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Seraing-le-chateau, Belgium

Eurogentec is an international biotechnology supplier, based in Belgium, that specializes in genomics and proteomics kits and reagents as well as cGMP biologics. The company was founded in 1985 as a spin-off from the University of Liège. Eurogentec's contract manufacturing organization facilities are licensed by the Belgian Ministry of Health to produce clinical trial and commercial biopharmaceutical material and also licensed by the US FDA to manufacture a commercial recombinant protein product for the US market. Eurogentec operates two manufacturing facilities in Belgium that provide custom biologics and oligonucleotide-based components for diagnostic and therapeutic applications. Wikipedia.


Mercier F.,University of Liege | Paris J.,University of Liege | Kaisin G.,University of Liege | Thonon D.,University of Liege | And 4 more authors.
Bioconjugate Chemistry | Year: 2011

The alkyne-azide Cu(I)-catalyzed Huisgen cycloaddition, a click-type reaction, was used to label a double-stranded oligonucleotide (siRNA) with fluorine-18. An alkyne solid support CPG for the preparation of monostranded oligonucleotides functionalized with alkyne has been developed. Two complementary azide labeling agents (1-(azidomethyl)-4-[18F] fluorobenzene) and 1-azido-4-(3-[18F]fluoropropoxy)benzene have been produced with 41% and 35% radiochemical yields (decay-corrected), respectively. After annealing with the complementary strand, the siRNA was directly labeled by click chemistry with [18F]fluoroazide to produce the [ 18F]-radiolabeled siRNA with excellent radiochemical yield and purity. © 2010 American Chemical Society. Source


An apparatus and a method for obtaining a (poly)nucleotide sequence of interest include steps of cultivating hosts cells to produce a nucleotide sequence of interest and harvesting these cells, introducing these cells in a passageway and disintegrating them in a continuous process. In the continuous process, performing in the passageway a precipitation of contaminants by a mixing of the disintegrated cells with a solution containing one or more salt(s) and obtaining a mixture and allowing a precipitate to separate from the solution of this mixture, preferably to float and/or to sediment from the solution of this mixture for 1-48 hours and pumping out a soluble material from this solution, while excluding recovering the precipitate.


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All animal experiments were performed in accordance with the European Community and local ethics committee guidelines. Xenopus laevis were purchased (Nasco) and maintained in our animal facility. Ambystoma mexicanum (axolotls) were bred and maintained in our facility, where they were kept at 18 °C in Dresden tap water and fed daily with artemia or fish pellets. Five-to-six centimetre (snout to tail tip) axolotls were used for all the experiments. Animals were anaesthetized for all the surgical process as previously described20. Labelling of connective tissue was achieved by transplanting lateral plate mesoderm from GFP transgenic embryos to normal host embryos as previously described20. Total RNA was isolated from day-1, day-3 and day-5 limb and tail blastemas with TRIzol reagent (Invitrogen) according to the manufacturer’s manual. Total RNA from mature (not regenerating) limb tissues was isolated using the same procedure as blastema samples. Blastema RNAs from the different time points were equivalently pooled as limb–blastema total RNA or tail–blastema total RNA, respectively. mRNA was purified from limb–blastema or tail–blastema total RNA with the Poly (A) Quick mRNA isolation kit (Stratagene). Xenopus oocyte preparation and microinjection were performed essentially as previously described6, 7, 21. Briefly, mature oocytes were defolliculated with collagenase (Sigma). Purified mRNA (5.0 ng) was injected into the selected healthy oocytes after the defolliculation. Eight injected oocytes were cultured together in one well of a 96-well plate (Nunc) for 48 h and the supernatants were harvested for myotube assay (see later). Clones (110,592) from a 6-day tail blastema library were arrayed into 288 × 384-well plates22. To prepare the ‘pools’, all the saturated bacterial cultures on one 384-well plate were pooled in one conical tube, and 288 pools were prepared from the library in total. To prepare the ‘superpools’, 24 pools were combined together in one conical tube, and 12 superpools were prepared from the pools in total (Extended Data Fig. 1a–c). To obtain superpool plasmid, 500 μl of superpool bacteria were cultured in 50 ml LB medium (Extended Data Fig. 1d). To avoid losing low frequency clones in the superpools, the optical density (OD) of each culture was controlled and the cultures were harvested around OD 0.6. Superpool plasmids were purified with QIAGEN Plasmid Midi Kit (QIAGEN) according to the manufacturer’s manual. To reconstitute the whole library, 5 μg of each superpool’s plasmids were pooled in one tube before transfection. HEK293FT cells (Invitrogen) were maintained with the standard protocol from Invitrogen. To obtain the superpool supernatants, 8.0 × 105 of HEK293 cells were plated on one well of a 6-well plate (Nunc) and 1 μg of each individual superpool plasmids were transfected into HEK293 cells with Fugene 6 (Roche; Extended Data Fig. 1e) according to the manufacturer’s manual. For the first 24 h, the transfected HEK293 cells were kept in the 10% fetal calf serum (FCS) medium. Then the cells were rinsed with FreeStyle 293 expression medum (Gibco) that is a serum-free medium and cultured in the medium at 72 h after transfection. Individually harvested supernatants were concentrated approximately tenfold with a Vivaspin 10,000 MWCO (Sartorius). These concentrated supernatants were tested on A1 myotubes (Extended Data Fig. 1f). It should be noted that given the injury-specific extracellular activity of AxMLP, we infer that the Xenopus oocyte and HEK293 cell systems are likely to be in ‘wound-epithelium-like’ signalling states that permit at least some extracellular release of AxMLP, and that the 6-day regenerating tail blastema cDNA had a sufficient number of AxMLP clones for detection in the expression cloning system. We only detected 1 AxMLP clone among 100,000 clones, and this may reflect the levels of mRNA present at later regeneration time points. Maintenance of A1 myoblasts, myotube differentiation from A1 myoblasts, myotube purification and subsequent myotube assays were performed essentially as described previously8, 23, 24. Briefly, concentrated supernatants were individually added into myotube culture medium in a 96-well plate and incubated for 5 days, and BrdU (Sigma; final 10 μg ml−1) was added to the culture media for 18 h before the fixation with 1.5% PFA (Sigma)/PBS. Fixed myotubes were stained with anti-MHC and anti-BrdU antibodies (mouse monoclonal, 4a1025 and Bu20a) conjugated to FITC or rhodamine with DyLight Antibody Labelling Kit (Thermo Scientific) according to the manufacturer’s manual. For the quantification of BrdU incorporation activity, the total number of myotubes and BrdU+ myotubes were counted by hand under the microscope (Zeiss Axioplan 2). This biological evaluation of the BrdU-incorporation activity on myotubes is called the myotube assay. The high-serum (15% FCS) condition was used as a positive control for myotube assays and the low-serum (0.5% FCS) or serum-free condition was used as a negative control. For the second screen from superpool number 9, the clones from the 24 384-well plates contained in superpool 9 were divided into smaller sub-pools according to a two-dimensional matrix (Extended Data Fig. 2a). For example, sub-pool A contained pools from plates 193–198 and sub-pool 1 contained pools from plates 193, 199, 205, 211. These sub-pools were cultured to OD 0.6 before plasmid preparation. For the third screen from pool number 212, 384 single clones were arrayed by 96-pin plastic replicators (Genetix) on 96-well plates (SARSTEDT) filling 150 μl LB medium per well (Extended Data Fig. 2b; groups A–D). Individual clones on the 96-well plates were statically cultured until they were saturated and 24 clones were pooled together (Extended Data Fig. 2b; sub-pools A1–D4). Plasmids from each pool were purified with QIAprep spin miniprep kit (QIAGEN). For the fourth screen from A1, 24 clones were individually cultured in LB medium and the plasmids were purified with QIAprep spin miniprep kit (QIAGEN). To construct sub-pools, 1 μg of the plasmid from each single clone was pooled according to the diagram in Extended Data Fig. 2c. The process from the transfection into HEK293 cells to myotube assay in the second to the fourth screen was the same as the first screen. To validate transfection efficiency during whole expression cloning, 50 pg of secreted alkaline phosphatase (SEAP)-pCMV-SPORT6 plasmid was co-transfected with the samples as a spike and a portion of the supernatants was assayed by Great EscAPe SEAP Chemiluminescence Kit 2.0 (Clontech). The luminescence of the supernatants was measured by GENiosPro Microplate Reader (TECAN). We confirmed that there was no significant difference of transfection efficiency during the expression cloning (data not shown). There were no cell line misidentification and cross-contamination in the experiments. We used a single mammalian cell line (HEK293FT cells) provided by Invitrogen and single amphibian cell line (newt A1 cells) in the experiments. These two cell lines have totally different morphologies and are cultured under mutually incompatible culture conditions. The growth of both cells were carefully monitored during the experiments and cells samples were constantly stained with Hoechst 33342 (Sigma, final 0.5 μg ml−1) to test mycoplasma contamination. Human and mouse MLP cDNA clones were purchased from OriGene Technologies (clone ID: human, SC112373; mouse, MC208965). Zebrafish and Xenopus mlp cDNA clones were purchased from Source BioScience (clone ID: zebrafish, 6795591; Xenopus, 8330180). All oligonucleotide sequences and the restriction enzyme sites using for cloning are shown in Supplementary Table 2. Since the backbone vector of the cDNA library is pCMV-SPORT6, we subcloned following genes into pCMV-SPORT6 vector (Invitrogen) or pCMV-SPORT6-3C-His vector. PCR-amplified fragments with the oligonucleotides numbers 1 and 2 from pSEAP2-Basic (Clontech) were subcloned into pCMV-SPORT6. The oligonucleotides numbers 3 and 4 were attached to pCMV-SPORT6 to generate a backbone vector, pCMV-SPORT6-3C-His (Extended Data Fig. 3c, bottom left). The AxMlp open reading frame (ORF) was amplified by PCR with the oligonucleotides numbers 5 and 6 from the original AxMlp clone (BL212a101) and subcloned in the pCMV-SPORT6-3C-His vector (Extended Data Fig. 3c, top left). N-terminal deletion AxMlp was amplified by PCR with the oligonucleotides numbers 7 and 8 from the original AxMlp clone (BL212a101) and subcloned in the pCMV-SPORT6-3C-His vector (Extended Data Fig. 6a, bottom). Human, mouse, zebrafish and Xenopus MLP ORFs were amplified from purchased cDNA clones with specific primers (for human, oligonucleotides numbers 9, 10; mouse, oligonucleotides numbers 9, 11; zebrafish, oligonucleotides numbers 12, 13; Xenopus, oligonucleotides numbers 14, 15, respectively), and were subcloned in the pCMV-SPORT6-3C-His vector. The oligonucleotides numbers 16 and 17 were inserted into to the pEGFP-N1 plasmid (Clontech) to generate a backbone vector, pEGFP-N1-3C (Extended Data Fig. 3c, bottom right). The AxMlp ORF was amplified by PCR with the oligonucleotides numbers 18 and 19 from the original AxMlp clone (BL212a101) and subcloned into pEGFP-N1-3C (Extended Data Fig. 3c, top right). Newt Mlp ORF was amplified by PCR from newt limb blastema cDNA with the oligonucleotides numbers 28 and 29 and the PCR fragments were subcloned in the pCMV-SPORT6-3C-His vector. These constructs were confirmed by sequencing. For measuring the GFP intensity of supernatants, 8.0 × 105 of HEK293 cells were plated on 6-well plates and 1 μg of AxMLP-3C-pEGFP-N1 plasmid or 1 μg of empty-pEGFP-N1 plasmid were transfected into HEK293 cells. The supernatants were harvested at 24 h post-transfection (hpt), 48 hpt and 72 hpt and concentrated with Vivaspin 10,000 MWCO (Sartorius) individually. The fluorescence intensity was measured using a GENiosPro Microplate Reader (TECAN). To determine the percentage of GFP+ cells in the culture, the transfected cells were detached with Trypsin-EDTA (Gibco, final 0.05%)/PBS from the well, then spread on improved Neubauer chamber. The number of GFP+ cells and total cells in the grids were counted by hand and the percentage was calculated. Time-lapse imaging was performed under Axiovert 200M (Zeiss) with humidity, temperature and CO control chamber. Images were taken every 30 min from 5 to 72 hpt. For the antibody-based blocking assay in vitro, 1 μg of AxMlp-3C-His-pCMV-SPORT6 plasmid or empty-pCMV-SPORT6-3C-His plasmid was transfected into HEK293 cells with Fugene 6 (Roche). The supernatants were harvested at 72 hpt and concentrated. Ten micrograms of AxMLP-3C-His protein (22.7 kDa) were treated with 70 μg or 350 μg of anti-AxMLP polyclonal antibody (see later) or anti-GFP polyclonal antibody (MPI-CBG antibody facility) as a negative control, respectively, at room temperature for 2 h. These antibody-treated supernatants were used for the myotube assay. For the in vivo antibody blocking assay, anti-full-length AxMLP polyclonal antibody (see later), anti-GFP polyclonal antibody (MPI-CBG antibody facility) or PBS as a negative control were injected into mature (not regenerating) tail as the first injection (3 h before amputation) and injected into the blastema as the second injection (12 h post-amputation) and as the third injection (1 day post-amputation) (Extended Data Fig. 9a). These samples were co-injected with tetramethylrhodamine dextran MW 70,000 (Molecular Probes; final 2.5 mg ml−1) as a tracer. The injection efficiency was confirmed based on the intensity of the rhodamine under the fluorescence dissecting microscope (SZX 16, OLYMPUS). No animals were excluded from the analysis. In each injection 500 ng, then, in total 1.5 μg antibody or equivalent volume of PBS were injected. Injected animals were kept in clean tap water for 3 days at room temperature. The animals were injected intraperitoneally with 30 μl of 2.5 mg ml−1 BrdU (Sigma) 4 h before collecting the tails. The injected blastemas were fixed, embedded, cryosectioned and immunostained as described later. For the imaging, the tiled images of the entire cross-section of the tails taken on a Zeiss Observer.Z1 (Zeiss) were then stitched by Axiovision software or Zen 2 (Zeiss). For the quantification at least a total of 1,000 cells per one animal were counted from four different animals in each condition (PBS, anti-GFP antibody or anti-AxMLP injection, respectively), and the marker-positive nuclei (BrdU+, PAX7+, MEF2+ or Hoechst+) on the sections were counted by hand. The cells in spinal cord, epidermis and cartilage/notochord were separately counted based on morphology. The label “Other tissues” in Fig. 2d, contained mainly mesenchymal cells and endothelial cells and was calculated by the subtraction from the total number to the number of all the other specific cell types. For His-tagged AxMLP purification, AxMLP-3C-His-pCMV-SPORT6 plasmid was transfected into HEK293 cells and the supernatant was harvested at 72 hpt. His-tagged protein in the supernatant was purified in native conditions on a 1 ml HisTrap HP column (GE Healthcare) using FreeStyle 293 expression medium including 500 mM imidazole step elution. The eluate (purified AxMLP) and depleted media (flow-through) were concentrated with Vivaspin 10,000 MWCO (sartorius) 40 fold and the final concentration of purified AxMLP was 1.31 μg μl−1. Both concentrated eluate (purified AxMLP) and flow-through fractions were dialysed with Spectra/Por Dialysis Membrane MWCO 6-8000 (Spectrum Laboratories) in AMEM (MEM medium (Gibco) diluted 25% with distilled water) for biological assays. The fractions from the purification were tested by silver staining and western blotting (Extended Data Fig. 3g, h). The washing fraction was concentrated about tenfold to load the same volume as other fractions on 4–20% gradient SDS–PAGE gels (anamed Elektrophorese). Western blotting and silver staining were performed with a standard protocol. Briefly, the fractions were treated with 2× Sample Buffer including dithiothreitol (DTT; Sigma, final 0.2 M) and boiled at 95 °C for 10 min. The proteins were blotted on PROTRAN