Notch signaling activity regulates hematopoiesis in Drosophila and vertebrates alike. Parasitoid wasp infestation of Drosophila larvae, however, requires a rapid downregulation of Notch activity to allow the formation of encapsulation-active blood cells. Here we show that the Drosophila CSL transcription factor Suppressor of Hairless [Su(H)] is phosphorylated at Serine 269 in response to parasitoid wasp infestation. As this phosphorylation interferes with the DNA-binding of Su(H), it reversibly inhibits Notch activity. Accordingly, phospho-deficient Su(H)S269A mutants are immune compromised. A screen for kinases involved in Su(H) phosphorylation identified Pkc53E, required for normal hematopoiesis as well as for parasitoid immune response. Genetic and molecular interactions support the specificity of the Su(H)-Pkc53E relationship. Moreover, phorbol ester treatment inhibits Su(H) activity in vivo and in human cell culture. We conclude that Pkc53E targets Su(H) during parasitic wasp infestation, inducing downregulation of Notch activity, thereby remodeling the blood cell population required for wasp egg encapsulation.
The study by Deichsel et al. reports valuable findings that suggest a new, possibly conserved, mechanism by which post-translational modification of a Notch regulator mediates the cellular immune response. However, the claims are only partially supported as the data and analysis are incomplete. The work will be of interest to biologists working on immune cell development or regulation of Notch.
Drosophila melanogaster harbours a sophisticated cellular immune system to fight invaders. The larval hematopoietic system comprises a circulating and sessile compartment of embryonic origin and the developing larval lymph gland, constituting a hematopietic organ. The circulating and sessile compartment consists primarily of macrophage-like plasmatocytes, plus a small number of crystal cells involved in wound healing and melanization responses to neutralize pathogens. Both cell types differentiate from hemocyte precursors within the lymph gland as well; they are released during pupal stages to serve the adult with immune cells. Finally, lamellocytes represent the third blood cell type in Drosophila. While merely absent in healthy animals, their formation is induced within hours of wasp infestation or wounding (reviewed in: Banerjee et al., 2019; Letourneau et al., 2016; Hultmark and Andó, 2022). In fact, parasitism by endo-parasitoid wasps represents one of the most severe naturally occurring immune challenges to Drosophila, invariably causing death if not defended off properly. Parasitoid wasps deposit their egg into the live Drosophila larva, where it develops into the next wasp generation egressing from the pupa instead of a fly. Hence, wasp infestation is a life-threatening challenge to the Drosophila host, demanding an immediate immune response to overwhelm the parasite. This involves a massive increase in circulating hemocytes, and most energy resources are devoted to fight the invader. Here, lamellocytes play a critical role: they encapsulate the wasp egg and, by expressing Prophenoloxidase, induce a melanization reaction retarding further wasp development (Dudzic et al. 2015; reviewed in: Banerjee et al., 2019; Letourneau et al., 2016; Hultmark and Andó, 2022).
Wasp infestation dramatically remodels the composition of the Drosophila hemocyte population (Cho et al., 2020; Tattikota et al., 2020; reviewed in: Csordás et al., 2021). There is a vast increase in plasmatocytes and intermediate precursors in both hematopoietic compartments, from which lamellocyte differentiate, requiring the combined activity of several pathways, including JAK/STAT, JNK, Toll, Notch, EGFR and INR pathways, which in fact regulate blood cell homeostasis in general (reviewed in Banerjee et al., 2019; Letournau et al., 2018; Csordás et al., 2021). Simultaneous to the massive expansion of lamellocytes, crystal cells derived from the same precursors are significantly reduced (Crozatier et al., 2004; Krzemien et al., 2010; Ferguson and Martinez –Agosto, 2014; Cho et al., 2020; Tattikota et al., 2020; reviewed in Csordás et al., 2021). Crystal cells are generated both in the larval lymph gland as well as by trans-differentiation from plasmatocytes in the sessile compartment. Their formation, differentiation and survival strictly depend on Notch signaling activity (reviewed in: Banerjee et al., 2019; Csordás et al., 2021; Hultmark and Andó, 2022). Lamellocyte and crystal cell lineages are mutually exclusive. Accordingly, while promoting crystal cell fate, Notch activity inhibits differentiation of lamellocytes within the lymph gland, however, is reduced upon wasp infestation (Small et al., 2014). Lamellocyte induction involves the formation and sensing of reactive oxygen species triggered by wasp egg injection (Nappi et al., 1995; Sinenko et al., 2011; Small et al., 2014; Louradour et al., 2017; reviewed in: Banerjee et al., 2019; Csordás et al., 2021). The molecular mechanism underlying the simultaneous crystal cell fate inhibition, however, is less well understood. Obviously, an effective and rapid downregulation of Notch activity is required to fend off parasitic wasps.
The Notch pathway is highly conserved between invertebrates and vertebrates, where it regulates numerous cell fate decisions. Notch signals are transduced by CSL type proteins (abbreviation of human CBF1/RBPJ, Drosophila Suppressor of Hairless [Su(H)], and worm Lag1), that bind the DNA to direct transcriptional activity of Notch target genes by help of recruited cofactors (reviewed in: Giaimo et al., 2021; Kopan and Ilagan, 2009; Siebel and Lendahl, 2017). Hence, CSL is central to Notch pathway activity as no signal transduction could occur in its absence or in the instance of a lack of DNA-binding. Earlier, we observed CSL, i.e. Su(H) phosphorylation at Serine 269 in cultured Drosophila Schneider S2 cells (Nagel et al., 2017). Of note, Schneider S2 cells have hemocyte characteristics (Cherbas et al., 2011; Schneider, 1972; Terriente-Felix et al., 2013). As a consequence of the negative charge conferred by the phosphorylation at Serine 269, Su(H) loses its affinity to the DNA, and without its DNA-binding ability, also its function as a transcriptional regulator (Nagel et al., 2017). Accordingly, a phospho-mimetic Su(H)S269D mutant behaved like a Su(H) loss of function mutant in all respects. In contrast, the phospho-deficient Su(H)S269A variant, appeared wild type at the first glance, indicating some tissue-specificity of this regulatory mechanism. In fact, the Su(H)S269A allele displayed a gain of Notch activity particularly during embryonic and larval hematopoiesis with increased numbers of crystal cells in both hematopoietic compartments (Frankenreiter et al., 2021). Moreover, we found that the general Notch antagonist Hairless is not involved in constraining crystal cell numbers, suggesting that in the context of blood cell homeostasis Notch activity is regulated by the phosphorylation of Su(H) (Maier, 2006; Frankenreiter et al., 2021). As the same set of genes affect blood cell maintenance and differentiation in homeostasis and upon immune challenge (Cho et al., 2020; Tattikota et al., 2020), obviously phospho-mediated downregulation of Notch activity might also occur in response to parasitoid wasp infestation, allowing the formation of encapsulation active lamellocytes at the expense of crystal cells. Briefly, phosphorylation of Su(H) is an elegant mechanism to promptly and transiently curb Notch activity.
