, at least 20 lymph glands were analyzed.

ChIP assays were performed as described previously (Kim et al., 2005). In brief, S2 cells (CVCL_Z232, Invitrogen, Cat#R690-07) were transfected with the pMK33-Flag-jumu full CDS construct using the Effectene Transfection kit (Qiagen). S2 cells stably expressing Flag-jumu were fixed in 1% formaldehyde at room temperature. The samples were then sonicated to obtain DNA fragments between 200 and 800 bp in length. The fragmented chromatin was incubated with the anti-Flag antibody (Sigma) or IgG (Santa Cruz) bound to Salmon Sperm DNA/Protein A Agarose Slurry (Upstate) overnight at 4°C. The immunoprecipitated DNA fragments were recovered using a MiniElute purification kit (Qiagen) and analyzed by PCR amplification. Similar results were obtained in three independent ChIP experiments. The primer sequences are shown in Supplementary file 1.

The dmyc promoter −643 to +467 was cloned into the pGL3 basic vector (firefly luciferase reporter plasmid, Promega) using KpnI and XhoI sites. The two base mutations of FKH-binding sites in the dmyc promoter were generated by the overlapping PCR method (Vallejo et al., 2008). The recombinant pGL3 plasmid with the dmyc promoter −643 to +467 was used as a template. The first and second rounds of PCR were performed using a primer pair composed of the forward primer amplifying dmyc promoter −643 to +467 (F) and the reverse mutated primer containing the two base exchanges in the FKH-binding sites (M1R, M2R or M3R) of the dmyc promoter and a second primer pair corresponding to the complementary forward mutated primer (M1F, M2F, or M3F) and the reverse primer amplifying dmyc promoter −643 to +467 (R), respectively. The two amplified overlapping fragments were used as templates for a subsequent overlap extension PCR with the primer pair amplifying the dmyc promoter −643 to +467 (F+R). The primer sequences used are shown in Supplementary file 1. The PCR fragment containing the mutated FKH-binding site was then cloned into the pGL3 basic vector. The introduction of the mutations was verified by DNA sequencing.

A total of 5 × 10⁵ S2 cells (CVCL_Z232, Invitrogen, Cat#R690-07) stably expressing the pMK33-Flag-jumu full CDS construct or empty pMK33 vector were plated per well in a 24-well plate and then co-transfected with 200 ng of dmyc promoter firefly luciferase reporter plasmid and 20 ng of Renilla luciferase pRL-TK reporter vector. The cells were harvested after 48 hr of incubation. Luciferase assays were performed using the Dual-Luciferase Reporter Assay System (Promega), and the normalized luciferase values were calculated as the ratio of the firefly to the Renilla luciferase activities. Plasmid transfection was performed in at least triplicate, and luciferase activity was measured in duplicate or triplicate.

The total RNA from the entire third-instar larvae or the dissected hemocytes (from 300 to 500 third-instar larvae) was isolated with TRIzol (Invitrogen). The obtained total RNA was used to generate cDNA with M-MLV Reverse Transcriptase (Promega). The real-time PCR amplification was performed using SYBR$\text{R}$ Select Master Mix (Applied Biosystems) on a Roche 4500 real-time PCR system. The results were normalized to the level of RpL32 mRNA in each sample. Three experiments per genotype were averaged. A biological replicate was performed, and the same results were obtained. The primer sequences used are shown in Supplementary file 1.

The statistical analyses were performed through two-tailed unpaired Student’s t-tests or one-way ANOVAs using GraphPad Prism software. The threshold for statistical significance was established as *p<0.1, **p<0.01 and ***p<0.001.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Alan M Michelson, Utpal Banerjee, Lucas Waltzera, Dominique Ferrandon, Jiwon Shim, Wuzhou Yuan and Bruno Lemaitre for providing the numerous fly strains used in this study. We acknowledge István Andó, Michèle Crozatier, and Ed Laufer for providing the anti-P1, anti-L1, anti-Col, and anti-P-Mad antibodies. We thank the GenExel Drosophila Stock Center, Kyoto Drosophila Genetic Resource Center, Vienna Drosophila RNAi Center, and Tsinghua Fly Center for sharing the numerous fly stocks utilized in this research. We also thank Michèle Crozatier for the valuable suggestions provided regarding this work. This work was supported by the National Natural Science Foundation of China (31270923).

1. Received: January 13, 2017
2. Accepted: March 20, 2017
3. Accepted Manuscript published: March 28, 2017 (version 1)
4. Accepted Manuscript updated: April 3, 2017 (version 2)
5. Version of Record published: April 13, 2017 (version 3)

© 2017, Hao et al.

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