nitrocellulose transfer membrane (Whatman) by TE 77 Semi-Dry Transfer Unit (Amersham). The membrane was blocked with 5% skim milk. Primary antibodies used: mouse anti-His (QIAGEN, 1/5,000), mouse anti-α-tubulin (MPI-CBG antibody facility, DM1A 1/5,000), rabbit anti-AxMLP-full length (1/2,500), rabbit anti-AxMLP-C terminus (1/2,500). Secondary antibodies used: goat anti mouse-HRP (Jackson ImmunoResearch Laboratories, 1/5,000), goat anti rabbit-HRP (Jackson ImmunoResearch Laboratories, 1/5,000). Cell lysates for western blotting were obtained by directly adding 2× Sample Buffer on top of the cultured cells and were boiled at 95 °C for 10 min. For the preparation of anti-full-length AxMLP polyclonal antibody, a glutathione S-transferase (GST) fusion protein with full-length amino acids of AxMLP was expressed in bacteria and purified by standard methods on GS-trap, glutathione sepharose (GE Healthcare). Purified GST fusion protein as an antigen was used to immunize rabbits (Charles River). Anti-serum was affinity purified using maltose-binding protein fused with full-length AxMLP conjugated to NHS-Sepharose resin (GE Healthcare). To raise C-terminal AxMLP polyclonal antibody, keyhole limpet haemocyanin (KLH)-tagged peptides, PPVEPQVEEVAAPAP, was used to immunize rabbits and the affinity-purified polyclonal antibody was provided (Eurogentec). Both anti-full-length and anti-C-terminal AxMLP polyclonal antibodies were tested on the cell lysate from AxMLP-transfected HEK293 cells (Extended Data Fig. 3f). Limb blastema and tail blastema preparations for sectioning were produced essentially as previously described25. Briefly, limb blastemas amputated at the wrist level were collected from the level of the shoulder, and tail blastemas amputated at the 12th myotome from the cloaca were collected at the 10th myotome of the regenerating tail. These limb and tail blastemas were immunostained as previously described15, 25, 26. Briefly, the samples were fixed with MEMFA fixative at 4 °C overnight, and were rinsed with PBS several times. The buffer was replaced from PBS to 10%, 20% and 30% sucrose (Sigma)/PBS, then the samples were embedded in Tissue-Tek O.C.T. Compound (Sakura) for cryosection and the tissues were sectioned at 10-μm thickness with Microm HM 560 cryostat (Thermo). Primary antibodies used: mouse anti-BrdU (MPI-CBG antibody facility, Bu20a 1/400), rabbit anti-BrdU (antibodies-online, 1/600), mouse anti-PAX7 (MPI-CBG antibody facility, PAX7 1/450), rabbit anti-MEF2 (Santa Cruz, 1/200), rabbit anti-AxMLP-C terminus (1/200), rabbit anti-GFP (Rockland, 1/400), rabbit anti-FITC (Invitrogen, 1/400), mouse anti-FITC (Jackson ImmunoResearch Laboratories, 1/400), rat anti-MBP (GeneTex, 1/200). The following appropriate fluorophore-conjugated secondary antibodies were used (all in 1/200 dilution): donkey anti-mouse Alexa Fluor (AF) 647 (Molecular Probes), goat anti-mouse AF 647 (Jackson ImmunoResearch Laboratories), donkey anti-mouse AF 488 (Molecular Probes), goat anti-rabbit AF 647 (Jackson ImmunoResearch Laboratories), donkey anti-rabbit AF 488 (Molecular Probes), donkey anti-rat AF 488 (Molecular Probes). The cell nuclei were stained with Hoechst 33342 (Sigma, final 0.5 μg ml−1). Imaging for the stained sections was performed with Zeiss Observer.Z1 (Zeiss) controlled by Axiovision software or Zen2 (Zeiss). Total RNA preparation, reverse transcription and qRT–PCR were essentially described in the previous work2. Briefly, three biological replicas were prepared for each time point and they were technically independent in all the processes (tissue collection, RNA preparation, cDNA synthesis and qRT–PCR). Eight to approximately ten limb or tail blastemas from one time point were collected in one tube and homogenized by POLYTRON PT1600E (KINEMATICA). Total RNA was purified with RNeasy Mini or Midi Kit (QIAGEN) according to the manufacturer’s manual. cDNA was synthesized from 300 ng of total RNA using SuperScript III First-Strand Synthesis System (Invitrogen) and qRT–PCR was performed with Power SYBR Green Master Mix (Invitrogen) in total volume of 12 μl with the final primer concentration of 300 nM on the LightCycler 480 (Roche). To obtain the values of fold change for each time point, the relative concentration of the PCR products was calculated by the standard curve method. To obtain the standard curves of the limb time course or the tail time course respectively, the dilution series (1/4, 1/16, 1/64, 1/256 and 1/1,024) were made from the mixture of cDNAs that were equivalently collected from the cDNA samples in all the different time points. These dilution series were used as the template for the PCR and the relative concentrations were calculated by LightCycler 480 Software (Roche) based on the standard curves. The concentration of AxMlp was normalized with that of Rpl4 (large ribosomal protein 4). Primers used for PCR were showed in Supplementary Table 2 (for AxMlp, oligonucleotides numbers 20, 21; for Rpl4, oligonucleotides numbers 22, 23). The raw values of qPCR data are shown in Supplementary Table 1. The dialysed protein samples: purified AxMLP, depleted media (flow-through) (see earlier) or PBS as a negative control were injected into mature (not regenerating) tails with a pressure injector, PV830 Pneumatic Picopump (World Precision Instruments). These protein samples were co-injected with tetramethylrhodamine dextran MW 70,000 (Molecular Probes, final 2.5 mg ml−1) as a tracer. A glass capillary (Harvard Apparatus) for the injection was pulled with P-97 Micropipette Puller (Sutter Instrument) and sharpened manually (external tip diameter: 30 μm). The injection efficiency was confirmed based on the intensity of the rhodamine under the fluorescence dissecting microscope (SZX 16, OLYMPUS). No animals were excluded from the analysis. In total, 270 ng of purified AxMLP or equivalent volume of controls were injected into one side of the tail. Injected animals were kept in clean tap water for 3 days at room temperature. The animals were injected intraperitoneally with 30 μl of 2.5 mg ml−1 BrdU (Sigma) 4 h before collecting the tails (Fig. 2a). The injected part of the tails was identified by rhodamine-positive myotomes and these tails were fixed, embedded, cryosectioned and immunostained as described earlier. For the quantification, the tile images of whole cross-sections of the tails from Zeiss Observer.Z1 (Zeiss) were stitched by Axiovision software (Zeiss). Three sections from five different animals in each condition (PBS, flow-through or purified AxMLP injection, respectively) were taken, and the marker-positive nuclei (BrdU+, PAX7+, MEF2+ or Hoechst+) on the sections were counted by hand. The cells in spinal cord, epidermis and notochord were separately counted based on morphology. The label “Other tissues”, contained mainly mesenchymal cells and endothelial cells, and was calculated by the subtraction from the total number to the number of all the other specific cell types. For the protein injection into the limb, the procedure was essentially as described earlier. Purified AxMLP protein was injected into the mature (not regenerating) right lower limbs at the centre between the elbow and the wrist. The control samples (flow-through fraction or PBS) were injected into the left limbs of the same animal that were injected with purified AxMLP on their right limbs. In total 2.0 μg purified AxMLP or equivalent volume of controls were injected into the limbs. The animals were injected intraperitoneally with 30 μl of 2.5 mg ml−1 BrdU (Sigma) 12 h before collecting the limbs (Extended Data Fig. 4e). For the quantification, at least a total of 1,000 cells per one animal were counted from four different animals in each condition (PBS, flow-through or purified AxMLP injection, respectively), and the marker-positive nuclei (BrdU+, PAX7+, MEF2+, MBP + or Hoechst+) on the sections were counted by hand. The cells in epidermis and bone/perichondrium were separately counted with their morphology. The label “Other tissues”, contained mainly mesenchymal cells and endothelial cells, and was calculated by the subtraction from the total number to the number of all the other specific cell types. For the acceleration experiment, purified AxMLP, flow-through or PBS as a negative control were injected into mature (not regenerating) tail as the first (3 days before amputation) injection and as the second (1 day before amputation) injection and injected into the blastema as the third (2 days post-amputation) injection (Extended Data Fig. 10a). These samples were co-injected with tetramethylrhodamine dextran MW 70,000 (Molecular Probes, final 2.5 mg ml−1) as a tracer. The injection efficiency was confirmed based on the intensity of the rhodamine under the fluorescence dissecting microscope (SZX 16, OLYMPUS). No animals were excluded from the analysis. The samples were injected into both side of the tail and in each injection, 600 ng, then, in total 1.8 μg protein or equivalent volume of controls were injected. Injected animals were kept in clean tap water for 4 days at room temperature. The length of the blastema was measured from the amputation plane to the tip at the spinal cord level at 4 dpa based on the stereoscope images (SZX 16, OLYMPUS). A1 myoblasts were transfected with original clone BL212a101, AxMLP-3C-His, ΔN-AxMLP-3C-His or empty pCMV-SPORT6-3C-His plasmids and co-transfected with AxMLP-specific morpholinos (Gene Tools; Supplementary Table 2: oligonucleotides numbers 24, 26) or control morpholinos (Gene Tools; Supplementary Table 2: oligonucleotides numbers 25, 27) using Microporator (Digital Bio) according to the manufacturer’s manual with some modifications. All morpholinos were modified with FITC at the 3′ end. A1 myoblasts were re-suspended in 1× Steinberg solution at a density of 5.0 × 106 cells per ml followed by incubation of 10 μl cell suspension with 0.5 μg of plasmid and 1 μl of the morpholino (final 100 μM in the incubation). Electroporation was performed at 1,000 V, 35 mS pulse length and 3 pulses and the electroporated cells were spread in 10% FCS AMEM media24 on a 24-well plate (Nunc), immediately after the electroporation. The culture medium was replaced by new media at 24 h post-electroporation and the cells were kept in culture at 72 h post-electroporation. The electroporated cells were fixed with 1.5% PFA/PBS, and the cell lysates were prepared for western blotting. Primary antibodies used for immunostaining: mouse anti-His (QIAGEN, 1/200), mouse anti-FITC (Jackson ImmunoResearch Laboratories, 1/400), rabbit anti-FITC (Invitrogen, 1/400), rabbit anti-AxMLP-full length (1/1,000). Secondary antibodies used for immunostaining (all in 1/250 dilution): goat anti-mouse Cy3 (Jackson ImmunoResearch Laboratories), goat anti-mouse AF488 (Jackson ImmunoResearch Laboratories), donkey anti-rabbit AF 488 (Molecular Probes), goat anti-rabbit Cy3 (Jackson ImmunoResearch Laboratories). Images of the stained cells were taken with Zeiss Observer.Z1 (Zeiss) controlled by Axiovision software (Zeiss). Electroporation to the spinal cord was performed as previously described with some modifications27. To deliver morpholino into the spinal cord and both sides of the tail epidermis, the tail required electroporation twice with NEPA 21 electroporator (Nepa Gene). The first electroporation was for the spinal cord and one side (left) of the epidermis, and the second electroporation was for the other side (right) of the epidermis. 1.5 μl of morpholino (1.0 mM) was loaded onto a small piece of tissue paper on the left side of the epidermis. Approximately 3 μl of morpholino (1.0 mM) was injected into the spinal cord and immediately electroporated (first electroporation). Sequentially, 1.5 μl of morpholino (1.0 mM) was loaded onto a small piece of tissue paper on the right side of the epidermis and electroporated (second electroporation). The first electroporation conditions: poring pulse, 70 V, 5.0 mS pulse length and 1 pulse; transfer pulse, 55 V, 55 mS pulse length, 5 pulses and 15% decay. The second electroporation conditions: poring pulse, 70 V, 10 mS pulse length and 1 pulse; transfer pulse, 30 V, 30 mS pulse length, 7 pulses and 5% decay. FITC dextran MW 70,000 (Molecular Probes, final 5 mg ml−1) was used as a negative control, since morpholinos were labelled with FITC. The electroporation efficiency in the spinal cord and epidermis was examined based on the intensity of the FITC under the fluorescence dissecting microscope (SZX 16, OLYMPUS). The animals with low FITC intensity were excluded from the next step of the experiments. Three days post-electroporation, the tails were amputated at the level of the maximal morpholino electroporated part. One day post-amputation, a total of 360 ng (180 ng for the spinal cord and 180 ng for blastema) of purified AxMLP or equivalent volume of control flow-through fraction was injected into the spinal cord and the blastema to rescue the morpholino effect. The length of the blastema was measured from the amputation plane to the tip at the spinal cord level on 1, 3, 6, 10 and 14 dpa based on the stereoscope images (SZX 16, OLYMPUS). To detect BrdU incorporation, the animals were injected intraperitoneally with 30 μl of 2.5 mg ml−1 BrdU (Sigma) 4 h before collecting the tails at 3 dpa. Fixation, embedding, cryosection, staining and imaging were described earlier. For the quantification, 3 cross-sections of the blastema (200–300 μm posterior to the amputation plane) from four different animals in each condition (FITC/flow-through, FITC/purified AxMLP, control morpholino/flow-through, control morpholino/purified AxMLP, AxMLP-specific morpholino/flow-through, AxMLP-specific morpholino/purified AxMLP, respectively) were taken, and the marker-positive nuclei (BrdU+, PAX7+, MEF2+ or Hoechst+) on the sections were counted by hand. Red-spotted newts, Notophthalmus viridescens, were supplied by Charles D. Sullivan Co. Animals were anaesthetized in 0.1% ethyl 3-aminobenzoate methanesulfonate (Sigma) for 15 min. Forelimbs were amputated above the elbow, and the bone and soft tissue were trimmed to produce a flat amputation surface. Animals were left to recover overnight in an aqueous solution of 0.5% sulfamerazine (Sigma). At specified time points, the uninjured or regenerating limbs were collected. All surgical procedures were performed according to the European Community and local ethics committee guidelines. The general condition in the newt experiments: 2 μl of 5 mg ml−1 purified AxMLP protein or equivalent volume of flow-through (AxMLP depleted fraction) was injected into the newt limbs. For EdU labelling, animals were injected intraperitoneally with 50–100 μl of 1 mg ml−1 EdU. To investigate the effect of AxMLP on intact newt limbs, purified AxMLP or flow-through was injected into the uninjured limb twice at day 1 and day 3. EdU was administered daily from day 1 to day 5 (Fig. 3a, top). To investigate the effect of AxMLP on regenerating limbs, purified AxMLP or flow-through was injected into the regenerating limbs at 7 and 10 dpa (Fig. 3b, top). EdU was administered daily from 8 to 13 dpa. For labelling myofibre progeny, a H2B-YFP reporter construct was introduced into myofibres before amputation as previously described15 (Fig. 3c, top). Cell cycle re-entry was quantified by EdU incorporation in the YFP+ myofibre progeny at 13 dpa. Frozen sections (5–10 μm) were thawed at room temperature and fixed in 4% formaldehyde for 5 min. Sections were blocked with 5% donkey serum and 0.1% Triton-X for 30 min at room temperature. Sections were incubated with anti-GFP (Abcam 6673), anti-PAX7 (DSHB) or anti-MHC (DSHB) overnight at 4 °C and with secondary antibodies for 1 h at room temperature. Antibodies were diluted in blocking buffer and sections were mounted in mounting medium (DakoCytomation) containing 5 μg ml−1 DAPI (Sigma). EdU detection was performed as previously described15. An LSM 700 Meta laser microscope with LSM 6.0 Image Browser software (Carl Zeiss) was used for confocal analyses. One in every eight sections was selected and labelled. For PAX7+ satellite cell counting, three sections were randomly selected and counted. For blastema YFP+ cell counting, all the sections in the region from regenerate tip to the bone were counted. Statistical analyses were performed using GraphPad Prism 6.0 (GraphPad Software). Student’s t-test, parametric, two-tail testing was applied to populations to determine the P values indicated in the figures. Significance was considered to have been reached at P values from <0.05. No statistical methods were used to predetermine sample size. In vivo axolotl experiments were not randomized and no blind tests were applied.