In this work, we followed the tempting hypothesis that following wasp infestation, a specific kinase might be activated to phosphorylate Su(H) thereby blocking Notch activity. In this case, the phospho-deficient Su(H)S269A allele should be immune-compromised, as it cannot respond to phosphorylation, i.e. remaining active even upon infection. Indeed, Su(H)S269A displayed an increased sensitivity towards parasitoid wasp infestation accompanied by an increase of crystal cells at the expense of lamellocytes. Accordingly, phosphorylation of Su(H) protein was detected in infested wild type larvae but not in the Su(H)S269A mutant. In a screen for kinases regulating this process, Pkc53E, the homologue of human PKCα, was identified as an important player. In agreement with a role in blood cell homeostasis, a Pkc53EΔ28 null mutant displayed increased crystal cell numbers. Moreover, genetic and molecular interactions between Su(H) and Pkc53E support the specificity of their relationship. Finally, Pkc53EΔ28 was impaired in its immune response to wasp infestation as well. Together, these data show that Su(H) is a target of Pkc53E during parasitic wasp infestation, inducing phosphorylation and subsequent downregulation of Notch activity to allow the mass production of lamellocytes required for wasp defense.
Impaired immune response to parasitoid wasps in the phospho-deficient Su(H)S269A mutant
Wasp infestation alters the course of hematopoiesis, as lamellocyte differentiation is massively increased at the expense of crystal cells. This process requires a rapid downregulation of Notch activity that may be implemented by the phosphorylation of Su(H) at Serine 269 (S269). To test this model, we exposed control larvae and larvae of the phospho-deficient Su(H)S269A variant to the parasitoid wasp Leptopilina boulardi (L. boulardi) and analysed the consequences on blood cell homeostasis two days post-infection. To exclude any influence of the engineered genomic background, we used Su(H)gwt for the comparison, carrying a genomic wild type construct in place of the mutant (Praxenthaler et al., 2017; Frankenreiter et al., 2021).
Earlier we noted an excess of crystal cells in the phospho-deficient Su(H)S269A allele, which we interpret as a gain of Notch activity in consequence of the inability to be downregulated by Su(H) phosphorylation (Frankenreiter et al., 2021) (Fig. 1A,B). In response to wasp infestations, however, crystal cell numbers should drop to allow formation of lamellocytes (Small et al., 2014; Csordás et al., 2021). Indeed in the Su(H)gwt control, both the sessile crystal cells as well as those within the larval lymph glands were significantly lessened in response to wasp infestation (Fig. 1A,B). In contrast, the higher crystal cell numbers in the Su(H)S269A mutant larvae dropped to control-level only, demonstrating the inability of the mutant to detect this immune challenge or to respond to it (Fig. 1A,B).
Increased abundance of lamellocytes upon wasp infestation was monitored in vivo in larval hemolymph and lymph glands, using either the L1-atilla-GFP reporter (Honti et al., 2009) or PPO3-Gal4::UAS-GFP (Dudzic et al., 2015), respectively. In the harvested larval hemolymph of the infested Su(H)gwt control, the hemocytes contained about 15% PPO3- and 23% atilla-labelled lamellocytes (Fig. 1C,D). These numbers were significantly lower in the wasp infested Su(H)S269A larvae (Fig. C,D). Consistently, both lamellocyte reporters were robustly induced in the lymph glands of the infested control, but rarely in glands of the Su(H)S269Amutant (Fig. 1E,F). Obviously, the Su(H)S269A mutant barely responds to the immune challenge raised by the parasitic wasp infestation. Overall, these data support the model that S269 in Su(H) is a molecular target for a kinase, phosphorylated upon immune challenge to downregulate Notch activity, thereby allowing lamellocyte formation at the expense of crystal cells.
The phospho-deficient Su(H)S269A allele cannot combat wasp infestation
Parasitoid wasp infestation constitutes an extreme immune challenge for the Drosophila larva: if not combatted by the immune system, a wasp egg, which is deposited in the larval body cavity, will develop into an adult wasp, thereby killing the larval host during pupal stage. Indeed, depending on the wasp species used, we measured a high mortality rate with less than 5% up to about 15% of surviving flies (Fig. 2A). According to our working hypothesis, the phospho-deficient Su(H)S269A variant should not be able to properly respond to the immune challenge. To test this directly, we measured the survival of Su(H)S269A animals upon wasp infestation compared to the wild type control.
The two closely related wasp species L. boulardi and Leptopilina heterotoma (L. heterotoma) very efficiently parasitized both the control Su(H)gwt and the Su(H)S269A variant, though the latter appeared slightly more sensitive (Fig. 2A). The difference in mortality became more apparent with the wasp species Asobara japonica (A. japonica), allowing 14.5% of the Su(H)gwt control flies to escape parasitism, whereas only 3.8% of the infested Su(H)S269A pupae emerged as flies. Thus, the Su(H)S269A mutants are less robust in resisting parasitoid wasp infestation consistent with a defective immune response.
Serine 269 of Su(H) is phosphorylated upon wasp infestation
Next, we wanted to directly monitor Su(H) phosphorylation at S269 in response to wasp infestation. To this end, polyclonal antibodies directed against a phosphorylated peptide containing the sequence motif NRLRpSQTVSTRYLHVE were generated (⍺-pS269). Specificity was first tested by Western blot analysis using bacterially expressed GST fusion proteins containing the entire beta-trefoil domain (BTD) of Su(H), as well as with phospho-mimetic (S269D) and phospho-mutant (S269A) versions. All three variants were detected by the antisera. The S269D version, however, was strongly preferred, indicating that this antibody does preferably recognize phospho-S269 Su(H) protein (Fig. 2 - figure supplement 1). Encouraged by this result, we used this antiserum on protein extracts derived from larvae infested and not infested by L. boudardi. For this experiment we used genome engineered fly strains expressing mCherry-tagged Su(H) proteins, Su(H)S269A-mChand Su(H)gwt-mChfor control (Praxenthaler et al., 2017). Su(H) proteins were trapped using RFP-nanobody coupled agarose beads, and the precipitates were then probed in Western blots with antibodies directed against mCherry and pS269. Indeed, ⍺-pS269 antibodies recognized Su(H)gwt-mCh protein specifically in wasp infested larvae, indicating respective phosphorylation of Su(H) protein. No such signals were seen in precipitates from the non-infected larvae nor from any of the Su(H)S269A-mCh mutant larvae (Fig. 2B). These data clearly show that wasp infestation induced the phosphorylation of Su(H) at S269. Apparently, parasitoid wasp infestation starts a cascade of events resulting in the inhibitory S269 phosphorylation of Su(H) protein to promptly interrupt Notch signalling activity and to allow lamellocyte formation. Hence, the question arose on the kinase/s involved in this process.