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No statistical methods were used to predetermine sample size. The experiments were not randomized. The investigators were not blinded to allocation during experiments and outcome assessment. The strains used in this study are listed in Supplementary Table 1. Unless otherwise specified, for all deletion mutants, the corresponding alleles from the Keio collection22 were transferred into the MC4100 wild-type strain using P1 transduction standard procedures23 and checked by PCR. To excise the resistance cassette, we used pCP20 (refs 22, 24). Strain AG227, deleted for the entire yedYZ operon, was constructed as follows. First, a cat-sacB cassette, encoding chloramphenicol acetyl transferase and SacB, a protein conferring sensitivity to sucrose, was amplified from strain CH1990 using primers yedYZ::cat-sacB_Fw and yedYZ::cat-sacB_Rv. The resulting PCR product shared a 40-base-pair (bp) homology to the 5′ untranslated region of yedY (msrP) and to the 3′ untranslated region of yedZ (msrQ) at its 5′ and 3′ ends, respectively. After purification, the PCR product was transformed by electroporation into CH1940. These cells harbour the pSIM5-tet vector, which encodes the Red recombination system proteins Gam, Beta and Exo under the control of the temperature-sensitive repressor cI859, encoded by the same vector. Induction of the Gam, Beta and Exo proteins was induced by shifting the cells to 42 °C for 15 min before making them electrocompetent. Recombinant cells were selected on chloramphenicol-containing plates (25 μg ml−1) at 37 °C for 16 h. At this temperature, the pSIM5-tet vector, which has a temperature-sensitive origin of replication, is lost. Colonies were also tested for the presence of the cat-sacB cassette by negative selection on sucrose-containing media (5% sucrose, no NaCl). Finally, we verified that the cat-sacB cassette replaced the msrPQ operon in the resulting strain (AG219) by sequencing across the junctions. The cat-sacB cassette was subsequently moved from AG219 to TP1004 by P1 transduction. The cat-sacB cassette was eliminated from the resulting strain (AG220) by transforming it with the pSIM5-tet plasmid, electroporating it with the oligonucleotide Delta_yedYZ (300 ng) and performing lambda red recombination as described above. Recombinants were selected on sucrose-containing media at 30 °C for 16 h. To eliminate the plasmid, the selected colonies were grown at 37 °C for 16 h. Loss of the cassette in the resulting AG227 strain was verified by positive (sucrose resistance) and negative (chloramphenicol sensitivity) selection and by PCR. The msrQ deletion mutant (strain BE105) was generated using the PCR knockout method developed in ref. 24. Briefly, a DNA fragment containing the cat gene flanked with the homologous sequences found upstream and downstream of the yedZ gene was PCR-amplified using pKD3 as template and the oligonucleotides P1_Up_YedZ and P2_Down_YedZ. Strain BE100, carrying plasmid pKD46, was then transformed by electroporation with the amplified linear fragment. Chloramphenicol-resistant clones were selected and verified by PCR. The msrP::lacZ fusion was constructed using the method described in ref. 25. Briefly, the msrP promoter region lying between nucleotide −797 and nucleotide +63, using the A nucleotide within the initiation triplet as a reference, was amplified by PCR with the appropriate oligonucleotides (lacI-msrP and lacZ-msrP' ). Using mini-lambda-mediated recombineering, the PCR product was then directly recombined with the chromosome of a modified E. coli wild-type strain (PM1205), carrying a P -cat-sacB cassette inserted in front of lacZ, at the ninth codon. Recombinants were selected for loss of the cat-sacB genes, resulting in the translational fusion of msrP to lacZ. The plasmids and primers used in this study are listed in Supplementary Tables 2 and 3, respectively. The YedY-His (MsrP-His ) expression vector was constructed as follows. Site-directed mutagenesis using primers pTAC_NdeI_Fw and pTAC_NdeI_Rv was performed using pTAC-MAT-Tag-2 as template to introduce an NdeI restriction site in the vector, yielding vector pAG177. yedY (msrP) DNA was amplified from the chromosome (MC4100) using primers pTAC_yedY_Fw and pTAC_yedY-His _Rv, which resulted in the fusion of a His tag coding sequence at the 3′ end. The PCR product was subsequently cloned into pAG177 using NdeI and BglII restriction sites, generating plasmid pAG178. To construct IPTG-inducible pTAC-MAT-Tag-2 vectors expressing either MsrP (without tag) or both MsrP and MsrQ, we first amplified the corresponding coding DNA sequences (msrP or the msrPQ operon) from the chromosome of strain MC4100 using primer pairs pTAC_yedY_Fw/ pTAC_yedY_Rv and pTAC_yedY_Fw/ pTAC_yedZ_Rv, respectively. The PCR products were then cloned into pAG177 using restriction sites NdeI and BglII, yielding pAG192 (MsrP) and pAG195 (MsrPQ). The complementation pAM238 vectors constitutively expressing either MsrP or MsrQ alone (without tag) or both MsrP and MsrQ were constructed as follows. We first amplified the corresponding coding DNA sequences (msrP, msrQ or the msrPQ locus) in addition to a 50 bp upstream region from each start codon (to include a ribosomal binding site) from the chromosome of strain MG1655 using primer pairs pAM238_yedY_Fw/ pAM238_yedY_Rv, pAM238_yedZ_Fw/ pAM238_yedZ_Rv and pAM238_yedY_Fw/ pAM238_yedZ_Rv, respectively. The PCR products were then cloned into pAM238 using restriction sites KpnI and PstI, yielding pAG264 (MsrP), pAG275 (MsrQ), and pAG265 (MsrPQ). The vector allowing the arabinose-inducible expression of SurA was constructed as follows. The surA-encoding DNA and its 50 bp upstream region (to include a ribosomal binding site) were amplified from the chromosome of strain MG1655 using the primer pair surA_Fw/surA_Rv. The PCR product was then cloned into pBAD33 using restriction sites KpnI and XbaI, yielding vector pAG290. Expression levels of the yedYZ (msrPQ) mRNA were assessed in M63 minimal medium supplemented with 0.5% glycerol, 0.15% casamino acids, 1 mM MgSO , 1 mM MoNa O , 17 μM Fe (SO ) and vitamins (thiamine 10 μg ml−1, biotin 1 μg ml−1, riboflavin 10 μg ml−1 and nicotinamide 10 μg ml−1). Overnight cultures of MG1655 were diluted to A  = 0.04 in fresh M63 minimal medium (100 ml) and cultured aerobically at 37 °C until A  = 0.8. Cells (10 ml) were then pelleted, resuspended in TriPure (Roche) and homogenized. After mixing with chloroform, RNA was isolated by centrifugation (15 min, 15,700g, 4 °C), precipitated with isopropanol, washed with ethanol 70%, dried and finally resuspended in DEPC water. Any residual DNA was eliminated by treatment of the sample with DNase (Turbo DNA-free Kit, Ambion). A RevertAid RT kit (Thermo Scientific) was used to generate complementary DNA (cDNA) from 1 μg RNA extracted from each of the cultured strains. cDNAs were then diluted 1/10 and submitted to qPCR, using a qPCR Core kit for SYBR Green I No ROX (Eurogentec) and a MyiQ Single-Colour Real-Time PCR Detection System (Bio-Rad). Expression levels of yedYZ were normalized to the expression of gapA. Primers used for qPCR analysis were qPCR_yedYZ_Fw and qPCR_yedYZ_Rv for yedYZ, and qPCR_gapA_Fw and qPCR_gapA_Rv for gapA (Supplementary Table 3). Synthesis of MsrP in strains JB590 and BE100 was assessed as follows. Overnight cultures were diluted to A  = 0.04 in fresh M63 minimal medium (100 ml) and cultured aerobically at 37 °C until A  = 0.8. Nine hundred microlitres of each culture were then precipitated with 10% ice-cold trichloroacetic acid (TCA), pellets were washed with ice-cold acetone, dried, resuspended and heated at 95 °C in Laemmli SDS sample buffer (SB buffer) (2% SDS, 10% glycerol, 60 mM Tris-HCl, pH 7.4, 0.01% bromophenol blue), and loaded on an SDS–PAGE gel for immunoblot analysis. The protein amounts loaded were standardized by taking into account the A values of the cultures. To monitor the MsrP expression levels after NaOCl or H O treatment, overnight cultures of wild-type cells (MG1655) were diluted to A  = 0.04 in fresh lysogeny broth (LB) medium (100 ml) and grown aerobically at 37 °C to A  = 0.5. NaOCl (2 mM) or H O (1 mM) was then added to the cultures. Samples were TCA-precipitated, washed with ice-cold acetone, dried, suspended in SB buffer, heated at 95 °C and loaded on an SDS–PAGE gel for immunoblot analysis. The protein amounts loaded were standardized by taking into account the A values of the cultures. The specificity of the anti-MsrP antibody was verified (Supplementary Fig. 5). l-Methionine sulfoxide ([α] 24 = +14.3° (water)), triethylamine (>99%) and methanol (>99.6%) were obtained from Sigma-Aldrich, picric acid from Prolabo and D O from SDS. Water was purified using Millipore Elix Essential 3 apparatus. 1H and 13C NMR were recorded on a Bruker Avance III Nanobay spectrometer (1H: 400 MHz; {1H}13C: 100 MHz). Chemical shifts (δ) were referenced to dioxane (1H: δ = 3.75 p.p.m.; 13C: δ = 67.19 p.p.m.)26, which was added as an internal reference; resonances are detailed as follows: 1H, δ in parts per million (multiplicity, J-coupling in hertz, integration, signal attribution); {1H}13C, δ in parts per million (signal attribution). For each diastereoisomer, chemical shifts are similar to those previously reported27. 13C resonance assignments were confirmed by heteronuclear single quantum coherence experiments. Optical rotations were measured on an Anton Paar Modular Circular Polarimeter 200 instrument at 25 °C and 589 nm from aqueous solution containing 0.8–1.2 g per 100 ml of l-methionine sulfoxide. The values reported are the average and s.d. relative to three independent measurements recorded on distinct solutions. The commercial mixture of diastereoisomers was separated following the previously reported method28. Briefly, 10 ml of water was added to l-methionine sulfoxide (1.333 g, 8.069 mmol) and picric acid (1.849 g, 8.071 mmol). The suspension was heated to reflux until complete dissolution and then slowly cooled to room temperature (~25 °C). The suspension was filtered on a sintered funnel and the solid was washed with cold water (10 ml in total). Both the solid (dextro) and filtrate (levo) were collected separately for further purification. Dextro. To the dried solid, 20 ml of water were added and the mixture was heated to reflux then allowed to cool slowly to room temperature. The solid was filtered out, washed with 10 ml water and dried. Again, 11 ml of methanol were added to the resulting solid and the mixture heated to reflux. After slow cooling, the yellow crystals were filtered, washed with 5 ml methanol and dried. A portion was used for structure determination by X-ray analysis. To the dextrogyre picrate salt (1.345 g, 3.42 mmol), ~1.1 equivalents of triethylamine were added as a dilute aqueous solution (22 ml, 175 mM, 3.85 mmol). Subsequently, 200 ml of acetone were added portion-wise to the above stirring suspension and a white solid precipitated. This was filtered, washed, triturated with acetone and finally dried in vacuum (533 mg, 80%). Levo. The volume of the filtrate was reduced in vacuum at 40 °C to about 3–4 ml to obtain a saturated solution and a small amount of precipitate. Then, 1.5 ml of water were added, the suspension was filtered and the solid washed with minimal water (2 ml). The whole step was repeated once (reduce the volume, dilute, filter and wash), and the resulting solution was then completely dried in vacuum. To the resulting yellow residue, 15 ml of methanol were added and the suspension was heated to reflux. In our hands, no solid precipitated upon cooling (in contrast with the reported method28); therefore the solution was dried again in vacuum. Following the same protocol as before, to the levogyre-enriched picrate salt (1.354 g, 3.44 mmol), ~1.1 equivalents of triethylamine were added as a concentrated aqueous solution (3.8 ml, 1 M, 3.8 mmol). Afterwards, 200 ml of acetone were added portion-wise and a white solid precipitated. This was filtered, washed, triturated with acetone and finally dried in vacuum (515 mg, 77%). Levo (l-methionine-R-sulfoxide): [α] 25 = −72.7 ± 0.5° (water); 1H NMR (400 MHz, D O pD = 6.5): 3.86 (t, 3J = 6.3, 1H, Hα ), 3.12 (ddd, J = 13.4, 9.6, 7.0, 1H, Hγ or Hγ ), 3.02 (m, 2H, Hγ ), 2.93 (ddd, J = 13.5, 9.1, 6.8, 1H, Hγ or Hγ ), 2.74 (s, 3H, Hε ), 2.31 (m, 2H, Hβ ); {1H}13C NMR (100 MHz, D O): 173.9 (COO ), 54.2 (Cα ), 54.0 (Cα ), 48.9 (Cγ ), 37.2 (Cε ), 37.0 (Cε ), 24.4 (Cβ ). Literature values from ref. 28: [α] 26 = −71.6° (water), from ref. 27: [α]  = −78° (water, room temperature); 1H NMR (300 MHz, D O): 4.10 (m, 1H), 3.08–2.78 (m, 2H), 2.59 (s, 3H), 2.32–2.13 (m, 2H); 13C NMR (75 MHz, D O): 171.1, 52.1, 48.4, 37.0, 23.7. In the 1H NMR spectra, the resonance centred at 3.02 p.p.m. was attributed to the S- enantiomer. The relative integral values suggest that R-Met-O is contaminated by 3% of the S- diastereoisomer. Moreover, comparing the measured [α] 25 values with those reported in ref. 27, the data are consistent with the presence of 3% S- diastereoisomer as a contaminant. Such purity is in line with previous reports using the same separation method28, 29. The absolute configuration of the l-methionine-S-sulfoxide was confirmed by X-ray structural analysis and matches previous assignments27, 30. To synthesize N-acetyl-Met-O, Met-O (30 mg; Sigma-Aldrich) was solubilized in 2 ml 100% acetic acid. After addition of 2 ml of 97% acetic anhydride, the resulting mixture was incubated 2 h at 23 °C. Then, 2 ml of water were added and the mixture was lyophilized overnight. Finally, the lyophilized N-acetyl-Met-O was washed three times with 6 ml of water, re-lyophilized and suspended in 500 mM Na HPO , pH 9.0 to a final concentration of 1.5 M. The pH was then adjusted to 7 with NaOH. The MsrP reductase activity was followed spectrophotometrically at 600 nm by monitoring the substrate-dependent oxidation of reduced benzyl viologen, serving as an electron donor. Reactions were performed anaerobically at 30 °C in degassed and nitrogen-flushed 50 mM MOPS, pH 7.0 using stoppered cuvettes. Benzyl viologen was used at a final concentration of 0.4 mM (molar extinction coefficient, ε, of reduced benzyl viologen = 7,800 M–1 cm–1) and reduced with sodium dithionite. The final reaction volume was kept constant, with the ordered addition of benzyl viologen, sodium dithionite, 1–32 mM N-acetyl-methionine sulfoxide (NacMet-O) and 10 nM MsrP-His . The concentrations used for the R- and S-Met-O diastereoisomers were 1–64 mM. The Michaelis–Menten parameters (maximum velocity (V ) and K ) were determined using Graphpad Prism software. The reductase activities of MsrA and MsrB were followed spectrophotometrically at 340 nm by monitoring the substrate-dependent oxidation of NADPH (ε = 6,220 M–1 cm–1). Reactions were performed at 37 °C in HEPES–KOH 20 mM, pH 7.4, NaCl 10 mM, and the final reaction volumes were kept constant, with the ordered addition of 250 μM NADPH (Roche), 2.6 μM TrxR, 40 μM Trx, 64 mM substrate and 1.5 μM of either MsrA or MsrB. The identification of the MsrP substrates was performed as follows. AG89 cells (2L) were grown aerobically at 37 °C in terrific broth to A  = 0.8. Periplasmic extracts were prepared as described previously31. Briefly, cells were pelleted by centrifugation at 3,000g for 20 min at 4 °C and incubated on ice with gentle shaking for 30 min in 100 mM Tris-HCl, pH 8.0, 20% sucrose, 1 mM EDTA. This mixture also contained 20 mM N-ethylmaleimide to alkylate reduced cysteine residues in proteins to prevent their subsequent oxidation. Periplasmic proteins were then isolated by centrifugation of the cells at 3,000g for 20 min at 4 °C. The periplasmic extract was subsequently concentrated by ultrafiltration in an Amicon cell (3,000 Da cutoff, YM-3 membrane) and loaded on a PD-10 column (GE Healthcare) equilibrated with 50 mM NaPi, pH 8.0, 50 mM NaCl. After concentration using a 5 kDa cutoff Vivaspin 4 (Sartorius) concentrator, the extract was finally separated in three samples. Two samples were incubated 10 min at 37 °C with 2 mM NaOCl whereas the third was left untreated to serve as reduced control. NaOCl was then removed by gel filtration using a NAP-5 column (GE Healthcare) equilibrated with 50 mM MOPS, pH 7.0. The untreated sample was also subjected to the NAP-5 gel filtration. One of the NaOCl-oxidized fractions was then reduced in vitro by incubation for 1 h at 37 °C with 10 μM MsrP, 10 mM benzyl viologen and an excess of sodium dithionite. The other NaOCl-oxidized fraction, used as an oxidized control, and the non-oxidized fraction were incubated with 10 mM benzyl viologen and an excess of sodium dithionite but without MsrP. The three samples were then de-salted by dialysis against 50 mM MOPS, pH 7.0 by using Slide-A-Lyzer 3,500 MWCO G2 cassettes (Thermo Scientific). The three samples (500 μg) were precipitated by adding TCA to a final concentration of 10% w/v. The resulting pellets were washed with ice-cold acetone, dried in a Speedvac, suspended in 0.1 M NH HCO , pH 8.0, digested overnight at 30 °C with 3 μg sequencing-grade trypsin, and analysed by two-dimensional LC–MS/MS essentially as