Screening of kinase candidates mediating phosphorylation of Su(H) at Serine 269
To identify Ser/Thr kinases involved in the phosphorylation of Su(H) at S269, we commenced with a combination of in silico and biochemical approaches aiming to generate a list of candidate kinases which can be further analysed by genetic means (Fig. 3). First, by using GPS 3.0 software that encompasses a substantial database of kinases and their preferred recognition motives (Xue et al., 2011), 36 potential human kinases were predicted to recognize S269 as substrate, represented by 30 kinases in Drosophila (Fig.3 – supplement Table 1). In addition, the BTD domain of Su(H) was bacterially expressed as a GST fusion protein and subjected to phospho-assays using 245 different human Ser/Thr kinases. Our reasoning for using the entire BTD domain was to ensure a normal folding of the domain (Kovall and Blacklow, 2010), and to reduce the number of potential phospho-sites at the same time present in full length Su(H). With this approach, we ended up with 62 human Ser/Thr kinases (25% of the tested kinases) that use the BTD of Su(H) as an in vitro substrate. These kinases correspond to 40 different kinases in Drosophila belonging to six different kinase families (Fig.3 –supplement Table 2). Ten of these kinases were also predicted in silico, representing members of the AGC, CAMK, CMGC, STE and OPK family of kinases (Fig.3 –supplement Table 2). Both sets of data, biochemical and bioinformatics, were used to generate a list of 44 candidates to be analysed by genetic means (Fig.3 –supplement Table 3). The candidates were further screened for an imbalanced hematopoiesis. We reasoned that mutants affecting a relevant kinase gene involved in the phosphorylation of Su(H) should display increased crystal cell numbers similar to what was observed in the Su(H)S269A mutant (Frankenreiter et al., 2021). Larvae of 13 different kinase mutants and the progeny of 44 UAS-RNAi and/or UAS-kinase dead transgenes crossed with hml-Gal4, a blood cell-specific Gal4 driver line, were tested (Fig.3 –supplement Table 3). To this end, the larvae were subjected to heating. This procedure allows a comfortable visualization and quantification of mature larval crystal cells through the cuticle, which blacken due to the activation of Prophenoloxidase (Rizki, 1957; Lanot et al., 2001). Fourteen kinase mutants tested were similar to the control, whereas mutations in four kinases impeded crystal cell development or prevented it altogether. Unexpectedly, the majority of the tested kinase mutants exhibited elevated crystal cell numbers, however to a different degree (Fig.3 –supplement Table 4). Seven kinase mutants substantially exceeded crystal cell numbers seen in Su(H)S269A larvae; they were hence excluded from further analysis. Nineteen kinase mutants matched closely the Su(H)S269A phenotype, making those the most promising candidates for being involved in the phosphorylation of Su(H) at S269. Six of those were within the cluster of ten candidates singled out by the in silico and the in vitro screens (Fig. 3 A,B). Using commercially available, activated human kinases, we were able to test five candidates, AKT1, CAMK2D, GSK3B, S6 and PKCα in vitro by MS/MS analysis on the Su(H) peptide ALFNRLRS8QTVSTRY, where Serine 8 corresponds to S269 in Su(H). Only PKCα unambiguously phosphorylated the given peptide at Serine 8. Whereas GSK3B did not phosphorylate the peptide at all, AKT1, CAMK2D and S6 piloted Threonine 10, corresponding to Threonine 271 in Su(H) (Fig. 3D-figure supplement 1). Together, these data support the idea, that PKCα corresponding to Pkc53E in Drosophila, is part of the kinase network mediating the phosphorylation of Su(H) at S269.
Role of Pkc53E in the phosphorylation of Su(H)S269
Confirming the MS/MS data, human PKCα was able to phosphorylate the respective Su(H) peptide (Swt) similar to its defined pseudo-substrate PS (Kochs et al., 1993), whereas the S8A mutant peptide (SSA) was accepted only half as well in an ADP-GloTM assay, indicating that S269 is a preferred substrate (Fig. 4A). Bacterially expressed and purified Drosophila Pkc53E, however, did neither accept the PS nor the Su(H) peptides (Fig. 4B). Pkc53E activity, however, was stimulated by the agonistic phorbol ester PMA (phorbol 12-myristate 13-acetate) (Blumberg et al., 1983; Nakashima, 2002) in phosphorylating the PS and Su(H) peptide Swt but not the S8A mutant peptide SSA (Fig. 4C). To generate an activated form of Pkc53E, we exchanged four codons by in vitro mutagenesis, three (T508D, T650D and S669D) mimicking phosphorylation in the kinase and C-terminal domains, respectively, and one in the pseudo-substrate domain (A34E) (Fig.4 – figure supplement 1) (Gould and Newton, 2008). The resultant Pkc53EEDDD protein accepted the PS, and the Su(H) peptide Swt even better, but not the S8A mutant peptide SSA, demonstrating the specificity of the phosphorylation (Fig. 4D). As predicted for a fully activated kinase, PMA was unable to boost Pkc53EEDDD protein activity any further (Fig. 4E).
The PKC-agonist PMA influences Su(H) activity and blood cell homeostasis
Our data so far indicated that Su(H) is a phospho-target of Pkc53E which reduces its activity by affecting its DNA-binding. In this case, we might expect an influence of the general PKC-activator PMA on both, Su(H) activity as well as Notch-mediated crystal cell formation. We tested the former in a RBPJko HeLa cell system (Wolf et al., 2019), measuring Notch-reporter gene activation by Su(H)-VP16, as this Su(H) variant is independent of Notch activity itself, allowing to directly monitor the influence of PMA on Su(H) activity. Indeed, Su(H)-VP16 ability to activate reporter gene transcription was reduced by more than half in the presence of PMA (Fig. 5A). This is in agreement with an PMA-mediated activation of endogenous PKC in the transfected HeLa cells, resulting in Su(H)-VP16 phosphorylation, loss of DNA-binding activity and reduced transcriptional activation. Accordingly, repression could be reverted by the addition of kinase-inhibitor Staurosporine (Karaman et al., 2008). In fact, Staurosporine alone already increased Su(H)-VP16 transcriptional activity, indicating that PKC-driven phosphorylation occurs to a considerable degree in the normal situation in HeLa cells already (Fig. 5A).
Next, we assayed the effect of PMA on larval crystal cell formation. If, as expected, PMA increased Pkc53E activity, Su(H) should be inactivated by phosphorylation with decreased crystal cell numbers as a consequence. We fed PMA to Drosophila larvae and assayed the numbers of sessile crystal cells. Indeed, crystal cell numbers dropped to nearly zero, similar of what is observed in the phospho-mimetic Su(H)S269Dmutant (Frankenreiter et al., 2021), suggesting a very efficient phosphorylation and inactivation of Su(H) protein (Fig. 5B). In contrast, Su(H)S269A mutant larvae displayed an increased number of crystal cells which can be attributed to the fact that here, Su(H) can no longer be phosphorylated and hence, is overactive in this context. Feeding PMA to Su(H)S269A larvae only caused a drop of excessive crystal cell numbers down to a wild type level (Fig. 5B). In conclusion these data show that Su(H) activity is regulated in vitro and in vivo by PKC activity in the context of blood cell homeostasis.
Pkc53E is required for normal blood cell homeostasis in Drosophila larvae
Su(H)S269A mutant larvae develop an excess of crystal cells, both in the hemolymph as well as in the lymph glands, due to a failure to downregulate respective Notch activity in the hemocyte precursors via the phosphorylation of Su(H) protein (Frankenreiter et al., 2021) (see Fig. 1A). Assuming Pkc53E has a major role in the phosphorylation of Su(H), we should expect a similar phenotype in a Pkc53E mutant due to the inability to phosphorylate any substrate. In order to test this assumption directly, we assessed the number of crystal cells in larval lymph glands as well as in sessile crystal cells in several loss of function backgrounds of Pkc53E. To this end, we used the Pkc53EΔ28 allele, which by RT-PCR is a null mutation (Fig. 6 – figure supplement 1). In addition, we used two different RNAi-lines and one sgRNA line under UAS-control to knock down Pkc53E activity specifically within the developing lymph gland using lz-Gal4 and within the hemocytes using hml-Gal4, respectively (Lebestky et al., 2000). In any context tested, the number of crystal cells was strongly increased matching those of the Su(H)S269A mutant (Fig. 6). The similar phenotypes imply that Pkc53E acts through the phosphorylation of Su(H). However, as outlined above, the majority of kinase mutants displayed increased crystal cell numbers, raising the possibility of a fortuitous accordance. If the increase of crystal cell numbers in Pkc53E mutants is independent of Su(H) phosphorylation, we should expect an additive effect if we combine the two mutants. The double mutants Su(H)S269A Pkc53EΔ28 were generated by genetic recombination; they displayed the same range of excessive crystal cell numbers as the single mutants (Fig. 7A,B). Moreover, the strongly reduced number of crystal cells observed in the overactive Su(H)S269D mutant was not increased by Pkc53EΔ28 (Fig. 7A,B), indicating that Pkc53E indeed acts upstream of Su(H), or directly on Su(H). If the latter is the case, we may expect the two proteins to form complexes in vivo. Indeed, we could co-precipitate Su(H)-Pkc53E protein complexes, both from Drosophila heads containing hemocytes (Sanchez Bosch et al., 2019), as well as from the larval hemolymph (Fig. 7C,D). Specific co-precipitation was eased by using fly strains expressing m-Cherry tagged Su(H) (Praxenthaler et al., 2017) and HA-tagged Pkc53E, respectively. Together, these data demonstrate that Pkc53E has an important role in blood cell homeostasis that can be largely explained by its activity to phosphorylate Su(H), thereby regulating Notch activity during hemocyte and lymph gland development.
Pkc53E mutants are immune compromised
According to our hypothesis, Pkc53E phosphorylates Su(H) in response to an immune challenge by parasitic wasp infestation. Hence, we would expect that a loss of Pkc53E function should affect the ability of Drosophila larvae to fight wasp infestations similarly to the Su(H)S269A mutant. Indeed, when the Pkc53EΔ28 null mutant was infested with the parasitic wasp L. boulardi, lamellocyte numbers in the hemolymph did not reach wild type levels, and they were absent from the larval lymph glands (Fig. 7E,F). Apparently, the Pkc53EΔ28 null mutant is unable to recognize parasitic wasp infestation or is unable to respond to it, for example by the phosphorylation of Su(H), demonstrating the involvement of this kinase in the immune response of Drosophila to parasitoid wasp infestation.
The Notch pathway is highly conserved between invertebrates and vertebrates, with regard to both the underlying molecular principles as well as the biological processes it is involved, including hematopoiesis and immune defence. During mammalian hematopoiesis, Notch plays a fundamental role in stem cell maintenance and proliferation as well as in the differentiation of blood cell precursors, notably in T-cell development. Accordingly, aberrant Notch signaling activity has profound consequences for blood cell homeostasis that may result in leukemia (reviewed in: Radtke et al., 2010; Siebel and Lendahl, 2017; Banerjee et al., 2019; Gallenstein et al., 2023). Hence, the principles of the regulation of Notch signaling activity are of great interest.
Notch signals are transduced by CSL-type DNA-binding proteins that are pivotal to Notch pathway activity, as no signal transduction could occur in their absence or in the instance of a lack of DNA-binding. CSL proteins are extremely well conserved. They act as a molecular switch, either activating or repressing Notch target genes, depending on the recruited cofactors. Upon ligand binding, the Notch receptor is cleaved, and the biologically active Notch intracellular domain (NICD) is released, to assemble an activation complex with CSL and further co-factors. Notch pathway repression, however, entails for example the recruitment of co-repressors by CSL; in Drosophila mediated by the binding of Hairless (reviewed in: Maier, 2006; Borggrefe and Oswald, 2009; Kovall and Blacklow, 2010; Bray, 2016; Giaimo et al., 2021).
Studies of Drosophila hematopoiesis have uncovered a pleiotropic role of Notch in all the hematopoietic compartments, where Notch activity needs to be precisely regulated to ensure blood cell homeostasis (reviewed in: Banerjee et al., 2019; Csordás et al., 2021). During blood cell formation, Notch directs the crystal cell lineage. Accordingly, a downregulation of Notch activity causes a lack of crystal cells, whereas a gain of Notch activity results in increased numbers (Duvic et al., 2002; Lebestky et al., 2003; Terriente-Felix et al., 2013; Gosh et al., 2015; Frankenreiter et al., 2021). In our earlier work, we have shown that the regulation of Notch activity during hemocyte differentiation is independent of the general Notch antagonist Hairless. Instead, it relies on the post-translational modification of the Drosophila CSL protein Su(H) (Frankenreiter et al., 2021). Phosphorylation at Ser269 in the beta-trefoil domain of Su(H) impedes DNA-binding activity, and hence the capability of Su(H) to act as a transcriptional regulator in the context of blood cell development, without affecting Su(H) protein expression (Nagel et al., 2017; Frankenreiter et al., 2021). Accordingly, phospho-deficient Su(H)S269A mutants develop an excess of crystal cells. Now we provide evidence that Su(H) phosphorylation is likewise involved in parasitoid wasp defense, pointing to a dual use of the Notch pathway in blood cell homeostasis as well as in stress response, which is typical for the myeloid system (Banerjee et al., 2019).