described32. Briefly, peptides were first separated on a first-dimension hydrophilic interaction liquid chromatography (HILIC) column with a reverse acetonitrile gradient and 25 fractions of 1 ml collected (2 min per fraction). After drying, peptides were analysed by LC–MS/MS on a C18 column. The MS scan routine was set to analyse by MS/MS the five most intense ions of each full MS scan; dynamic exclusion was enabled to assure detection of co-eluting peptides. Raw data collection of approximately 230,000 MS/MS spectra per two-dimensional LC–MS/MS experiment was followed by protein identification using SEQUEST. All MS raw files have been deposited in the ProteomeXchange Consortium33 via the PRIDE partner repository with the data set identifier PXD002804. In detail, peak lists were generated using extract-msn (ThermoScientific) within Proteome Discoverer 1.4.1. From raw files, MS/MS spectra were exported with the following settings: peptide mass range 350–5,000 Da; minimal total ion intensity 500. The resulting peak lists were searched using SequestHT against a target-decoy E. coli protein database (release 07.01.2008, 8,678 entries comprising forward and reverse sequences) obtained from Uniprot. The following parameters were used: trypsin was selected with proteolytic cleavage only after arginine and lysine, number of internal cleavage sites was set to 1, mass tolerance for precursors and fragment ions was 1.0 Da, and considered dynamic modifications were +15.99 Da for oxidized methionine and +125.12 Da for N-ethylmaleimide on cysteines. Peptide matches were filtered using the q value and posterior error probability calculated by the Percolator algorithm ensuring an estimated false positive rate below 5%. The filtered SEQUEST HT output files for each peptide were grouped according to the protein from which they were derived using the multiconsensus results tool within Proteome Discoverer. Then the values of the spectral matches of only Met-containing peptides were combined from the three two-dimensional LC–MS/MS experiments and exported in a Microsoft Excel spreadsheet, with the rows referring to the peptide sequences and the columns to the fractions of the HILIC column. Oxidation of Met residues to Met-O by NaOCl causes a hydrophilic shift, which influences their retention time and makes them elute later (4–8 min) than their reduced counterpart on a HILIC column. If these Met-O are reduced by MsrP, they will then show a hydrophobic shift and elute at the same retention time on the HILIC column as in the control sample. By comparing the retention times and the number of peptide spectral matches of the Met-O-containing peptides in a periplasmic extract under three experimental conditions (control, oxidized by NaOCl with and without MsrP), one can identify ‘bona fide’ potential MsrP substrates. TP1004 cells harbouring plasmid pAG178 and overexpressing MsrP-His protein were grown aerobically at 30 °C in terrific broth (Sigma-Aldrich) supplemented with sodium molybdate (1.5 mM) and ampicillin (200 μg ml−1). When cells reached A  = 0.8, expression was induced with 0.1 mM IPTG for 3 h. Periplasmic proteins were then extracted as in ref. 32. MsrP-His was then purified by loading the periplasmic extract on a 1 ml HisTrap FF column (GE Healthcare) equilibrated with buffer A (NaPi 50 mM, pH 8.0, NaCl 300 mM). After washing the column with buffer A, MsrP-His was eluted by applying a linear gradient of imidazole (from 0 to 300 mM) in buffer A. The fractions containing MsrP-His were pooled, concentrated using a 5 kDa cutoff Vivaspin 15 (Sartorius) device and de-salted on a PD-10 column (GE Healthcare) equilibrated with 50 mM NaPi, pH 8.0, 150 mM NaCl. VU1 CaM, MsrA and MsrB were expressed and purified as described previously34, 35. Trx was expressed and purified as follows. BL21 (DE3) cells harbouring plasmid pMD205, overexpressing Trx with a carboxy (C)-terminal His tag, were grown aerobically at 37 °C in LB supplemented with kanamycin (50 μg ml−1). Expression was induced at A  = 0.6 with 1 mM IPTG for 3 h. Cells were then pelleted, resuspended in buffer A (NaPi 50 mM, pH 8.0, NaCl 300 mM) and disrupted by two passes through a French pressure cell at 12,000 psi. The lysate was then centrifuged at 30,000g and at 4 °C for 45 min, to remove cell debris, and Trx was purified as described for MsrP-His . Ni-NTA-purified Trx was then loaded on a 120 ml HiLoad 16/60 Superdex 75 PG column (GE Healthcare) previously equilibrated with HEPES–KOH 50 mM, pH 7.4, NaCl 100 mM. The resulting Trx-containing fractions were pooled and concentrated using a 5 kDa cutoff Vivaspin 15 device. Thioredoxin reductase (TrxR) was expressed and purified as follows. BL21 (DE3) cells harbouring plasmid pPL223-2, overexpressing TrxR with an amino (N)-terminal His tag, were grown aerobically at 37 °C in LB supplemented with ampicillin (200 μg ml−1). Expression was induced at A  = 0.6 with 1 mM IPTG for 3 h. Protein extraction was performed as described for Trx and purification was performed as described for MsrP-His . BL21 (DE3) cells harbouring plasmid pKD11, overexpressing SurA with a C-terminal His tag, were grown aerobically at 37 °C in LB supplemented with kanamycin (50 μg ml−1). Expression was induced at A  = 0.6 with 1 mM IPTG for 3 h. Protein extraction and purification were performed as described for MsrP-His . MG1655 cells harbouring plasmid pKD84, overexpressing SurA with a C-terminal Strep-tag, were grown aerobically at 37 °C in LB supplemented with ampicillin (200 μg ml−1). Expression was induced at A  = 0.7 with a final concentration of 200 μg l−1 anhydrotetracycline (AHT) for 5 h. Protein extraction was performed as described for MsrP-His . SurA-Strep was then purified by loading the periplasmic extract on a 5 ml Strep-Tactin Superflow cartridge H-PR (IBA) equilibrated with buffer A (Tris-HCl 100 mM, pH 8.0, NaCl 150 mM, EDTA 1 mM). After washing the column with buffer A, SurA-Strep was eluted by applying a linear gradient of desthiobiotin (from 0 to 2.5 mM) in buffer A. The fractions containing SurA-Strep were pooled, concentrated using a 5 kDa cutoff Vivaspin 15 (Sartorius) device and de-salted on a PD-10 column (GE Healthcare) equilibrated with 50 mM NaPi, pH 8.0, 150 mM NaCl. A modified version of Pal lacking the signal sequence and in which the first cysteine of the lipobox was replaced by an alanine (Pal ) was expressed with an N-terminal His tag from the pEB0513 vector in BL21 (DE3) cells. Cells were grown aerobically at 37 °C in LB supplemented with ampicillin (200 μg ml−1). Expression was induced at A  = 0.6 with 1 mM IPTG for 3 h. Protein extraction was performed as described for Trx and purification was performed as described for MsrP-His . CaM was oxidized in vitro as described previously36. SurA-His and Pal were oxidized in vitro by incubating the purified proteins (50 μM) for 2 h 30 min at 30 °C with 100 mM H O in a buffer containing 50 mM NaPi, pH 8.0, 50 mM NaCl. H O was then removed by gel filtration using a NAP-5 column (GE Healthcare) equilibrated with 50 mM NaPi, pH 8.0, 150 mM NaCl. In vitro repair of oxidized CaM (CaMox), SurA (SurA ox) and Pal (Pal ox) was assessed by incubating the oxidized proteins (2 μM of CaMox and SurA ox, 5 μM of Pal ox) with purified MsrP-His (2 μM for CaMox and SurA ox, 5 μM for Pal ox), 10 mM benzyl viologen and an excess of sodium dithionite at 37 °C for 1 h. As controls, the oxidized proteins were incubated separately with either MsrP-His or the inorganic reducing system (benzyl viologen and sodium dithionite). The reactions were stopped by adding SB buffer and heating at 95 °C for the CaM and SurA samples or by adding 0.1% trifluoroacetic acid for the Pal samples. The CaM and SurA samples were then loaded on an SDS–PAGE gel and the proteins visualized with the PageBlue Protein Staining Solution (Fermentas). For the Pal samples (20 μg), proteins were separated by reverse-phase high-performance liquid chromatography on a C4 column (Vydac 214TP54, 4.6 mm × 250 mm) at a flow rate of 400 μl min−1 with a linear gradient of acetonitrile in 0.1% trifluoroacetic acid (0–70% acetonitrile in 90 min). Absorbance was monitored at 214 nm and the peaks were collected. The fractions were dried in a Speedvac and the proteins resupsended in 25 μl of 100 mM NH HCO before overnight digestion at 30 °C with 0.5 μg of trypsin or EndoGlu-C. The peptides were then analysed as described below. For CaM and SurA, the gel bands corresponding to the different oxidation states were in-gel digested with trypsin and the resulting peptides analysed by LC–MS/MS on a C18 reverse-phase column as described above. Relative abundances of every Met-containing peptide in its different oxidation state were obtained by integration of peak area intensities, taking into account the extracted ion chromatogram of both doubly and triply charged ions. The in vivo repair of SurA ox and Pal ox by the MsrPQ system or MsrP alone expressed from plasmids pAG195 and pAG192, respectively, was performed as follows. Overnight cultures of AG233 (containing the empty pAG177 vector), AG234 (containing the pAG195 plasmid) and AG289 (containing the pAG192 plasmid) were diluted to A  = 0.04 into fresh LB medium (100 ml) and cells were grown aerobically at 37 °C in the presence of 0.1 mM IPTG and 200 μg ml−1 ampicillin. At A  = 0.5, cells were subjected to NaOCl treatment (3.5 mM) and protein synthesis was blocked by the addition of chloramphenicol (300 μg ml−1). Samples were taken at different time points after NaOCl addition and precipitated with TCA. The pellets were then washed with ice-cold acetone, suspended in SB buffer, heated at 95 °C and loaded on a SDS–PAGE gel for immunoblot analysis using anti-Pal37 and anti-SurA antibodies. The specificity of the anti-SurA antibody was verified (Supplementary Fig. 6). The protein amounts loaded were standardized by taking into account the A values of the cultures. SurA-Strep was oxidized in vitro by incubating the purified protein (200 μM) for 3 h at 30 °C with 100 mM H O in a buffer containing 50 mM NaPi, pH 8.0, 150 mM NaCl. H O was then removed by gel filtration using a NAP-5 column (GE Healthcare) equilibrated with 50 mM NaPi, pH 8.0, 150 mM NaCl. For the in vitro repair of oxidized SurA (SurA ox), the oxidized protein (30 μM) was incubated with purified MsrP-His (30 μM), 10 mM benzyl viologen and 10 mM of sodium dithionite at 37 °C for 1 h. Following repair, SurA was purified by passing the sample through a gravity flow column containing 200 μl Strep-Tactin Sepharose beads (from a 50% suspension, IBA), previously equilibrated with buffer A (Tris-HCl 100 mM, pH 8.0, NaCl 150 mM, EDTA 1 mM). After washing with buffer A, repaired SurA was eluted using buffer A containing 2.5 mM desthiobiotin. The elution fractions were pooled and submitted to buffer exchange using a NAP-5 column (GE Healthcare) equilibrated with 50 mM NaPi, pH 8.0, 150 mM NaCl. To check for the correct oxidation, repair and purification of SurA, samples were loaded on an SDS–PAGE gel and the proteins visualized with the PageBlue Protein Staining Solution (Fermentas). The ability of SurA to act as a chaperone preventing the thermal aggregation of citrate synthase (Sigma, reference C3260) was assessed as follows. The aggregation of citrate synthase (0.15 μM) was monitored at 43 °C in 40 mM HEPES–KOH, pH 7.5, in the absence or in the presence of 0.6 μM SurA, SurA ox or MsrP-repaired SurA ox using light-scattering measurements. To avoid effects that might have been caused by the protein buffer, all samples were added to the assay in constant volume. SurA ox and MsrP-repaired SurA ox were obtained as described above. Light-scattering measurements were made using a Varian Cary Eclipse spectrofluorometer both with excitation and with emission wavelengths set to 500 nm at a spectral bandwidth of 2.5 nm. Data points were recorded every 0.1 s. The ability of various E. coli strains (BE100, JB08, CH193, BE104) to assimilate Met-O was assessed on M9 minimal medium supplemented with either Met or Met-O at 20 μg ml−1. Plates were incubated at 37 °C for 72 h. Overnight cultures of strains AG272, AG273, AG279 and AG274 were diluted to A  = 0.04 into fresh M63 minimal medium (100 ml) supplemented with 0.5% glycerol, 150 μg ml−1 of each amino acid, 1 mM MgSO , 1 mM MoNa O , 17 μM Fe (SO ) , vitamins (thiamine 10 μg ml−1, biotin 1 μg ml−1, riboflavin 10 μg ml−1, and nicotinamide 10 μg ml−1) and 100 μg ml−1 spectinomycin, and grown aerobically at 37 °C. When A reached 0.5, cells (5 ml) were washed three times with M63 medium containing 150 μg ml−1 Met-O instead of methionine, and serially diluted in the same medium. Five microlitres of each dilution were then spotted on M63 plates containing either Met or Met-O at 150 μg ml−1, and plates were subsequently incubated at 37 °C for 40 h. The msrP::lacZ-containing strains (CH183, CH186 and CH187) were grown at 37 °C with shaking in M9 minimal medium. When cells reached A ≈ 0.2, cultures were split into two plastic tubes, one of them containing HOCl (200 μM). These tubes were then incubated with an inclination of 90° with shaking at 37 °C. After 30 min of incubation, 1 ml was harvested and the bacteria were resuspended in 1 ml of β-galactosidase buffer. Levels of β-galactosidase were measured as described38. NR744, NR745, CH0127 and AG190 cells were grown aerobically at 37 °C with shaking in 50 ml of LB medium in 500 ml flasks. When cells reached A ≈ 0.45, 5 ml samples were transferred to conical polypropylene centrifuge tubes (50 ml; Sarstedt) and HOCl (2 mM) was added. Cells were then incubated at 37 °C with shaking (150 r.p.m.) at 90° inclination. Samples were taken at various time points after stress, diluted in PBS buffer, spotted on LB agar and incubated at 37 °C for 16 h. Cell survival was determined by counting colony-forming units (c.f.u.) per millilitre. The absolute c.f.u. at time-point 0 (used as 100%) was ~108 cells per millilitre in all experiments. For strains CH194, CH196 and CH197, the same protocol was used with chloramphenicol (25 μg ml−1) and arabinose (0.2%) added to the cultures. Cells (MG1655 and BE107) were grown at 37 °C with shaking in 10 ml of LB (in 100 ml flasks). When cells reached A ≈ 0.8, 5 ml samples were transferred to conical polypropylene centrifuge tubes (50 ml, Sarstedt) and HOCl (2 mM) was added. After 5 min of incubation, samples were taken and diluted in PBS buffer to ~2 × 103 cells per millilitre. Aliquots (100 μl) were then spread on LB agar plates containing SDS (1%). Colonies were counted the next day. A non-redundant local protein database containing 1,342 complete prokaryotic proteomes available in NCBI (http://www.ncbi.nlm.nih.gov/) as of 30 July 2014 was built. This database was queried with the BlastP program (default parameters)39, using YedY (NP_416480) and YedZ (NP_416481) of E. coli strain K-12 substrate MG1655 as a seed. Distinction between homologous and non-homologous sequences was assessed by visual inspection of each BlastP output (no arbitrary cut-off on the E value or score). To ensure that we did not overlook divergent YedY or YedZ proteins, iterative BlastP queries were performed using homologues identified at each step as new seeds. The list of YedY and YedZ homologues is provided in Supplementary Data 1. The retrieved sequences were aligned using MAFFT version 7 (default parameters40; Supplementary Data 2 and 3). Each alignment was visually inspected and manually refined when necessary using the ED program from the MUST package41. Regions where the homology between amino-acid positions was doubtful were removed by using BMGE software (BLOSUM30 similarity matrix42). For each homologue, the genomic context was investigated using MGcV (Microbial Genomic context Viewer43). The domain composition and protein location of each homologue was also analysed using pfam version 27.0 (ref. 44), SignalP version 4.1 (ref. 45) and TMHMM server version 2.0 (ref. 46), respectively. For the YedY protein, preliminary phylogenetic analysis used FastTree version 2 and a gamma distribution with four categories47. On the basis of the resulting tree, the subfamily containing the sequence from E. coli was identified and selected for further phylogenetic investigations. The corresponding sequences were realigned using MAFFT version 7. The resulting alignment was trimmed with BMGE as previously described. Maximum likelihood trees were computed using PHYML version 3.1 (ref. 48) with the Le and Gascuel model (amino-acid frequencies estimated from the data set) and a gamma distribution (four discrete categories of sites and an estimated alpha parameter) to take into account variations in evolutionary rate across sites. Branch robustness was estimated by the non-parametric bootstrap procedure implemented in PhyML (100 replicates of the original data set with the same parameters). Bayesian inferences were performed using MrBayes 3.2 (ref. 49) with a mixed model of amino-acid substitution including a gamma distribution (four discrete categories). MrBayes was run with four chains for one million generations and trees were sampled every 100 generations. To construct the consensus tree, the first 2,000 trees were discarded as ‘burn in’.