The primary cellular immune response of Drosophila to fight parasitoid wasp infestation is an encapsulation of the parasite egg to terminate its further development. Albeit metabolically extremely costly, Drosophila larvae have to remodel their entire hematopoietic system to generate the masses of lamellocytes required for the encapsulation (reviewed in Letournau et al., 2016; Kim-Jo et al., 2019; Csordás et al., 2021; Hultmark & Ando, 2022). There are two major sources for the lamellocytes. One is the transdifferentiation of circulating plasmatocytes, released from the sessile compartment and proliferating upon immune challenge (Márkus et al., 2009; Honti et al., 2010; Stofanko et al., 2010; Vanha-Aho et al., 2015; Anderl et al., 2016). Second is a massive expansion of prohemocytes in the lymph gland followed by a differentiation to lamellocytes and their release due to the premature disintegration of the gland (Lanot et al., 2001, Sorrentino et al., 2002, Louradour et al., 2017; Cho et al., 2020). Wasp infestation hence provokes a biased commitment to the lamellocyte lineage at the expense of crystal cells, as the two lineages are mutually exclusive (Krzemien et al., 2010; Tattikota et al., 2020; Cho et al., 2020).
The processes underlying the Drosophila immune response to wasp parasitism are well-understood, sharing many molecular details with the inflammatory responses of vertebrates (reviewed in Letourneau et al., 2016; Kim-Jo et al., 2019; Banerjee et al., 2019; Kharrat et al., 2022). Interestingly, sterile injury of Drosophila larvae initiates a likewise immune response covering all aspects of wasp-mediated immune challenge (Evans et al., 2022). The epidermal penetration by the wasp ovipositor causes a burst of hydrogen peroxide at the injury site via the activation of the NADPH oxidase DUOX. The oxidative stress then induces a systemic activation of Toll/NF-κB and JNK-signaling within circulating hemocytes as well as within the cells of the posterior signaling center of the lymph gland. Following cytokine release, JAK/STAT and EGFR signalling pathways are activated non-cell autonomously driving the trans/differentiation of pro/hemocytes to lamellocyte fate and lymph gland dispersal (Nappi et al., 1995; Schlenke et al., 2007; Sinenko et al., 2011; Gueguen et al., 2013; Razzell et al., 2013; Louradour et al., 2017; Chakrabarti and Visweswariah, 2020; Evans et al., 2022; reviewed in Letourneau et al., 2016; Banerjee et al., 2019). Whereas the switch to lamellocyte fate is well elaborated, less is known about the processes underlying the simultaneous suppression of crystal cell fate. Obviously, this step requires a downregulation of Notch signalling activity, as the Notch pathway is instrumental to crystal cell fate in all the hematopoietic compartments (Duvic et al., 2002; Lebestky et al., 2003; Schlenke et al., 2007; Small et al., 2014; reviewed in Banerjee et al., 2019). Our work now reveals that a parasitoid wasp attack causes a phosphorylation of Su(H) on Ser269 as means of an efficient, rapid and reversible way to inhibit Notch activity. This idea is consistent with earlier observations, whereby wasp parasitism quenched the expression of a Su(H)-lacZ reporter (Small et al., 2014). Moreover, we provide evidence that Pkc53E is involved in Su(H) phosphorylation during blood cell development as well as during parasitoid wasp-defense in Drosophila. The antagonistic role of Pkc53E in blood cell homeostasis is fully consistent with earlier reports demonstrating that a loss of Pkc53E activity increased the expansion of prohemocytes in a genetic model of myeloid leukemia in Drosophila (Sinenko et al., 2010). Moreover, the involvement of PKCs in wasp defense is not completely unexpected given their well-documented role in the immune responses of mammals and Drosophila alike. For example, Drosophila flies ensure a healthy gut-microbiota homeostasis by modulating DUOX activity as a pathogen-specific defense line (reviewed in Kim and Lee, 2014). DUOX activation and ROS production, induced by gut pathogens was shown to be mediated by Pkc53E and Ca2+ via phospholipase Cβ (Ha et al., 2009; Lee et al., 2015). Sterile wounding of the Drosophila embryo triggers an instantaneous Ca2+ flash followed by hydrogen peroxide production (Razzel et al., 2013). It is conceivable that the epidermal breach by the wasp’s sting similarly induces a Ca2+ flash that may spark Pkc53E activation as a result. However, PKCs may be activated directly by oxidative modification even in the absence of Ca2+. Moreover, they may promote endogenous ROS production in a positive feed back loop (reviewed in Cosentino-Gomes et al., 2012), that could support the systemic response in the larval lymph gland.
The role of PKCs in the mammalian hematopoietic and immune systems is well documented. PKCs in this context act redundantly, and presumably, this holds also for Drosophila where several PKCs exist. Upon activation, PKCs may translocate into the nuclear compartment, where they can directly influence gene expression programs, for example by piloting chromatin factors as substrates, pivotal for regulating immune cell differentiation (reviewed in Lim et al., 2015). PKCs, including PKCα, are present in CD34+ long-term hematopoetic stem cells. Moreover, specific roles in both the myeloid and the multilymphoid lineages are known. Notably, some PKC isoforms including PKCα may play a role in Notch-dependent T-cell development. Notch pathway activity is indispensable for T-cell lineage commitment of early thymic progenitors, however, is rapidly downregulated after the β-selection phase. As treatment with phorbol ester upregulates the transcription of TCRα and β chains, a role of PKC in Notch-dependent commitment of αβ T-cells has been proposed. With the expressing of the invariant TCR components, fate commitment of T-cells driven by Notch signaling is completed. Subsequent cell survival and differentiation into CD4+CD8+ double positive cells, however, depends on the formation of a pre-TCR complex and requires PKCα activity (reviewed in Altman and Kong, 2016; Yui and Rothenberg, 2014). During initial T-cell development, Notch1 signaling increases in intensity in the pre-β-selected thymocytes due to an autoregulatory feedback loop directly controlled by E proteins and by Notch1-CSL itself. Following β-selection, as Notch signaling activity becomes dispensable, Notch1 is abruptly downregulated at the transcriptional level. This rapid downregulation is mediated by the inhibition of the E proteins (Yashiro-Ohtani et al., 2009), however, might in addition involve the phosphorylation of CSL by PKCα. Cell fate in more mature thymocytes is then fixed by the silencing of Notch target genes through chromatin modulators (reviewed in Yui and Rothenberg, 2014).