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Klp98AΔ47 is a null allele generated by homologous recombination with the ‘ends-out’ strategy29, 30 using a pW25 plasmid containing two homology fragments flanking the coding region of Klp98A as described31 (2,848 bp of homology in the 5′ region and 4,011 bp in the 3′ region, Extended Data Fig. 1g). Upon recombination, this construct replaces the coding sequence of Klp98A by the whs gene flanked by two loxP sites followed by an AttP ΦC31 site. The whs gene is subsequently floxed to generate Klp98AΔ47 which corresponds to a deletion of the Klp98A coding sequence (Extended Data Fig. 1h). Gene deletion was confirmed by PCR (Extended Data Fig. 1i) and sequencing. Three zinc-finger nuclease pairs targeting Klp98A were designed and produced by Sigma-Aldrich (product number CSTZFNY-1KT, lot number 03041026MN). The target sequences were (cut site indicated with underlining): no. 1, CAGAGCACTGGGCATG GGTGCGGGAGCATCG; no. 2, CTTCGACTACTCCTATTG GATGCGGAGGATCCG; and no. 3, CTCTTTGCCCGCATG GGCCAGGAGTCGGGCA. The mRNAs corresponding to the three pairs were injected together at 40 ng μl−1 in w1118 embryos by BestGene Inc. Adults from these embryos were crossed with w;Df(3R)BSC497/TM6c. Df(3R)BSC497 is a deletion spanning the Klp98A gene (Flybase and our own unpublished data). The relevant progeny (about 50 individuals) was then analysed by PCR using primers flanking the three cut sites and the amplicons sequenced. We found deletions only in the region corresponding to the zinc-finger pair no. 1. We studied three of them in more detail: Klp98AΔ6, Klp98AΔ7 and Klp98AΔ8. Klp98AΔ6 is a G to C substitution at position 500 of the coding sequence of Klp98A (CG5658-PA) followed by a six-nucleotide deletion. The amino acid sequence at position 167 is therefore changed from 164TGHGLRVRE172 to 164TGHA—VRE170 (see Extended Data Fig. 1j). This two amino acid deletion maps into the L8 loop of the motor domain of Klp98A and does not affect the stability of the protein (see Extended Data Fig. 1k) but behaves like a strong mutant in transheterozygocity with Klp98AΔ47 (see Fig. 1h, Extended Data Fig. 3a, b, d–f). Klp98AΔ7 is a deletion of seven nucleotides at position 502 in the coding sequence of Klp98A, leading to a frameshift starting at amino acid 168 and causing a stop codon after amino acid 209. Klp98AΔ8 is a deletion of eight nucleotides at position 501, leading to a frameshift starting at amino acid 168 and causing a stop codon after amino acid 186. Full-length Klp98A protein is undetectable in homozygous Klp98AΔ47, Klp98AΔ7 and Klp98AΔ8 animals (Extended Data Fig. 1a, k). In this work, transheterozygous animals (that is, Klp98AΔ47/Δ6 and Klp98AΔ47/Δ8) were used in phenotypic analyses in order to avoid the effects of potential linked mutations. These transheterozygous combinations are viable and fertile. However, these mutants show Notch-dependent asymmetric cell fate assignation phenotypes when the other two systems controlling these events, that is, Numb and Neuralized, are compromised (see Fig. 1i, j and Extended Data Fig. 3). Transgenes used in this study included Ubi > mCherry-Pavarotti (generated for this study), UAS > Jupiter-mCherry (this study), UAS > Klp98A-mCherry (this study), UAS > Klp98A-GFP (this study), UAS-GFP-Patronin (this study), Asense > GFP-Pon (this study), Asense > mCherry-Pon (this study), Asense > GFP-Sara (this study), UAS-GBP-Pon (this study), UAS-GBP-mCherry-Pon (this study), UAS-GBP-Bazooka (this study), Jupiter-GFP knock-in at the endogenous locus (ref. 14, Bloomington no. 6836), UAS-mRFP-Pon (ref. 32), UAS-mRFP-Sara (ref. 1), Neur > Gal4 (ref. 33), Ubi > GFP-Pavarotti (ref. 34), UAS > GFP-Pon (ref. 35), pnr > Gal4, phyllopod > GFP-Pon (ref. 36), pnr > Gal4 (Bloomington no. 3039), UAS > DsRed (kind gift from François Karch), UAS > PatroninRNAi#1 (VDRC no. 108927, referred to as Patronin RNAi in the main text), UAS > PatroninRNAi#2 (VDRC no. 27654), UAS > Klp10ARNAi (ref. 37, VDRC no. 41534), UAS > Klp98ARNAi (VDRC no. 40605), UAS > NeuralizedRNAi (VDRC no. 108239), UAS > NumbRNAi (gift from W. Zhong, ref. 38), Df(3R)BSC497 (Bloomington no. 25001), Klp98AΔ47 (this study), Klp98AΔ6 (this study), Klp98AΔ7 (this study), Klp98AΔ8 (this study), NumbSW (ref. 9, gift from R. Stanewsky), Numb2 (kind gift from Roland Le Borgne), Numb15 (kind gift from Roland Le Borgne), UAS > lgl3A (ref. 39), GFP-Rab5 knock-in at the endogenous locus (ref. 40), YFP-Rab11 knock-in at the endogenous locus (ref. 41), YFP-Rab7 knock-in at the endogenous locus (ref. 41) and tub > Gal80ts (Bloomington no. 7017). The genotypes of mutant stocks were verified by PCR and sequencing, as well as the genotypes of the F1 progeny generated for interaction studies (Fig. 1i, j and Extended Data Fig. 3). Since the Jupiter–GFP gene trap is viable, fertile and does not induce visible phenotypes in the SOP lineage, we used it as an alternative to balancers for controls in gene interaction studies (Extended Data Fig. 3). Flies co-expressing GBP–Pon and GFP–Patronin (Fig. 4 and Extended Data Figs 8 and 9) displayed occasional polarity defects reflected by loss of mRFP–Pon asymmetry (See Extended Data Fig. 8d for quantification). Cells showing such polarity defects were excluded from subsequent analysis. We used Gal80ts to achieve low levels of Klp98A–GFP expression to prevent endosome fusion (Fig. 1a, Extended Data Figs 1c–e and 2a, b). Extended Data Fig. 3c: Klp98AΔ47/+: w1118;UAS > NeurRNAi/+;pnr > Gal4 UAS > DsRed Klp98AΔ47/TM6B (29 °C; outside the pnr expression region). pnr > NeurRNAi Klp98AΔ47/+: w1118;UAS > NeurRNAi/+;pnr > Gal4 UAS > DsRed Klp98AΔ47/TM6B (29 °C; inside the pnr expression region). pnr > NeurRNAi Klp98AΔ47/Δ8: w1118;UAS > NeurRNAi/+;pnr > Gal4 UAS > DsRed Klp98AΔ47/Klp98AΔ8 (29 °C; inside the pnr expression region; sibling of fly above). Extended Data Fig. 3d: w1118 (25 °C). numb2/SW Klp98AΔ47/Jupiter–GFP: w1118(/+);Numb2/NumbSW;Jupiter-GFP/Klp98AΔ47 (25 °C). numb2/SW Klp98AΔ47/Δ8: w1118(/+);Numb2/NumbSW;Klp98AΔ8/Klp98AΔ47 (25 °C; sibling of the fly above). pnr > numbRNAi Klp98AΔ47/+: w1118;pnr > Gal4 UAS > DsRed Klp98AΔ47/UAS > numbRNAi (29 °C). pnr > numbRNAi Klp98AΔ47/Δ8: w1118;pnr > Gal4 UAS > DsRed Klp98AΔ47/Klp98AΔ8 UAS > numbRNAi (29 °C). Extended Data Fig. 3e: numb2/SW Klp98AΔ47/Jupiter–GFP:w1118(/+);Numb2/NumbSW;Jupiter-GFP/Klp98AΔ47 (25 °C). numb2/SW Klp98AΔ47/Klp98AΔ6: w1118(/+);Numb2/NumbSW;Klp98AΔ6/Klp98AΔ47 (25 °C; sibling of the fly above). numb2/SW Klp98AΔ47/Jupiter–GFP: w1118(/+);Numb2/NumbSW;Jupiter–GFP/Klp98AΔ47 (25 °C). numb2/SW Klp98AΔ47/Klp98AΔ8:w1118(/+);Numb2/NumbSW;Klp98AΔ8/Klp98AΔ47 (25 °C; sibling of the fly above). numb15/SW Klp98A−/+: w1118(/+);Numb15/NumbSW;Klp98AΔ47/TM6B (25 °C) or w1118(/+);Numb15/NumbSW;Klp98AΔ6/TM6B (25 °C). numb15/SW Klp98AΔ47/Klp98AΔ6:w1118(/+);Numb2/NumbSW;Klp98AΔ6/Klp98AΔ47 (25 °C; sibling of the fly above). Most of these genotypes correspond to the F1 of crosses performed at 25 °C. Embryos were laid at 25 °C. Larvae were then shifted to 16 °C until puparium formation and 16 h before SOP imaging they were shifted to 25 °C or 29 °C, as indicated. In Extended Data Fig. 1a, k, larvae homozygous for the Klp98A mutants were used for western blot analysis instead of animals deriving from an outcross. All the open reading frames (ORFs) cloned by PCR for this study were flanked by FseI and AscI sites for convenient shuttling between compatible plasmids. eGFP was amplified from pEGFP C1 (Clontech). Pavarotti (CG1258-PA), Klp98A (CG5658-PA), Bazooka (CG5055-PA) and Patronin (CG33130) were amplified from cDNAs prepared from adult w1118 flies total RNA extracted in TRIzol (Life Technologies), followed by reverse transcription (Super Script II kit, Life Technologies). The Patronin cDNA that we cloned encodes a splicing isoform slightly smaller than previously reported Patronin cDNAs16, 23 and has been deposited in the NCBI database (BankIt1865736 Patronin KT953618). The Pon localization domain (corresponding to amino acids 474–670 of Pon35) was similarly cloned from cDNA from w1118 flies. In various transgenes in this work (driven by UAS or Ase promoters), this Pon localization domain is referred to as ‘Pon’ for simplicity. Sara was subcloned from pUAST–GFP–Sara42. For antibody production, we also cloned smaller fragments of Patronin (corresponding to amino acids 1,039–1,384, named Patronin-Cter thereafter) and Klp98A (corresponding to amino acids 401–1,265, named Klp98A-Cter thereafter). Jupiter–mCherry was generated by cloning Jupiter–GFP from cDNA prepared from Jupiter–GFP flies14 and by replacing the GFP (in the middle of the gene) by mCherry using site-directed mutagenesis. We also cloned the GFP-binding peptide (GBP), or so called GFP nanobody, a lama VHH single chain antibody against GFP43 either for protein production (His–GBP) or for expression of fusion proteins in the fly (GBP–Pon and GBP–Bazooka, see below). Klp98A–GFP–PC: for stable expression of Klp98A in S2 cells, the Klp98A ORF described above was subcloned into a modified pMT vector (Life Technologies), to which a Puromycin selection gene (amplified from the pCoPuro plasmid44) and a C-terminal tag (eGFP followed by PC, the Protein C epitope tag: EDQVDPRLIDG) were added. GST–Klp98A-Cter, and GST–Patronin-Cter: for expression of GST–Klp98A-Cter and GST–Patronin-Cter in bacteria, the ORFs described above were subcloned into a modified pGEX vector45. His–Klp98A-Cter, His–eGFP, and His–GBP: for expression of (His) -tagged Klp98A-Cter, GFP and GBP, these ORFs were subcloned into a modified pET28b vector, which tags the ORF at its N terminus with a (His) tag. UAS > GFP–Patronin, UAS > Jupiter–mCherry, UAS > GBP–Pon, UAS > GBP–mCherry–Pon UAS > GBP–Bazooka, UAS > Klp98A–mCherry, UAS > Klp98A–GFP: for expression in flies with the UAS/Gal4 system, the Patronin, Bazooka, Klp98A, Pon localization domain and Jupiter–mCherry ORFs described above were subcloned into modified pUAST4 vectors tagging the ORF with either N-terminal PC–eGFP (for Patronin, referred to as GFP–Patronin for simplicity), C-terminal mCherry–PC (for Klp98A), C-terminal eGFP (for Klp98A), N-terminal GBP (for the Pon localization domain and Bazooka), N-terminal GBP–mCherry (GBP followed by mCherry separated by a GGG linker, for the Pon localization domain) or leaving it untagged (for Jupiter–mCherry). N-terminal GFP tagging of Patronin has been previously shown to be functional16, as well as N-terminal tagging of Bazooka46. Ubi > mCherry–Pavarotti: for ubiquitous expression of mCherry–Pavarotti, Pavarotti was subcloned into a modified pUbi vector allowing the expression of mCherry–Pavarotti under the ubiquitin promoter. Ase > GFP–Pon, Ase > GFP–Sara and Ase > mCherry–Pon: for specific expression in SOPs independently of the UAS/Gal4 system, the Pon localization domain and Sara were subcloned into the pAsense GFP vector, which was created by inserting a 1,943-bp fragment upstream of the start codon of the Asense gene (amplified from w1118 flies genomic DNA) into the Green Pelican GFP plasmid (Drosophila Genomics Resource Center), which results in tagging the Pon localization domain (or Sara) with an N-Terminal GFP (Ase > GFP–Pon). Alternatively, the GFP was exchanged by quick-change PCR into mCherry to generate the pAsense mCherry vector, in which the Pon localization domain was subcloned. Injection of plasmids into Drosophila embryos to generate transgenics was performed by BestGene Inc. SDS–PAGE was performed using NuPAGE 4–12% Bis-Tris gels (Life Technologies) according to the manufacturer’s instructions. Colloidal Coomassie blue (Life Technologies) was used for total protein staining of gels. Gels were transferred on nitrocellulose membranes using iBLOT (Life Technologies) according to the manufacturer’s instructions. For western blot, we used all primary antibodies at 1 μg ml−1 in TBS, 0.2% BSA, 1 mM CaCl , 0.02% Thymerosal O/N at 4 °C. Western blots were revealed using HRP coupled antibodies (Jackson immunoResearch 1:10,000 dilution), Western Bright Quantum (Advansta) or SuperSignal West Pico (Pierce) chemiluminescence reagents and a Vilber Lourmat Fusion imager. Alternatively (Extended Data Fig. 6a), western blots were performed with fluorescently labelled anti-tubulin antibodies and imaged with an Ettan DIGE Imager (GE Healthcare). For gel source data, see Supplementary Fig. 1. For total fly extracts (Extended Data Fig. 1a, k), dissected brains, imaginal discs and salivary glands of second instar larvae were squashed into 500 μl of lysis buffer (25 mM NaF, 1 mM Na VO , 50 mM Tris pH 7.5, 1.5 mM MgCl2, 125 mM NaCl, 0.2% IGEPAL, 5% glycerol, 1 mM DTT and protease inhibitor cocktail (benzamidine (1 mM, Applichem), chymostatine (40 μg ml−1, Applichem), antipain (40 μg ml−1 Applichem), leupeptine (1 μM Applichem), pefabloc (1 mM) and PMSF (0.5 mM)). The extract was incubated 40 min at 4 °C with rocking, then cellular debris were cleared by centrifugation at 16,000g for 10 min at 4 °C. Extracts were then diluted in LDS sample buffer (Life Technologies) enriched with 2.5% β-mercaptoethanol and analysed by SDS–PAGE and western blot as above. For RNAi-treated S2 total cell extracts (Extended Data Fig. 6a), Drosophila S2 cells (UCSF, mycoplasm-free judged by DAPI staining) were cultured and incubated with 5 μg dsRNA for 4 days as previously described47. This dsRNA sequence corresponds to the sequence in the UAS > PatroninRNAi#1 fly stock (VDRC no. 108927). Cells were washed in XB (20 mM HEPES, 150 mM KCl, pH 7.7), resuspended in LDS sample buffer, boiled for 2 min, then treated with Benzonase (30 units μl−1, Sigma) and analysed by SDS–PAGE and western blot as above. Unless stated otherwise, reagents were from Sigma. All purification steps were performed at 4 °C. Protein concentrations were determined spectrophotometrically using absorbance at 280 nm or after SDS–PAGE using purified BSA as a standard, followed by quantifications by densitometry using ImageJ (http://imagej.nih.gov/ij/). GST- and His-tagged Klp98A-Cter were expressed in E. coli BL21 Rosetta 2 (Stratagene) by induction with 0.5 mM IPTG in Terrific Broth medium (Sigma) at 23 °C. Bacteria expressing GST–Klp98A-Cter were lysed enzymatically using 0.7 mg ml−1 lysosyme and 10 μg ml−1 DNase I (Roche) in lysis buffer (50 mM Tris, 150 mM NaCl, 1% Triton X-100, 1 mM DTT, 5% Glycerol, pH 7.6) enriched with protease inhibitors (Roche Mini) for 1 h at 4 °C with rocking. After clarification (12,000 r.p.m., Beckman JA 25.5), lysate was incubated with glutathione sepharose resin (glutathione sepharose 4B, Amersham) for 2 h at 4 °C and washed extensively in 50 mM Tris, 2 mM β-mercaptoethanol, 100 mM NaCl, 5 mM MgCl pH 7.5. Glutathione-sepharose-bound GST–Klp98A-Cter was then cleaved on column by an overnight incubation at 4 °C with 40 μg of TEV protease per mg of fusion protein. Klp98A-Cter was subsequently dialysed against PBS, concentrated to 1 mg ml−1 by ultrafiltration (Amicon Ultra-4 3k Millipore) and injected into rabbits for polyclonal antibody production (see Antibodies). For affinity purification of polyclonal anti-Klp98A antibodies, we purified His–Klp98A-Cter following a protocol similar to the one described above, but using NiNTA resin (Ni Sepharose High Performance, Amersham) and 10 mM imidazole in lysis and wash buffers. His–Klp98A-Cter was eluted by 20 mM HEPES, 150 mM KCl, 300 mM imidazole, 1 mM DTT, pH 7.7, dialysed against 20 mM HEPES, 150 mM KCl, 10% glycerol, 1 mM DTT, pH 7.7, concentrated by ultrafiltration to 7.3 mg ml−1 and finally coupled to amino-link sepharose resin (Pierce). His–GFP and His–GBP were expressed and purified from E. coli BL21 Rosetta 2 following the same procedure as for His–Klp98A-Cter. Final dialysis buffer was (20 mM HEPES, 150 mM NaCl, pH 7.7) for His–GFP and (20 mM HEPES, 150 mM NaCl, 5% glycerol, 15 mM imidazole, pH 7.7) for His–GBP. His–GFP and His–GBP were concentrated by ultrafiltration to 7.5 mg ml−1 and 2.34 mg ml−1, respectively, flash frozen in liquid N and kept at −80 °C. GST-tagged Patronin-Cter purification was similar to the one of GST–Klp98A, except that TEV was removed by using NiNTA resin before final dialysis. Tag-free Patronin-Cter was injected into rabbits for polyclonal antibody production. Alternatively, tag-free Patronin-Cter was coupled to amino-link sepharose resin for affinity purification these anti-Patronin antibodies (see Antibodies). Klp98A–GFP–PC (that is, full length Klp98 fused to GFP and the PC tag in Cter) was purified from a puromycin-resistant Schneider S2 stable cell line expressing Klp98A–GFP–PC under the inducible metallothionein promoter. To obtain this cell line, S2 cells were transfected with pMT Puro Klp98A–GFP–PC plasmid (see above) using Effectene (Qiagen). This stable cell line was subsequently grown and selected in Schneider medium (Life Technologies) enriched with 10% vol/vol fetal calf serum and 5 μg ml−1 puromycin (Applichem). The concentration of inducer (CuSO ) was subsequently gradually increased from 0.05 mM to 0.6 mM over 1 month so as to select clones able to express high amounts of Klp98A (whose overexpression is toxic). We then grew 100 15-cm plates of this pseudo-clone. Cells were harvested, washed in XB buffer (20 mM HEPES, 150 mM KCl, 1 mM CaCl , pH 7.7) then lysed in 100 ml of lysis buffer (20 mM HEPES, 150 mM KCL, 1% Triton X-100, 1 mM CaCl , 2 mM MgCl , 0.1 mM ATP, pH 7.2) supplemented with a protease inhibitor cocktail (benzamidine/chymostatine/antipain/leupeptine/pefabloc/PMSF, see SDS–PAGE and western blot section). Lysate was rocked for 1 h at 4 °C to ensure microtubule depolymerization. Cell debris were removed by centrifugation at 3,300g for 10 min at 4 °C in a swinging bucket rotor (Heraeus Megafuge) followed by an ultracentrifugation at 200,000g for 30 min at 4 °C (Beckman Ti 60). Clarified lysate was subsequently incubated with 1 ml of pre-equilibrated Protein C affinity resin (Roche) for 4 h at 4 °C with recirculation. The column was then washed extensively with 50 ml lysis buffer, then with 50 ml of Klp98A buffer (20 mM HEPES, 150 mM KCL, 2 mM MgCl , 0.1 mM ATP, 10% glycerol, pH 7.2) enriched with 1 mM CaCl , followed by 50 ml Klp98A buffer. Elution was then performed by incubating the 1 ml resin with 1 ml of Klp98A buffer enriched with 5 mM EGTA overnight at 4 °C with rocking. Eluted Klp98A–GFP–PC was then mixed with Klp98A buffer enriched with 2 mM DTT in a 50:50 volume ratio, concentrated by ultrafiltration (Amicon Ultra-4 3k Millipore), and further purified by gel filtration on a Superdex 200 10/300 column (GE Healthcare Life Sciences) in (20 mM HEPES, 0.15 M KCl, 2 mM MgCl , 1 mM DTT, 0.1 mM ATP, pH 7.2) at 0.25 ml min−1. Fractions containing Klp98A–GFP–PC were pooled, mixed with Klp98A buffer containing 20% glycerol final in a 50:50 volume ratio, concentrated by ultrafiltration (Amicon Ultra-4 3k Millipore), flash frozen in liquid N and finally kept at −80 °C (Fig. 1b). Final Klp98A–GFP–PC buffer is (20 mM HEPES, 150 mM KCL, 2 mM MgCl , 0.1 mM ATP, 10% glycerol, 1 mM DTT pH 7.2). For motility assays were a high concentration of Klp98A–GFP–PC was critical to achieve a high density of Klp98–GFP–PC on the quantum dots, the gel filtration step was omitted. Unlabelled porcine tubulin or HiLyte488- and rhodamine-labelled porcine tubulin were purchased from Cytoskeleton, reconstituted at 10 mg ml−1 in BRB80 buffer (80 mM K-Pipes, 1 mM MgCl pH 6.9) supplemented with 1 mM GTP (Roche) or 1 mM GMPPCP (Jena Bioscience), flash frozen in liquid N and kept at −80 °C. GFP–MAP65-1 was a gift from V. Stoppin-Mellet, M. Vantard and J. Gaillard (ref. 13). Fly notum dissection and SOP imaging was performed in clone 8 medium after embedding into a fibrinogen clot48, 49 in order to diminish tissue movements during fast 3D image acquisition as described50. Fluorescent Delta antibody uptake to label the Sara endosomes was performed as previously described1, 50 with a 5-min pulse (3.4 μg ml−1 antibody in clone 8) and a 20-min chase (referred to as iDelta ). To address antibody bleaching, which hampers the accuracy of endosome tracking during acquisition, we replaced the original primary anti-Delta antibody coupled to a fluorescent Fab1, 50 by a primary anti-Delta antibody covalently coupled to the very stable Atto647N dye (see Antibodies). Under these labelling conditions, no bleaching is detectable (Extended Data Fig. 4n). For SiR-tubulin imaging, dissected nota were incubated in clone 8 medium enriched with 1 μM SiR-tubulin15 (Spirochrome) for 30 min at room temperature, then washed twice in clone 8 before fibrinogen clot embedding as above and imaging. Note that SiR-tubulin is less excluded from the Pavarotti-positive central spindle core than Jupiter–mCherry (Fig. 3a). For imaging of Sara endosomes dynamics in toto with neither iDelta uptake nor notum dissection (Extended Data Fig. 2c–h), pupae were mounted as described by Jauffred and Bellaïche51. Drift along the z axis resulting from muscle contractions was corrected by manually adjusting the focus during the acquisition. Compared to the signal in the primary culture preparation upon an antibody uptake, this in toto preparation shows a lower signal-to-noise ratio owing to the glow signal generated by the tissues underneath the epithelium of the epidermis. To address this, and only for visualization purposes, in Extended Data Fig. 2f, g we processed the images with a wavelet à trous filter (ImageJ plugin ‘Kymo Toolbox’ developed by Fabrice Cordelières). Imaging was performed using a 3i Marianas spinning disk confocal setup based on a Zeiss Z1 stand, a 63× PLAN APO NA 1.4 objective and a Yokogawa X1 spinning disk head followed by a 1.2× magnification lens and an Evolve EMCCD camera (Photometrics). Fast z-stack acquisition of entire SOP cells (0.5-μm steps) was obtained using a piezo stage (Mad City Labs). Single-emitter emission filters were always used to avoid bleed-through and each channel was acquired sequentially. To increase acquisition speed for iDelta endosome tracking, we acquired 3D stacks spanning only 3 μm along the z axis (with 0.5-μm steps), which is usually sufficient to contain most of the central spindle (and sufficient to distinguish particles along the z axis, given the PSF of the microscope at this wavelength). In addition, the Pavarotti channel was acquired once every 20 time points. The strong brightness of the Atto647N dye allowed us to perform 3D acquisition at 1.3 Hz on average. Unless stated otherwise, data presented in figure panels correspond to maximum-intensity projections. Dissected fly nota were fixed according to a method designed to preserve the microtubule cytoskeleton52. In brief, nota were first incubated in Hank’s balanced salt solution (Gibco) enriched with 1 mM DSP (Pierce) for 10 min at room temperature followed by a 10 min incubation in MTSB (microtubule stabilization buffer: 0.1 M PIPES, 1 mM EGTA, 4% PEG 8000, pH 6.9) enriched with 1 mM DSP, then finally in MTSB enriched with 4% PFA (Electron Microscopy Science). Nota were then permeabilized in MTSB enriched with 4% PFA and 0.2% Triton X-100 then processed for immunofluorescence as described1 and mounted in Prolong Gold anti-fade reagent (Molecular Probes). Unlabelled and fluorescently labelled (see Antibodies) primary antibodies were used at 1 μg ml−1. When non-labelled primary antibodies were used, we added Alexa647- and Alexa488-coupled secondary antibodies (Life Technologies) at a 1:500 dilution. For lineage staining (Extended Data Fig. 3c), fly nota were dissected 30 h after puparium formation and processed for immunofluorescence as above using primary rat anti-Elav at 22 μg ml−1 antibodies followed by Cy5-coupled secondary antibodies (Biozol) at a 1:100 dilution. For S2 cells immunofluorescence (Extended Data Fig. 6b), cells were plated onto glass coverslips pre-coated with Concanavalin A (Sigma, 0.05 mg ml−1 in water for 1 h) for 1 h at 25 °C in Schneider medium enriched with 10% serum. Cells were then fixed with 4% PFA (Electron Microscopy Science) for 20 min, then processed for immunofluorescence using standard techniques with Oregon-green 514-anti α-tubulin antibodies at 1 μg ml−1 final (see Antibodies). Coverslips were mounted in Prolong Gold anti-fade reagent. Image acquisition was performed on the 3i Spinning disk confocal microscope described above, but using a 100× PLAN APO NA 1.45 TIRF objective and a z step of 0.27 μm for optimal sampling along the z axis. Alternatively, for Extended Data Fig. 3c, images were taken on this setup using a 40× PLAN APO NA 1.3 objective and a Photometrics HQ2 CCD camera. For co-localization studies of iDelta with GFP–Sara (Extended Data Fig. 2a, b) and of Klp98–mCherry with GFP–Sara, iDelta , GFP–Rab5 knock-in and YFP–Rab7 knock-in (Extended Data Fig. 1c–e), dissected fly nota embedded in the fibrinogen clot were fixed using 4% PFA in PEM buffer (80 mM K-Pipes, 5 mM EGTA, 1 mM MgSO , pH 6.95) for 20 min at room temperature, then washed three times with PEM and imaged in PEM. Image acquisition was performed on the 3i Spinning disk confocal microscope described above with the 100× PLAN APO NA 1.45 TIRF objective, a z step of 0.27 μm and both channels were acquired sequentially at each z plane. Cells at various stages of the cell cycle were included into the analysis. Since signal of YFP–Rab11 at endogenous levels (knock-in) was lost upon fixation in our conditions, co-localization between Klp98–mCherry and YFP–Rab11 was addressed in living tissue (acquiring only one z plane, to address fast 3D movements of the endosomes). Polyclonal rabbit anti-Klp98A antibody was generated by injecting rabbits (Eurogentec Speedy program) with cleaved GST–Klp98A-Cter (see Protein purification). Immunized serum was subsequently affinity-purified with sepharose-bound His–Klp98A-Cter using standard glycine (0.1 M, pH 3.0) elution. Eluted antibody was subsequently dialysed against PBS then PBS-50% glycerol for storage at −20 °C. The characterization of this antibody is presented in Extended Data Fig. 1a, b. Polyclonal rabbit anti-Patronin antibody was generated using the same protocol. Its characterization is provided in Extended Data Fig. 6a. Mouse anti-Delta monoclonal antibodies (C594.9B, Developmental Studies Hybridoma Bank) were purified on a Protein G column (Pierce) from hybridoma culture supernatant obtained by cultivating the hybridoma in CELline devices (Integra) using RPMI medium (Gibco) supplemented with 10% ultra-low IgG fetal calf Serum (Gibco) and 1% pen-strep (Gibco). Antibodies were subsequently dialysed against fresh 0.15 mM sodium bicarbonate pH 8.3, concentrated to 4.11 mg ml−1 and labelled with NHS-Atto 647N (Atto tech) in a 5× molar excess of dye for 2 h in the dark at room temperature. Free dye was subsequently removed by gel filtration on a G-25 fine column (Sigma) in PBS. Degree of labelling was measured spectrophotometrically to be 2.6. Oregon Green 514-labelled mouse anti-β-tubulin (E7, Developmental studies hybridoma bank), Oregon Green 514-labelled mouse anti-α-tubulin (12G10, Developmental studies hybridoma bank) and Atto-647N-labelled anti-α K40 acetylated tubulin (C3B9, HPA Cultures) were purified and labelled in a similar fashion. Degree of labelling was measured spectrophotometrically to be 2.7 for Oregon Green 514-labelled anti-β-tubulin, 1 for Oregon Green 514 labelled anti-α-tubulin, and 1.6 for Atto-647N labelled anti-acetylated tubulin. Biotinylated GBP was obtained by in vitro biotinylation of purified GBP (see protein purification, or purchased from Chromoteck) with EZ-Link SulfoNHS biotin (Pierce) in a 1:5 ratio followed by extensive dialysis (SnakeSkin 3kD MWCO, Pierce) against PBS. All labelled antibodies were subsequently frozen in liquid N and kept at −80 °C. Mouse anti-PC (clone HPC4) antibodies were from Roche. Rat anti-Elav (7E8A10) was from Developmental studies hybridoma bank. Unlabelled mouse anti-β-tubulin (E7) was also used for loading controls in western blots. General tubulin handling as well as preparation of GMPPCP-stabilized, Taxol-stabilized and polarity marked fluorescent microtubules were performed accordingly to the protocols of the Mitchison laboratory (http://mitchison.hms.harvard.edu/resources). GTP and GMPPCP microtubules were polymerized at 5 mg ml−1 for 20 min at 37 °C in a water bath. Unpolymerized fluorescent tubulin dimers were removed by ultracentrifugation over a glycerol cushion. Motility assays of Klp98A were performed using purified full length Klp98A–GFP–PC (that is, full length Klp98A fused to GFP and the PC tag in Cter; see Protein purification). Imaging of motility assays were performed using a 3i TIRF microscope based on a Zeiss Z1 stand equipped with a TIRF Slider 3 module. Excitation was performed with a 488 nm laser and simultaneous detection of both microtubules and quantum dots (Qdots) was performed using a Dualview device (Photometrics) equipped with a 565dcxr dichroic (Chroma) and two emission filters (520/30 and 630/50, Chroma) in front of an EMCCD camera (Cascade II 512, Photometrics) at 6.66 Hz. The motility properties of Klp98A-bound Qdots were analysed on kymographs using the ImageJ plugin ‘Kymo Toolbox’ developed by Fabrice Cordelières. This plugin was also used to process images from the Qdot channel with a wavelet à trous filter for representation purposes (Supplementary Video 5). Motility of Klp98A-bound Qdots was analysed as described53 with the following modifications. In brief, glass coverlips (Agar Scientific) were cleaned using a plasma cleaner (Harrick_plasma) and assembled into a flow chamber using sticky slides (sticky-Slide VI 0.4 Luer, Ibidi). This flow chamber was connected to an Aladdin Syringe Pump (World Precision Instrument) used to change gently the solution in the chamber. The chamber was first perfused with anti-tubulin antibodies (SAP4G5, Sigma, 1/100 dilution in BRB80) for 5 min, then passivated using four chamber volumes of 0.1 mg ml−1 PLL-PEG (Susos) in BRB80 for 5 min followed by four chamber volumes of 0.5 mg ml−1 K-Casein (Sigma) in BRB80 for 5 min. A dilute solution of Taxol- or GMPPCP-stabilized microtubules (0.05 mg ml−1, 5% labelled with HiLyte 488) were then injected and let to adhere to the antibodies for 10 min. The chamber was then washed with four chamber volumes of imaging buffer (BRB80 enriched with 0.25 mg ml−1 K-casein, 1 mM ATP, 40 mM DTT, 20 μg ml−1 catalase, 160 μg ml−1 glucose oxydase and 40 mM d-glucose). Klp98A–GFP–PC (3 μM) was pre-incubated with 1.5 μM of biotinylated GBP for 5 min, before mixing in a 10:1 molar ratio with strepavidin-coated Qdots 605 (Molecular Probes). This ensured a high density of motors per Qdot, thus mimicking a bead assay, although bead diameter is small. These Klp98A-bound Qdots were then injected in the flow chamber in imaging buffer. Gliding assays of polarity-marked microtubules (Fig. 1c) were performed using the same flow chamber described above. Polarity-marked microtubules were obtained by elongating short bright GMPPC microtubule seeds (5 mg ml−1, 30% rhodamine labelled) with a dimmer tubulin mix (1.5 mg ml−1, 5% rhodamine labelled) followed by stabilization with 20 μM Taxol. The chamber was first perfused with Klp98A–GFP–PC (2.9 μM) then passivated with PLL-PEG as above. Polarity-marked Taxol-stabilized microtubules were then injected and let to adhere to Klp98A for 5 min. The chamber was then washed with two chamber volumes of imaging buffer enriched with 20 μM Taxol then imaged in the same buffer. As seen in Fig. 1c, the minus-end (short) is leading in these gliding assays, indicating that Klp98A is a plus-end motor. For motility of Klp98A-bound Qdots on antiparallel arrays of microtubules, antiparallel bundles were generated by incubating 50 nM GMPPCP microtubules (5% rhodamine-labelled) with 6.5 nM of GFP–MAP65-113 for 5 min at room temperature. These bundles were injected into the chamber and moving Klp98A-bound Qdots were observed as before using a 561 nm laser to excite rhodamine and a 405 nm laser to excite the Qdots 605. Due to the excess of Klp98A–GFP–PC over GBP–biotin in this assay, it is likely that all available GFP-binding sites of the Qdots are saturated, thus the presence of GFP-tagged MAP65-1 is not an issue. For the analysis of the frequency at which Qdots change direction, we only considered antiparallel overlaps composed of two microtubules. We first identified pauses in the motility of Qdots (a pause is defined by a Qdot immobile for at least three consecutive frames, which corresponds to 0.9 s). Then we scored the incidence of changes of direction after a pause, in order to compute the frequency of direction changes. Liposome flotation assays were performed as described45 with the following modifications. Small unilamellar vesicles (SUVs) were prepared by N. Chiaruttini in BRB80 buffer by sonication in a water bath with several lipid mixtures: DOPC:DOPS 90:10; DOPC:DOPS:PI(3)P 80:10:10; DOPC:DOPS:PI(4)P 80:10:10 and DOPC:DOPS:PI(5)P 80:10:10. All lipid mixtures were doped with 0.6% rhodamine phosphatidylethanolamine (PE). 70 μl of SUVs (1 mg ml−1) were incubated with 5 μl Klp98A–GFP–PC (0.05 mg ml−1) for 30 min at room temperature. Then 50 μl of 2.5 M sucrose in BRB80 was added and gently mixed. 100 μL of this solution was poured into a polyallomer tube (Beckman Coulter), and then overlaid with 100 μL of 0.75 M Sucrose in BRB80 then with 20 μL of BRB80. This discontinuous sucrose gradient was then ultracentrifuged at 100,000 r.p.m. for 20 min in a TLA 100.4 rotor (Beckman Coulter) at 25 °C with acceleration and deceleration settings set to level 5. The top 50 μl of the gradient, referred to as the ‘floating fraction’, was subsequently collected and liposome recovery was quantified by measuring rhodamine fluorescence using a Spectramax I3 plate reader (Molecular Devices). Equal amounts of recovered SUVs were then loaded onto a SDS–PAGE gel followed by western blot against the PC tag to analyse protein co-flotation with the SUVs. As controls, we also loaded samples devoid of liposomes as well as the input before centrifugation (Extended Data Fig. 1f). Flies were euthanized by exposure to diethyl ether for 20 min, then mounted on SEM holders using double-sided carbon tape (Electron Microscopy Sciences) and subsequently treated with a gold sputter coater (JFC-1200, JEOL). Imaging was performed using a JEOL JSM-6510LV scanning electron microscope operating in high-vacuum mode using a working distance of 10 mm and an acceleration of 10 kV. Alternatively, for Extended Data Fig. 3a, imaging was performed using a JEOL 7600F scanning electron microscope using a working distance of 25 mm and an acceleration of 5 kV. Two endocytic factors play major, independent roles during asymmetric Notch signalling in the SOP: Neuralized and Numb (reviewed in ref. 8). In Neuralized mutants, cells in the lineage become neurons and, conversely, in Numb mutants they become sockets. It has previously been shown that Neuralized complete loss of function causes a full conversion of all the SOP lineage into neurons leading to a bald notum cuticle54, 55. However, a partial depletion of Neuralized in the centre of the notum (pnr > neurRNAi) allows many sensory organs to perform asymmetric cell fate assignation and to develop, as in wild type, into structures containing at least the two external cells (shaft and socket; Fig. 1i, j, Extended Data Fig. 3a, b). Klp98A mutants reveal that the lineages which generated bristles in pnr > neurRNAi need Klp98A function to perform asymmetric cell fate assignation: in Klp98A−, pnr > neurRNAi double mutants, these lineages failed to perform asymmetric signalling, causing the notum to be largely bald (Fig. 1i, j, Extended Data Fig. 3a, b). This was confirmed with two independent Klp98A mutants. Conversely, these two different Klp98A mutant conditions in combination with three alternative hypomorphic mutant conditions for Numb (NumbSW/Numb2, NumbSW/Numb15 or pnr-gal4 driving NumbRNAi) all show a strong suppression (by half) of the multiple socket phenotype diagnostic of Numb mutants9 (Extended Data Fig. 3d–f). All together, these experiments demonstrate the role of Klp98A motility in Notch signalling. To quantify these cell-fate phenotypes in the SOP lineage in NeuralizedRNAi mutants (Fig. 1i, j and Extended Data Fig. 3a, b), we manually scored in each genotype the number of organs within the region between the left and right pairs of dorso-central macrochaetes (which corresponds to the panier expression region) at the dissecting scope or on SEM images. To focus on lineage specification phenotypes generated by cell-fate specification failures in the SOP division, we scored lineages which generated organs composed of one-shaft/one-socket or two-shafts. In these organs, the SOP division seems to have been normal and thereby generated a pIIa (and a pIIb cell). ‘Tufts’, which are characteristic of neuralized mutant phenotype, could be caused by SOP specification defects and were therefore excluded from the analysis. We verified that the absence of lineages generating bristles in the pnr > neurRNAi, Klp98AΔ8/Klp98AΔ47 double mutant conditions are not due to an earlier, SOP specification problem. The question is whether, in the double mutant condition, the notum is bald because SOPs were specified and the lineage has all been converted into neurons or, alternatively, whether SOPs were not specified in the first place. Immunostaining with a neural specific marker (elav) confirmed that, below the bald cuticle, clusters of elav-positive neurons are present like in the control animals (Extended Data Fig. 3c). To quantify cell-fate phenotypes in the SOP lineage in Numb mutants (Extended Data Fig. 3d–f), we manually scored on SEM images the number of organs showing multiple sockets (that is, Notch gain-of-function phenotype) in the dorsal-most region of the notum (between the left and right pairs of dorsocentral bristles) both in mutant and control flies and calculated the percentage of affected organs in each genotype. Lifetime imaging of GFP–Patronin was performed on a setup composed of an Olympus IX81 stand, a 60× NA 1.42 oil objective, a FV1000 confocal scanner head and time-correlated single-photon counting (TCSPC) hardware from Picoquant. Illumination was achieved with a pulsed 485 nm laser (Picoquant) operating at 40 Mhz, and detection was performed on a gated PMA hybrid 40 detector (Picoquant) behind a 520/35 nm bandpass filter (Semrock). Data analysis was performed using SymPhotime 2.0 software (Picoquant). GFP fluorescence lifetime was fitted to a dual exponential model after deconvolution for the instrument response function (measured using fluorescein in the presence of saturating potassium iodine). The lifetime reported in images and graphs corresponds to the intensity-weighted average lifetime. To