Appropriate silencing of Notch signaling activity in the course of T-cell development is of utmost importance, as prolonged Notch activity during β-selection predisposes the T-cells to leukemic transformation. Phosphorylation of CSL proteins by PKCα kinase offers a way for a rapid and reversible deactivation of Notch signals not only in Drosophila but also in the mammalian system. In fact, the amino acid sequences harboring the respective Serine residue in the CSL beta-trefoil domain are completely conserved between vertebrates and invertebrates (Wilson and Kovall, 2006; Nagel et al., 2017), raising the possibility of a likewise regulatory mechanism in mammalian hematopoiesis and immunity. Indeed, respective phosphorylation of the human CSL protein at the homologous position Ser195 was observed in human embryonic stem cells, where differentiation was induced by phorbol ester treatment (Rigbolt et al., 2011), consistent with an involvement of PKCα in this context as well. In conclusion, our work uncovers an important role for PKC-mediated downregulation of Notch activity via CSL-phosphorylation in blood cell homeostasis and in the immune response to parasitoid wasp infestation in Drosophila. Future work may uncover, whether the same mechanisms apply to mammalian hematopoiesis and immunity as well.
Materials and Methods
Key resources are listed in S1 Table
Maintenance of parasitoid wasps and infection assay of Drosophila melanogaster
Leptopilina boulardi (L. boulardi), Leptopilina heterotoma (L. heterotoma) and Asobara japonica (A. japonica) were kindly provided by B. Häußling and J. Stöckl, Bayreuth, Germany (Weiss et al., 2015). Wasp species were co-cultured with wild-type Drosophila larvae at room temperature. To this end, about forty 3-5 days old female wasps and twenty male wasps were co-incubated with second instar larvae for 5-7 days at room temperature. Every other day, fresh drops of honey water were added to the vial plug for feeding the wasps. After two weeks, all hatched flies were discarded. Wasps emerge about 30 days after the infestation.
For the infection assays, 50-100 staged Drosophila late second/early third instar larvae of the respective genotype were transferred onto apple juice plates with fresh yeast paste. 30 females and 20 males of the wasps aged between 3-6 days were added to the larvae, allowing to infect them for 4-6 hours. Afterwards, wasps were removed and infected larvae were allowed to develop further in vials with normal fly food. Wasps were only used once for each infection. After infestation, third instar larvae were prepared, or the survival rate of wasps versus Drosophila imago was recorded.
Fly work and genetic analyses
Fly crosses were performed with 30-40 virgin females and 20 males to avoid overcrowding and stress. Combination/recombination of fly stocks was monitored by PCR-genotyping using primers listed in S2 Table. A complete list of the Kinase mutant flies tested in the ‘larval kinase screen’ is found in Fig.3 – supplement Table 3. As reporter lines served atilla-GFP (BL23540) and PPO3-Gal4 UAS mCD8-GFP (named PPO3::GFP, Dudzic et al., 2015). For Gal4/UAS based overexpression and RNAi-mediated knockdown, we used hml-Gal4 (BL30141), lz-Gal4 (Lebestky et al., 2000; obtained form M. Crozatier, Université de Toulouse, France), UAS-μMCas9 (VDRC 340002), UAS-HA-Pkc53E (this study), UAS-white-RNAi (BL31231) and UAS-sgRNA-Pkc53E (VDRC341127). The strain vasa-ϕC31, 96E-attB/TM3 (Bischof et al., 2007) served for the generation of UAS-HA-Pkc53E flies. Su(H) controls and mutants comprised: Su(H)gwt, Su(H)gwt-mCh, Su(H)S269A and Su(H)S269D/CyO-GFP, (Praxenthaler et al., 2017; Frankenreiter et al., 2021). Su(H)S269A-mCh flies were produced in this study by a C-terminal in frame fusion of mCherry to the Su(H)S269A mutant gene followed by genomic integration of the construct via gene engineering as outlined before (Praxenthaler et al., 2017).
Generation of the UAS-HA-Pkc53E fly line
Pkc53E cDNA (DGRC GH03188) was PCR amplified and subcloned via XhoI/XbaI in a modified pBT-HA vector, harbouring three copies of an HA-Tag generated via annealed oligos cloned into Acc65I/XhoI to generate pBT-3xHA-Pkc53E. HA-Pkc53E was then shuttled via Acc65I/XbaI in likewise opened pUAST-attB vector (Bischof et al., 2007). All cloning steps were sequence verified. Primers used for cloning are included in S2 Table. Transgenic fly lines were then generated with the help of the PhiC31 integrase-based system using 96E as landing site (Bischof et al., 2007).
RT-PCR of Pkc53EΔ28 null mutants
Poly(A)+ RNA was isolated from 50 third instar larvae (Pkc53EΔ28 and y1w67c23) using PolyATract System Kit 1000 (Promega, Mannheim, Germany) according to the manufacturer’s protocol, followed by a DNase I treatment (New England Biolabs GmbH, Frankfurt, Germany, #M0303). Subsequent cDNA synthesis was conducted with qScriber cDNA Synthesis Kit (highQu, Kraichtal, Germany) according to the supplier’s protocol. For amplification, a Pkc53E primer pair overlapping the last three introns was chosen (Pkc53E_RT-PCR UP and Pkc53E_RT-PCR LP). Tubulin 56D primers (Tub56D_229 UP and Tub56D_507 LP) were used as internal controls. For primers, see S2 Table.
Analyses of Drosophila hematopoetic cells and tissues
Determination of sessile larval crystal cells
Larval crystal cells were counted according to Frankenreiter et al., 2021. Briefly, staged wandering third instar larvae of the respective genotype were heated to 60°C for 10-12 min. Pictures of the posterior dorsal side were taken with a Pixera camera (ES120, Optronics, Goleta, USA) mounted to a stereo-microscope (Wild M3Z, Leica, Wetzlar, Germany) with Pixera Viewfinder 2.5. Melanized crystal cells appear as black dots, and were counted in the last two larval segments with ImageJ 1.51 software using Cell Counter tool. 25-74 larvae were scored for the statistical evaluation. For the PMA-feeding experiment, 20-30 developmentally synchronized second instar larvae were selected and grown for 24 h in complete dark at 25°C on fly food with 500 µl of 1 mM Phorbol-12-myristat-13-acetat (PMA, Sigma-Aldrich, St. Louis, USA) added to the surface. Subsequently, wandering third instar larvae were heated and analysed as above.
Visualization and quantification of lamellocytes
The lamellocyte specific reporters atilla-GFP or PPO3::GFP strains were re/combined with Su(H)gwt, Su(H)S269A or Pkc53EΔ28 alleles by genetic means. Late second/early third larval instars were infested by L. boulardi and the number of lamellocytes was determined two days later and compared with those observed in non-infested larvae. Larvae were washed thoroughly in cold PBS and dried with a tissue and teared apart. The hemolymph of 10 larvae each was collected with a 20 μl Microloader tip (Eppendorf, Hamburg, Germany) and placed on a slide with 7 μl of Vectashield mounting medium containing DAPI (BIOZOL, Eching, Germany). GFP-positive cells, i.e. lamellocytes were counted in relation to the total number of DAPI labelled hemocytes with a Zeiss Axioskop and a PlanNeofluar 20x objective. 8-10 independent bleedings were performed each.
Immunostaining and documentation of larval lymph glands
Larval lymph glands were prepared one-two days after infection and treated as described before (Frankenreiter et al., 2021). For comparison, non-infested lymph glands were prepared. Primary antibodies used for staining: mouse anti-Hnt for crystal cells (DSHB 1G9, RRID: AB_528278, 1:20) and guinea pig anti-Pzg as nuclear marker (Kugler and Nagel, 2007; 1:500). GFP signals were monitored directly. Secondary fluorescent antibodies were from Jackson Immuno-Research Laboratories (obtained from Dianova, Hamburg Germany, 1:250 each). Mounted tissue was documented with a Zeiss Axioskop coupled with a BioRad MRC1024 confocal microscope using LaserSharp software 2000. For statistical evaluation at least 12 primary lobes were documented and statistically analyzed by using Image J software (Schindelin et al., 2012). Indices represent the number of cells in relation to the size/area of the tissue (in pixel) x 10000.
Determination of kinases and kinase assays
Screening of protein kinase candidates in silico and in vitro
To search for potential candidates in silico, GPS3.0 software was used at the lowest threshold levels, including the 40 kinases with the highest difference between score and cut-off value (Xue et al., 2011). The corresponding Drosophila kinases were determined with the help of flybase according to (Morrison et al., 2000). For the in vitro screen, a 0.5kb cDNA fragment (741-1242) encoding the Su(H) beta-trefoil domain (codons 247-414), was PCR-amplified and cloned via BamHI/ EcoRI into pGEX-2T vector (Smith and Johnson, 1988) for bacterial expression and purification of the BTD-GST fusion protein. Primers used for cloning are included in S2 Table. ProQinase GmbH (Freiburg, Germany) provided the ‘KinaseFinder assay service’. Briefly, BTD-GST and 33P-ATP served as substrates for 245 human Ser/Thr kinases in multi-well plates, analysed in a microplate scintillation reader. (A) Activity of each kinase was determined, (B) corrected for substrate background activity, and (C) auto-phosphorylation (kinase activity without substrate). A ratio value between phosphorylation of BTD-Su(H) and kinase auto-phosphorylation >1 (A-B/C) was considered as significant.
In vitro ADP-GloTM kinase assay
Drosophila pBT-3xHA-Pkc53E was mutated to generate the pseudo-activated form Pkc53EEDDD (A34E/T508D/T650D/S669D) stepwise by site directed mutagenesis using the Q5® Site directed Mutagenesis Kit (New England Biolabs, Frankfurt, Germany). Primers used for mutagenesis are included in S2 Table. Pkc53E as well as PKC53EEDDD were then shuttled into a modified pMAL vector (Riggs, 1994) where additional restriction sites for Acc65I, SacII and XhoI had been included in the multiple cloning site via primer annealing. The MBP-Pkc53E and MBP-Pkc53EEDDD fusion proteins were bacterially expressed and purified with Amylose resin (New England Biolabs, Frankfurt, Germany). Additionally, activated human kinase PKCα was obtained for a positive control (ProQinase, Freiburg, Germany). The PKCα pseudo-substrate PS (RFARLGSLRQKNV) (Kochs et al., 1993), the wild type Su(H) peptide Swt (ALFNRLRSQTVSTRY) and the phospho-deficient peptide SSA (ALFNRLRAQTVSTRY) were obtained (peptides & elephants, Henningsdorf, Germany).
To test kinase activity, the ADP-GloTM Kinase Assay system (Promega, Madison, USA) was used. Kinase assay reactions were performed in 96 well plates in a volume of 25 μl in the dark. Each reaction contained 100 μM of a kinase substrate peptide, 150 ng purified kinase and 500 μM ultra-pure ATP. 150 nM Phorbol 12-myristate 13-acetate (PMA) (Sigma-Aldrich, St. Louis, USA) was added to some reactions. The mixture was filled up with Kinase reaction buffer (40 mM Tris-HCl, 20 mM MgCl2, 0.1 mg/ml BSA, pH 7.4) and incubated at room temperature for 1 h in the dark. 25 μl ADP-Glo Reagent were added and incubated for 40 min to remove residual ATP. 50 μl of Kinase Detection Reagent was applied to convert ADP to ATP. The luminescent signal was measured after 45 min using GloMax® Discover Microplate Reader (Promega, Madison, USA), kindly provided by the Department of Zoology (190z), University of Hohenheim.
NanoLC-ESI-MS/MS analysis of Su(H) peptides
Nano-LC-ESI-MS/MS experiments were performed by the Mass Spectrometry Unit at the Core Facility Hohenheim (640) on an Ultimate 2000 RSLCnano system coupled to a Nanospray Flex Ion Source and a Q-Exactive HF-X mass spectrometer (Thermo Fisher Scientific, Waltham, USA). Peptides were separated with LTQ-Orbitrap XL coupled to a nano-HPLC operated under the control of XCalibur 126.96.36.199 software (Thermo Fisher Scientific, Waltham, USA). For all measurements using the Orbitrap detector, internal calibration was as described before (Olsen et al., 2005). MS/MS spectra were analyzed using Proteome Discoverer 2.2 (ThermoFisher Scientific, Waltham, United States), verified by manual inspection of the MS/MS spectra (Voolstra et al., 2010).
Generation of an ⍺-pS269 antiserum
Rabbit polyclonal p-S269 Su(H) antiserum was generated by DAVIDS Biotechnology GmbH (Regensburg, Germany) using the synthetic phospho-peptide NLRLpSQTVSTRYLHVE. Phospho-specific antibodies were enriched in a depletion-step by affinity purification against the non-phosphorylated peptide.
Immunoprecipitation of mCherry and Myc-tagged Su(H)
400 adult heads or 25 larvae of each genotype were homogenized on ice in 220 μl buffer 1 [150 mM NaCl, 1% Triton X-100, 50 mM Tris-HCl pH 7.5, 0.1% SDS, supplemented with protease inhibitor cocktail (Roche, Basel, Switzerland)] and incubated for 15 min. After a short spin, 20 μl of the supernatant was set aside as input fraction (‘protein extract’). The residual supernatant was diluted with 300 μl wash buffer I (see buffer I without Triton X-100). 15 μl of equilibrated magnetic RFP-Trap Agarose beads (ChromoTek, Planegg, Germany) were added, incubated for 1 h at 8°C and washed three times with wash buffer I. mCherry-trapped proteins were resolved on SDS-PAGE; Western blots were probed with rabbit anti-mCherry (GeneTex, Irvine, USA, 1:1000, #GTX128508) and rat anti-HA (Roche, Basel, Switzerland, 1:500, #11867423001). Goat secondary antibodies coupled with alkaline phosphatase (Jackson Immuno Research Laboratories, 1:1000) were used for detection.