measure the lifetime of GFP, we incubated 10 μl of TALON beads (Clontech) with 37.5 μg of purified His–GFP (see protein purification) in 10 μl clone 8 medium for 3 h at room temperature. After two washes in Clone 8 medium, we mounted the beads on a coverslip in 50 μl clone 8 and measured the intensity-weighted average lifetime in a region of interest (ROI) encompassing each bead by FLIM, followed by averaging over several beads. Similarly, to measure the lifetime of GFP in conditions where 100% of the molecules are bound to the GFP–nanobody (GBP), we incubated 10 μl streptavidin beads (GE healthcare) with 18 μg of biotinylated GBP (see Antibodies) for 10 min at room temperature. After extensive washing of unbound GBP, the resulting GBP-bound beads were incubated with 37.5 μg of purified His–GFP in 10 μl clone 8 medium for 3 h at room temperature. After two washes in clone 8 medium, the lifetime of GBP-bound GFP was measured as above. Alternatively, we used GFP-trap beads from Chromoteck, in which the GBP is directly cross-linked to beads. This gave similar values of increased GFP lifetime: τ = 2.627 ± 0.006 ns; n = 15 for the GBP-biotin/Streptavidin beads versus τ = 2.678 ± 0.004 ns; n = 10 for the GFP-Trap beads (GBP-free GFP has a lifetime of τ = 2.531 ± 0.003 ns; n = 29). Please note that for all FLIM measurements, either of purified GFP in vitro or of GFP–Patronin fusion in the fly, the term GFP refers to the enhanced GFP variant (eGFP). FRAP of GFP–Patronin (Extended Data Fig. 9e) was performed on the 3i Marianas spinning disk setup described above (63× NA 1.4 oil objective) equipped with a Micropoint Photomanipulation hardware driven by Slidebook 6.0. A region of interest (ROI) was drawn onto half of the mitotic spindle, bleached, and recovery was monitored by spinning disk confocal imaging at a frame-rate of 14.3 Hz (50 ms exposure, 20 ms transfer time). Owing to the fast recovery of GFP–Patronin (timescale of few seconds), recovery was monitored in 2D (that is, one z plane) to maximize frame-rate. FRAP movies were processed as follows: signal background was first removed homogenously using a ROI outside the cell as a reference, then, bleaching was corrected homogenously using the first frame as a reference. GFP–Patronin signal within the bleached ROI was then integrated overtime. Intensity was then normalized using the formula: With I(t), the integrated intensity at time point t; I , the intensity just after bleaching, and I the intensity before bleaching (averaged over five time points). Normalized intensity was then fitted to the equation: In this equation, A corresponds to the immobile fraction, the half-time of recovery is provided by and τ is an estimate of the k of GFP–Patronin for mitotic spindle microtubules (assuming that diffusion is faster than binding/unbinding kinetics). Averaging the values of A, t , and k for each curve gave similar results than the values obtained by fitting the average recovery: A = 0.90 ± 0.01, t  = 1.3 ± 0.1 s and k  = 0.53 ± 0.03 s−1, n = 11 for average of the individual fits versus A = 0.89 ± 0.02, t  = 1.31 ± 0.03 s and k  = 0.53 ± 0.01 s−1 for fit of the average curve (95% confidence intervals). Unless otherwise specified, image analysis was performed using custom codes written for ImageJ and Matlab (Mathworks), available on request. For representation purposes, intensity was sometimes colour-coded using the Rainbow or the Red Hot lookup tables in ImageJ. Videos were edited using Adobe Premiere Pro CS6. To automatically measure the co-localization between iDelta and GFP–Sara (Extended Data Fig. 2), as well as the co-localization between Klp98A–mCherry and various early endosome markers (Extended Data Fig. 1c, d), we developed a custom object-based method to determine the percentage of co-localization of signals detected in two different channels. Indeed, the fact the membrane of endosomes is organized as a mosaic of domains56, 57, 58 implies that the corresponding signals only partially overlap, which explain why classical co-localization methods relying on intensity correlation coefficients perform poorly in the case of endosomes. On the other hand, object-based methods rely on the segmentation of the signals in both channels followed by the measurements of the distances between all the objects: two objects are considered co-localized if the distance between their fluorescence centroid is below a certain threshold r (ref. 59). Current endosome segmentation methods rely on an intensity threshold for the fluorescent signal59. This is problematic when the signal intensity in different endosomes is heterogeneous (that is, to take dim endosomes into account, bright endosomes are over-segmented, and vice-versa). To avoid this issue, we adapted to 3D a threshold-free method for endosome segmentation, which is based on Gaussian fitting. In brief, signal-positive particles in both channels are first detected in 2D in each z plane by a 2D Gaussian fitting algorithm60, which does not rely on an intensity threshold, but rather on the fact that particles are characterized by fluorescent signals with a spatial Gaussian distribution with an offset which correspond to the local background. Then, the particles detected in each plane (2D), but corresponding to the same object in 3D, are connected based on the point spread function (PSF) of the microscope. From this, the 3D coordinates of the centroid of fluorescence of all the particle is determined in each channel. Once this automated detection (‘segmentation’) has been performed in the two channels, the distance d between all particles in 3D in the two channels (A and B) are computed and compared to a reference distance r . If d < r , the particles detected in the two channels do co-localize. When considering 2D data, r is routinely set to be the lateral resolution of the microscope resol (ref. 59). However, in 3D, since the axial (resol ) and lateral (resol ) resolutions of the microscope are not equal, the reference distance r has to take into account the relative position of the two particles in 3D. For instance, if the two particles are on the same z plane, then r has to be resol and conversely, if the two particles are on different z planes, but have identical x and y coordinates, then r has to be resol . Following a method implemented by Cordelières and Bolte in the ImageJ plugin JACop 2.0 (ref. 59), we calculated r for the 3D problem using the following equations: Here, x , y , z and x , y , z are the 3D coordinates of particles in channel A and B, respectively, and resol and resol correspond to the lateral and axial resolutions of the microscope, respectively. For our analysis, we measured resol  = 0.9 μm and resol  = 0.32 μm using 0.2-μm TetraSpeck beads from Invitrogen. Once all the particles have been detected and their co-localization state addressed (that is, d < r ), we measured the percentage of co-localization as the fraction of the total signal contained in particles that do co-localize, namely: This measurement was then averaged between cells and compared between genotypes. Similar values of the percentage of co-localization were obtained if the fraction of co-localizing particles rather than the fraction of total intensity was considered (data not shown). Since much of the signal of YFP–Rab11 at endogenous levels is lost upon fixation in our conditions, we measured the co-localization between Klp98–mCherry and YFP–Rab11 in living samples. We thus acquired only single planes and applied the algorithm describe above in 2D, considering r  = resol . All endosome tracks were recorded with a time interval of 12 s between frames. For each endosome track, a mean square displacement (MSD) analysis was performed using the MATLAB plugin MSD Analyser61. In brief, for each endosome track in data sets of different conditions, the MSD of segments of increasing duration (delay time t) was computed to obtain Extended Data Fig. 4a for wild type (103 tracks) and Extended Data Fig. 4b for Klp98A− (158 tracks). The ‘weighted mean’ of all individual MSD traces in each condition was then computed as described61: ; where n is the number of tracks, MSD (t) corresponds to the MSD value of the endosome track i for the delay time t, and w to the number of points averaged to compute MSD (t) (Extended Data Fig. 4c and Extended Data Fig. 4a, b, black curve). Note that the weighted mean gives more weight to MSD curves that have greater certainty. We fitted two fit functions to the measured weighted MSD of endosomes as a function of delay time: (i) motion with an average velocity v and a diffusive component with a diffusion D (diffusion + directed motion), which is captured by ; and (ii) simple diffusion, captured by . While simple diffusion (that is,  ) captures well the motion of Klp98A− endosomes (R2 = 0.999; D = (2.04 ± 0.02)×10−3 μm2 s−1; Extended Data Fig. 4c, 95% confidence interval), it poorly fits the data when considering the motion of wild-type endosomes (R2 = 0.8). This indicates that Klp98A is essential for the directed motility of endosomes beyond diffusion, as seen in wild type. Indeed, the ‘diffusion + directed motion’ fit function (that is, ) fits well the wild-type data (R2 = 0.99; Extended Data Fig. 4c). This fit provides an estimate for v = (5.75 ± 0.12)×10−3 μm s−1, while confirming the diffusion coefficient (D = (1.83 ± 0.13)×10−3 μm2 s−1; 95% confidence interval) observed in Klp98A− conditions. Furthermore, the ‘diffusion + directed motion’ fit function fits the Klp98A− data well (R2 = 0.999) only for very low values of v (v = (0.3 ± 0.5)×10−3 μm s−1; D = (2.11 ± 0.04)×10−3 μm2 s−1; 95% confidence interval), confirming that most of the directed motion of wild-type endosomes is mediated by Klp98A motor function. Since endosomes in Klp98A− mutants display simple diffusion, we used this mutant condition to independently evaluate the diffusion coefficient of endosomes by measuring the variance of the histograms of instantaneous speed and in both x and y dimensions. Indeed simple diffusion along the x axis is described by (ref. 61), where σ is the variance of the instantaneous speed over the x axis and Δ is the frame-rate (here Δ  = 12 s). A corresponding expression applies to the y axis. This provided an estimate of D  = 0.0024 μm2 s−1 (Extended Data Fig. 4d) and D  = 0.0023 μm2 s−1 (Extended Data Fig. 4e), confirming the results of the MSD analysis above. In this work, we used spatio-temporal registration of movies to generate a spatio-temporal endosome density plot during SOP division (Fig. 2d, Extended Data Fig. 7a). We also used this spatio-temporal registration to obtain a density plot of different microtubule markers to study the asymmetry of the spindle (Fig. 3a, b, Extended Data Figs 5, 6 and Supplementary Video 6). In addition, time registration allowed us to average data coming from several video data sets (Figs 1f, 3e and Extended Data Figs 2i, 4x and 7f), but also to compare the timing in different figure panels (for instance Figs 1f, 3e and Extended Data Fig. 2i) Spatial registration was performed by defining the centre of the central spindle as monitored by the Pavarotti fluorescent signal, which is also used to establish a Cartesian system of coordinates with respect to which all the other signals (including endosome tracks and density of microtubule markers) are referred. Time registration capitalizes in the stereotypic dynamics of Pavarotti contraction which allowed us to align the timing of our data set of videos (Extended Data Fig. 4q–s). In figure panels where data sets have been registered in time, we have set registered time = 0 to the onset of anaphase B (that is, when the Pavarotti signal starts to constrict, see Extended Data Fig. 4r). A custom code in ImageJ (available upon request) was generated to segment the Pavarotti signal over time. This allowed us to track the Cartesian reference frame of the central spindle, defined by an origin and two axes (x and y, where the y axis is aligned with the division plane; Fig. 2a, b, Supplementary Video 3). The orientation of the x axis is defined to be anterior to posterior (pIIb to pIIa) and was determined by automatic tracking of the mRFP–Pon signal at the anterior cortex of the SOP. In brief, the 3D stack of confocal slices in the Pavarotti channel (GFP– or mCherry–Pavarotti; 3 μm deep, Δz = 0.5 μm) is projected (maximum-intensity projection), then the Pavarotti-positive region is fitted by an ellipse after semi-automated thresholding. The long axis of the ellipse defines the y axis of the reference frame described above and the short axis, the x axis (see Fig. 2a). The length of the Pavarotti-positive region along each axis is determined by taking the full-width half-maximum (FWHM) of the Pavarotti signal along the two axes. For each time point, five parameters are measured: Pavarotti width (PW, size of the Pavarotti-positive region along the y axis); Pavarotti length (PL, size along the x axis); x and y , the 2D coordinates (with respect to the top/left corner of the image) of the position of the origin C of the central spindle reference frame and α, the angle defined by the x axis of this reference frame and the image horizontal axis (Extended Data Fig. 4f). The anterior to posterior orientation of the x axis was determined by detecting the position of the fluorescence centroid of mRFP–Pon signal after manual thresholding. To evaluate the accuracy of our central spindle tracking method, we applied this tracking code on movies of PFA-fixed fly nota acquired in identical imaging conditions. We calculated the deviation from the mean value of the different parameters (x , y and α) obtained from these movies of fixed material. We considered the FWHM of the histogram of these deviations as estimates for the accuracy of the parameters (Extended Data Fig. 4g, h, i). This analysis gave an estimated accuracy for x , y and α of 49 nm, 52 nm and 2.4°, respectively. Since the temporal profile of the shrinking Pavarotti width (PW) is stereotypic from cell to cell, we used it to register videos in time. For each cell, we plotted the temporal dynamics together with that of a reference cell ( ; Extended Data Fig. 4o). This reference cell video was arbitrarily chosen as one that spanned from anaphase to cytokinesis, the relevant phases for this work. We then determined the time delay τ that needs to be applied to the cell of interest to minimize the difference, in absolute value, between the two Pavarotti temporal profiles , that is, find the τ for which is minimum (Extended Data Fig. 4o, p). We then set the initial time of each movie to be equal to τ thereby registering all the movies into an ‘absolute time frame’. As expected, the registered PW curves collapsed (R2 = 0.93) if plotted all together (Extended Data Fig. 4q–s). Importantly, the registered PL curves (Pavarotti size along the x axis), which were not used in the registration process and is a parameter independent of PW contraction, also collapsed (R2 = 0.8; Extended Data Fig. 4t–v), validating our time registration method. In a fewcases where the Pavarotti signal was not recorded in the video (Fig. 3e, for instance), we used instead the contraction of the Jupiter signal over the y axis as a reference. Since Jupiter is excluded from the region where Pavarotti is (Fig. 3a), the absence of Jupiter (‘Jupiter gap’, defined as a FWHM) can be used as a proxy of the Pavarotti region. Extended Data Fig. 4w shows that the contraction of the Jupiter gap follows that of Pavarotti, thus either marker can be used to register data sets in time. Importantly, the contraction of Pavarotti/Jupiter is unaffected in Patronin depletion, Klp10A depletion and Klp98A mutants (Extended Data Fig. 4x), thus enabling temporal registration of videos acquired in these genetic backgrounds relative to control (Fig. 3e, h and Extended Data Fig. 7f). To generate average videos (Fig. 3a and Extended Data Fig. 5a, b and Supplementary Video 6) the Pavarotti tracking data was used to rotate and translate each image to display them in a common spatial reference frame, the centre of which is the centre of the central spindle and whose x axis is horizontal. In order to minimize rotation artefacts, rotation was performed with bicubic interpolation after image scaling by a factor of 4 (without interpolation). After time registration, frames corresponding to each time point were processed by performing homogenous background subtraction and signal normalization (to the brightest pixel). Finally, spatio-temporally registered videos corresponding to different cells were averaged to generate the ‘average video’. All these operations were performed on z-projected images generated by signal integration over the entire volume of the spindle (sum projection, 12 μm total, Δz = 0.5 μm). Images presented in Fig. 3a correspond to late cytokinesis (∼600 s registered time, see Extended Data Fig. 4r). Images of fixed samples (Extended Data Figs 5d–i, 6d–f) were obtained shortly before abscission, when PW and PL (Pavarotti size along y and x axes) do not change much (registered time > 600 s, Extended Data Fig. 4r) and therefore our time registration method (which relies on PW dynamics) cannot be applied anymore. At this stage, we thus used tubulin or Ac-tubulin stainings that had the characteristic ‘8’ shape pattern of late mitotic spindles. For spatial registration, we capitalized on the fact that late spindles have a well-defined elongated 8 shape, allowing image alignment by cross-correlation with a reference image, as used in structure determination from single-particle electron microscopy data62. All these operations were performed on z-projected images (sum projection, 6 μm total, Δz = 0.27 μm). To generate kymographs of endosome recruitment to the central spindle (Fig. 1g), we used the Pavarotti tracking data to rotate and translate each frame (as above for video averaging, but using maximum-intensity z projection in this case). Then each frame was y-projected onto its horizontal x axis and the y-projected movie was displayed as a kymograph. To measure endosome recruitment to the central spindle (Fig. 1f, Extended Data Figs 2i and 7f), we used the Pavarotti tracking data to measure the iDelta signal in the central spindle region over time. To quantify the iDelta signal, images were z-projected (sum projection) after homogeneous background subtraction using a region of the cell devoid of endosomes. This z projection was then segmented using a constant manual threshold to identify the endosomes and the iDelta intensity signal was integrated within the segmented endosomal regions. The iDelta intensity signal was measured both in the central spindle region and the entire cell including the central spindle. The central spindle region was operationally defined on the x axis as a 2 μm region centred at the centroid of the Pavarotti region. The central-spindle-associated signal was then expressed as a percentage of the total signal present in the cell. The Pavarotti tracking data was also used for precise time registration of these movies. Endosome tracking was performed using a custom Matlab code. In brief, the 3D stack containing the iDelta -Atto647N signal (3 μm deep, Δz = 0.5 μm) was z projected (maximum-intensity projection). Particles were detected using a 2D Gaussian fitting algorithm, then tracked using a modified Vogel algorithm, as previously described60. Tracks were rendered using the ImageJ plugin mTrackJ63. To evaluate the accuracy of our endosome tracking method, we applied this tracking code on movies of PFA-fixed fly nota acquired in identical imaging conditions. As an estimate of average accuracy of their position with respect to the image frame (x, y), we calculated the FWHM of their distribution in this fixed material (Extended Data Fig. 4j–l). This analysis showed a positional accuracy of 57 nm along the x axis and 53 nm along the y axis. As expected, we found that this measured positional accuracy decreases with the signal-to-noise (SNR) ratio of the particle considered (Extended Data Fig. 4m) and we thus excluded from the analysis all the particles displaying a SNR <15. The SNR of a diffraction limited object is defined as , where I is the intensity collected at the brightest pixel of the spot and σ is the standard deviation of the local background64. Importantly, due to the very high photostability of our Atto-647N anti-Delta probe, the SNR ratio of endosomes, and thus their positional accuracy, does not vary significantly over time (Extended Data Fig. 4n). Once we have determined the position of the tracked endosomes with respect to the reference frame of the image, we then expressed these coordinates into the Pavarotti Cartesian frame defined above. We did this in order to refer the motility of the endosomes with respect to the relevant structure: the Pavarotti-positive central spindle. If the endosome has the coordinates in the image reference frame, then corresponds to its coordinates in the central spindle reference frame. The central spindle reference frame is centred at and oriented at an angle α (see above) with respect to the image reference frame (Extended Data Fig. 4f). The coordinates in both reference frames are related by The precision of x′ and y′ thus depends on the relative precision of x, x , y, y and α. The variation of x′ relative to x, x , y, y and α is as follows In equation (1) we have so equation (2) becomes: Since errors are independent, an upper estimate of the accuracy of x′ (worst case scenario) is thus: We considered an experimental data set of x, x , y, y and α from a collection of 263 data points corresponding to endosome tracks close to the Pavarotti centroid, as well as the estimated accuracy by tracking endosomes and central spindles in fixed material described above (dx = 57 nm, dy = 53 nm, dα = 2.4° (0.042 rad), dx  = 49 nm and dy  = 52 nm; Extended Data Fig. 4g–l). Using this data to input into equation (4), we obtained an upper bound for the average accuracy of dx′ = 166 nm in the x axis, the axis relevant to the motility of endosomes on the central spindle microtubules. Note that the bidirectional movements that we observed at the central spindle (Fig. 2e and Extended Data Fig. 4y) are in the micrometre range, which is therefore one order of magnitude larger than the accuracy of our measurements. To generate spatio-temporal endosome density plots from our data set of endosome tracks (Fig. 2d and Extended Data Fig. 7a), we binned the data (time bins = 10 s and space bins = 0.5 μm), counted the number of tracks present in each bin and displayed this information as kymograph-type of image and applied the Red Hot lookup table. For residence time measurements (Extended Data Fig. 7d, e), subsets of 101 tracks for control and 30 for Patronin RNAi (‘high-quality tracks’, see also below) were selected after gap correction by manual inspection, if necessary (see Extended Data Fig. 4y for examples). Tracks were selected (i) to be long enough (200 time points on average, thereby allowing to determine residence time); (ii) to display low motility on the y axis (indicating endosomal motility on the central spindle microtubules; Fig. 2.f); and (iii) to contain at least one bidirectional motility event on the central spindle (that is, side-change event). We defined a side-change event as an event where an endosome is moving from the pIIa to the pIIb side of the spindle (or vice versa), that is, when the x coordinate of the moving endosome changes sign. On average, in our selected data, we observed 9 ± 1 side changes per track, which allow determination of the average residence time on each side of the central spindle. Residence time of endosomes on each side of the spindle was measured as follows. After detection of side-change events, the time spent by endosomes in each side of the spindle between these events was computed. Owing to the 166 nm precision of our tracks within the central spindle frame (see above), we excluded from this analysis the segments of the tracks between x = −83 nm and x = +83 nm, but the result did not qualitatively change if this region is considered in the analysis (data not shown). To measure the velocity of microtubule-based-motility, we manually selected segments within our track data where the orientation of movement in the x axis was occurring prominently in one direction for at least ten time points. These segments are referred to as ‘strides’. For each selected stride, we plotted x position versus time and performed a linear fit to estimate the velocity of the stride. This gave us an estimate of v = 0.173 ± 0.007 μm s−1. To measure the off-rate (k ) of endosomes from microtubules at the central spindle, we first automatically detected, on our central spindle tracks, which segments within the tracks correspond to events of transport on microtubules (‘transport segment’). We performed this track analysis on the subset of 101 control high-quality tracks (see above and Extended Data Fig. 10b for an example). The analysis is based on the study of the properties of each step (the displacement between two frames) and the correlation between successive steps. We operationally defined a transport segment using three criteria. (i) Instantaneous speed in each of the steps in the transport segment must be higher than 0.15 μm s−1. Since the velocity of microtubule-based-motility in vivo is v = 0.173 μm s−1, the diffusion coefficient is low and the frame rate is high (see below), this threshold decreases considerably the probability of incorrectly identifying a step of diffusion as a transport step. Two additional criteria help decreasing further this probability. (ii) Segments must last for at least two consecutive steps (three frames). (iii) The orientation of the movement must be the same for all the steps in a transport segment. These two additional criteria make negligible the probability of incorrectly identifying a diffusion segment (a segment composed only of diffusive steps) as a transport segment. Indeed, the probability that a rare fast step of diffusion is followed by yet another rare fast step in the same orientation is extremely low. We actually estimated by performing stochastic simulations (not shown) that, with our measured value D = 0.0021 ± 0.0001 μm2 s−1 and for the fastest frame rate used (1.4 Hz), the probability of incorrectly identifying a diffusion-segment as a transport-segment is about 1 × 10−3. We found k  = 0.90 ± 0.06 s−1 from exponential fits of the distribution of the duration of transport segments (95% confidence interval; see Extended Data Fig. 10c). To estimate k ρ, we considered the track segments in between transport segments which we defined operationally as diffusion segments. We then found k ρ = 0.05 ± 0.01 s−1 (95% confidence interval; Extended Data Fig. 10d). Note that, since we analyse the tracks regardless of their position within the central spindle, the value of the measured k ρ is an average of values for different microtubule densities (that is, ). Extended Data Fig. 10e shows the distribution of run lengths in the transport-state. To estimate the characteristic run length λ for the transport state, we used the method described by Thorn and Vale (ref. 65). In brief, we determined the cumulative distribution P(x) of the transport run lengths x (that is, the fraction of run lengths shorter than a given run length). We then fitted the observed cumulative distribution P(x) to the corresponding equation for x > x , where x  = 0.4 μm is the lower limit of runs included in the fit (x ≤ x corresponds to short runs, which are not measured with great accuracy and are thereby excluded from the analysis). The characteristic transport run length is λ = 0.31 ± 0.01 μm (R2 = 0.98; 95% confidence interval). The advantage of the Thorn and Vale procedure is that it allows us to fit the data directly without data binning. Indeed, it has been shown that performing the exponential fit directly on the binned run length distribution (like in Extended Data Fig. 10e) yields characteristic run lengths that depend strongly on the size of the bins. iDelta asymmetry was measured at late stages of cytokinesis when all endosomes had departed from the central spindle, and iDelta asymmetry had reached its maximum (∼600 s in registered time, see Fig. 1f and Extended Data Fig. 2i). Asymmetry was measured as follows. Endosomes were first detected by using the 2D Gaussian fitting algorithm described above. For all data sets, the same minimal fluorescence signal above local background was imposed to detect bona fide endosomes. Total intensity was then integrated for each endosomes, with the local background determined by Gaussian fitting subtracted. The pIIa and the pIIb cells were then segmented manually using the Pon channel as a reference. Finally, endosomes were assigned based on their coordinates to the segmented pIIa or the pIIb regions. The total endosomal signal for each daughter cell was subsequently computed. The percentage of iDelta in the pIIa daughter cell was then calculated as: We measured the percentage of iDelta signal in the pIIa daughter cell rather than the ratio of signal between the two cells (pIIa:pIIb) since our automatic detection method sometimes did not detect any particles in one of the two daughter cell, leading to a pIIa:pIIb ratio of 0 or infinity. Importantly, the iDelta asymmetries measured by this method were almost identical to results obtained with our previous method based on a 3D signal integration after manual background subtraction and thresholding1, 2, 50 (data not shown). In addition, the iDelta asymmetry measured by this method was similar if endosome numbers or area were considered instead of endosome intensity (data not shown). For correlative measurements of spindle asymmetry versus iDelta endosome asymmetry, and exploration of conditions where spindle asymmetry is inverted (Fig. 4 and Extended Data Fig. 10), we rather plotted the ratio of iDelta in pIIa, which is calculated as: In this work, we measured spindle asymmetry by two methods: the ‘pseudo-line-scan’ method and the ‘segmentation’ method (illustrated in Extended Data Fig. 5c). Both methods gave similar results in live material (Extended Data Fig. 5c corresponding to the samples displayed in Fig. 3a) and in fixed samples (Extended Data Fig. 5h, i). Unless stated otherwise, the pseudo-line-scan method was used. For measurements of spindle asymmetry on live material (Fig. 3b, e), we first projected z stacks containing the entire central spindle (6 μm depth, Δz = 0.5 μm) using sum-intensity projection. We then segmented the Pavarotti signal as described above (see spatio-temporal registration), which defined x/y axes of the spindle, as well as PW (Fig. 2a). Jupiter–GFP, GFP–Patronin or SiR-tubulin signal intensity was then measured along the x axis upon signal integration over the y axis within a region of interest (ROI) centred on the Pavarotti region centroid. This measurement thus conceptually resembles a line scan along the x axis of the spindle, but a rectangular ROI, rather than a line, is considered (ROI dimensions: 10 μm on the x axis and PW on the y axis). The signal intensity over the x axis determined this way displays two peaks: one in pIIa, one in pIIb, see Fig. 3b and Extended Data Fig. 5c. This reflects the facts that these signals are excluded (at least in part) from the Pavarotti region in the middle of the central spindle (see Fig. 2a). We then measured the value of each peak and subtracted the local background (average background was determined from five pixels adjacent to the spindle). Central spindle asymmetry was computed as the enrichment of the density of the marker in the pIIb relative to the pIIa according to Importantly, results were almost identical if a maximum intensity projection was used instead of a sum-intensity projection, and if microtubule density was measured along a line scan with a 1 pixel width instead of the entire width of the spindle by using the ROI, suggesting that spindle asymmetry is invariant along the y axis (data not shown). For measurement of spindle asymmetry on live material (Fig. 3e), we measured this marker enrichment in pIIb at each time point and subsequently averaged these values between different videos using the time registration method described above. In cases where frame rates were not identical among videos, the spindle asymmetry values were interpolated to the correct frame rate using spline interpolation. The kymograph of Jupiter–GFP depolymerization (Fig. 3f) was generated by plotting the pseudo-line-scan for each time point as a kymograph. We then applied the Red Hot lookup table. For correlative measurements of spindle asymmetry versus iDelta endosome asymmetry, and exploration of conditions where spindle asymmetry is inverted (Fig. 4 and Extended Data Fig. 10), we plotted Δ, the normalized enrichment of microtubule density in the pIIb side, rather than the enrichment on the pIIb. Δ is given by the formula: Note that Δ is symmetrical when pIIb and pIIa are inverted and that −1 ≤ Δ ≤ 1. For images of fixed samples (Extended Data Figs 5d–i, 6d–f and 8f, g), we capitalized on the fact that the spindle asymmetry as a function of time remains approximately constant at late stages of cytokinesis (Fig. 3e) and therefore measurements at those stages are unlikely to be affected by incorrect time registration. We fitted the microtubule marker signal to an ellipse to obtain the x and y axes of the spindle, determined manually (in the absence of Pavarotti signal) the cytokinesis plane and measured the microtubule enrichment in pIIb as described above considering a ROI of dimensions 10 μm on the x axis and 0.812 μm (4 pixels) over the y axis. In this method, we segmented the central spindle by considering an intensity threshold above the cytosolic background and computed the average intensity in the segmented regions in the pIIa and pIIb sides (see Extended Data Fig. 5c). This second methods considers the average density of the complete pool of microtubules at the central spindle. This gave comparable results to the pseudo-line-scan method (Extended Data Fig. 5c, h, i). To measure spindle asymmetry in metaphase (Extended Data Fig. 5n), we first projected z stacks containing the entire metaphase spindle (8.5 μm depth, Δz = 0.5 μm) using maximum-intensity projection. We then drew a line between the two spindle poles, which define the mitotic plane: the plane orthogonal to this line, located in the middle distance between centrosomes. We then measured the total signal in two ROIs of 4.6 μm (along the mitotic plane) × 2.3 μm (along the inter-centrosome line) on each side of the mitotic plane, in the pIIa and pIIb sides. Local background was subtracted by considering an adjacent ROI in the cell outside the spindle and the two ROIs described above. The signal enrichment on the pIIb side was then computed as Importantly, these ROIs do not contain the centrosomes so that spindle asymmetry measurements are not affected by centrosome asymmetry (Extended Data Fig. 5o, p). To measure centrosome asymmetry of different markers throughout mitosis (Extended Data Fig. 9d–h), we first projected z stacks containing the entire centrosome signal (6 μm depth, Δz = 0.5 μm) using maximum-intensity projection. We then measured the intensity of each centrosome by considering a circular ROI centred on the centrosome (1.4 μm diameter). Local background was subtracted by considering an adjacent ROI of identical diameter. We then calculated the ratio between the pIIa and the pIIb centrosome intensities. For prophase and prometaphase, the pIIa/pIIb centrosome identity could not be assigned since spindle rotates during metaphase. We therefore measured the ratio of the brighter centrosome over the dimer. To compare Jupiter–GFP intensity between different videos (Fig. 3g,h), a reference intensity was needed to account for the variations of the Jupiter–GFP signal, which occurs even in identical imaging conditions and with expression of Jupiter–GFP at endogenous levels, probably owing to different imaging depths into the tissue. We decided to use the intensity of the centrosome of the pIIa daughter cell, which Jupiter labels throughout the cell cycle (Fig. 3d and Supplementary Video 7) as a reference. We measured the intensity of the pIIa centrosome by considering a circular ROI centred on the centrosome (1.2 μm diameter) and integrating the signal intensity within the ROI on ten z planes (5 μm total depth). Local background was subtracted by considering an adjacent ROI of identical diameter for each plane. We then measured the Jupiter–GFP signal in both the pIIa and the pIIb daughter cells by using the same circular ROI dimensions and background subtraction as above. We then normalized the obtained signal intensity by the pIIa centrosome value. Interestingly, the centrosome of the pIIa daugther cell is 1.41 ± 0.06 (mean ± s.e.m.; n = 26 cells) times more intense than the one of the pIIb daughter at the late cytokinesis stage considered here (Fig. 3d, Extended Data Fig. 5o, p, Supplementary Video 7). Importantly, this difference is still present in Patronin RNAi (1.39 ± 0.17, n = 24) or Klp10A RNAi (1.25 ± 0.08, n = 23; Extended Data Fig. 5o, p) conditions, although the values of the normalized central spindle intensities are different from wild-type conditions (Fig. 3h), suggesting that using the pIIa centrosome is indeed a good way to normalize the Jupiter–GFP data. The fact that Patronin RNAi does not affect microtubule density around the centrosome is in agreement with a recent report showing that CAMSAP2, a mammalian orthologue of Patronin, does not act on astral microtubules28. Unless stated otherwise, measurements are given in mean ± s.e.m. Fit values (MSD analysis, Extended Data Fig. 9e and 10c–e), are provided with their 95% confidence interval. The experiments were not randomized and the investigators were not blinded to allocation during experiments and outcome assessment. No statistical methods were used to predetermine sample size. All statistical analyses were performed using SigmaStat 3.5 software (Systat) with an α of 0.05. Normality of variables was verified with Kolmogorov–Smirnov tests. Homoscedasticity of variables was always verified when conducting parametric tests. For Fig. 3h, a log transformation was applied to the data. In the case were variables failed normality and/or homoscedasticity tests, non-parametric tests were applied. In the main figures, we used Dunn’s post hoc test when performing Kruskal–Wallis tests (Fig. 1h, i) and Tukey’s post hoc test when performing ANOVA (Figs 3h and 4b). Post hoc tests used in Extended Data figures are indicated in their respective figure legends.

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