HeLa cell culture experiments and Luciferase assays
Generation of HSV-TK 2xMyc-Su(H)-VP16
Two myc-tags were added to Su(H) cDNA in pBT (Maier et al., 2011) by insertion of the two annealed oligonucleotides Myc-Tag UP and Myc-tag LP into the EcoRI site (for primers, see S2 Table). The construct was subsequently shuttled via EcoRI/XhoI into pCDNA3.1 (Invitrogen, Thermo Fisher Scientific, Waltham, USA). A VP16 activator domain was then cloned in frame at the C-terminus of 2xMyc-Su(H) by replacing the 829 bp BspEI/ApaI fragment of pCDNA3 2xMyc-Su(H) with a respective1060 bp fragment of the pUAST Su(H)-VP16 construct (Cooper et al., 2000). Then the CMV Promotor of pCDNA3 2xMyc-Su(H)-VP16 was replaced by the HSV-TK Promotor of the pRL TK Vector (Promega, Madison, USA). To this end, the 1023 bp BglII/NheI fragment from pRL TK was cloned into the BglII/SpeI opened pCDNA3 2xMyc-Su(H)VP16 construct.
Transfection of HeLa cells and reporter assay
RBPjKOHeLa cells were cultivated and transfected as described (Wolf et al., 2019). The following constructs were used: pGL3 NRE-reporter (Bray et al., 2005), pCDNA3 HSV-TK 2xMyc Su(H)VP16 and pRL TK (Promega, Madison, USA). For the Luciferase assay, 1 x 105 HeLa RBPjKO cells were seeded in each well of a 12-well cell culture plate. After 24 h the cells were transfected with 500 ng pGL3 NRE-reporter, 460 ng pCDNA3 HSV-TK 2xMyc Su(H)VP16 and 40 ng pRL-TK. 4 h after the transfection the cells were treated with 162 nM PMA (Sigma-Aldrich, St. Louis, USA), 162 nM PMA plus 21,4 nM stauporine (STAU, Sigma-Aldrich, St. Louis, USA) or 21,4 nM STAU alone. Control cells were treated with the same volume of DMSO present in the other treatments. 14 hours later, cells were washed twice in PBS pH 7.4 and lysed in 75 µl 1x Passive Lysis Buffer (Promega, Madison, USA). The Dual-Luciferase® Reporter Assay (Promega, Madison, USA) was performed according to the manufacturer’s instructions.
Statistical analysis and documentation of data
Statistical significance of collected data was determined by a two-tailed analysis of variance (ANOVA) approach for multiple comparisons according to Dunnett’s Test and Tukey-Kramer’s Honestly Significance Difference with p-values ***, p≤0.001; **, p≤0.01; *, p≤0.05; not significant, p>0.5. Boxplots were created with BoxPlotR software (Spitzer et al. 2014). Pictures were assembled using ImageJ, PhotoPaint, CorelDraw and BoxPlotR software.
All data are contained within the manuscript.
We are deeply grateful to Benedikt Häußling and Johannes Stöckl (University of Bayreuth, Germany) for sending us all wasp species used in this study and for giving LF and SD a basic course in the handling of wasps. We thank Michèle Crozatier (Toulouse, France), David Hipfner (Montréal, Canada), Bruno Lemaitre (Lausanne, Switzerland), Sarah Bray (Cambridge, UK), Dieter Maier (Hohenheim, Germany), the Bloomington Drosophila Stock Center (BDSC, NIH P40OD018537) and the Vienna Drosophila Stock Center (VDRC) for numerous fly stocks. We acknowledge the Drosophila Genomics Resource Center (DGRC, NIH 2P40OD010949) for sending the Pkc53E cDNA, and the Developmental Studies Hybridoma Bank (DSHB), created by the NICHD of the NIH and maintained at the University of Iowa, Department of Biology, Iowa City, IA 52242 for providing the Hnt (1G9) antibody, developed by HD Lipshitz. We very much acknowledge Franz Oswald (Ulm, Germany) and Tilman Borggrefe (Gießen, Germany) for the RBPjKO HeLa cell line. We are indebted to Lisa Lermer for her help in tissue preparations and screening of larvae and Janika Scharpf for the purification of the Pkc53E proteins. We are grateful to Armin Huber, Department of Biochemistry, for the use of the Apotome microscope, to Axel Schweickert, Department of Zoology, for use of the GloMax® Discover Microplate Reader and to Jens Pfannstiel at the Mass Spectrometry Unit (Core Facility of the University of Hohenheim) for the MS/MS and ESI spectra. We thank Dieter Maier for helpful comments on the manuscript.
This work was supported by a grant of the German Science Foundation DFG to ACN (NA 427/5-1) and by the University of Hohenheim. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
Sebastian Deichsel, Data curation, Formal analysis, Methodology, Investigation, Resources, Writing—review and editing, Validation, Visualization; Lisa Frankenreiter, Formal analysis, Methodology, Investigation, Resources, Writing—review and editing, Validation; Johannes Fechner, Formal analysis, Methodology, Investigation, Resources, Writing—review and editing, Validation; Bernd M. Gahr, Data curation, Methodology, Investigation, Resources, Writing—review and editing, Visualization; Mirjam Zimmermann, Formal analysis, Investigation, Resources, Writing—review and editing; Helena Mastel, Investigation, Writing—review and editing; Irina Preis, Investigation, Writing—review and editing; Anette Preiss, Formal analysis, Writing—original draft, Writing—review and editing, Visualization; Anja C. Nagel, Conceptualization, Data curation, Formal analysis, Methodology, Writing— original draft, Writing—review and editing, Validation, Visualization, Supervision, Funding acquisition, Project administration.
Figure 2 - figure supplement 1
The α-pS269 antiserum detects the phospho-mimetic Su(H) variant in vitro
Figure 3D – figure supplement 1
MS/MS spectra of the phosphorylated Su(H) peptide
Figure 4 – figure supplement 1
Conservation of Pkc53E and generation of an activated Pkc53EEDDD isoform
Figure 6 – figure supplement 1
The Pkc53EΔ28 allele is a null mutant
Figure 3 – supplement Table 1
List of kinases predicted to recognize S269 in Su(H) as substrate in silico
Figure 3 – supplement Table 2
List of kinases accepting the BTD domain of Su(H) as substrate in vitro
Figure 3 – supplement Table 3
Fly strains used for the larval crystal cell screen
Figure 3 – supplement Table 4
Larval crystal cell screen
Supplemental Tables, Materials and Methods
S1 Table Key resources
S2 Table Oligonucleotides
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