Jena, Germany
Jena, Germany

Time filter

Source Type

Licht V.,Leibniz Institute for Age Research | Licht V.,Friedrich - Schiller University of Jena | Noack K.,Friedrich - Schiller University of Jena | Noack K.,Jena University Hospital | And 6 more authors.
Oncotarget | Year: 2014

Signal Transducer and Activator of Transcription-1 (STAT1) is phosphorylated upon interferon (IFN) stimulation, which can restrict cell proliferation and survival. Nevertheless, in some cancers STAT1 can act in an anti-apoptotic manner. Moreover, certain malignancies are characterized by the overexpression and constitutive activation of STAT1. Here, we demonstrate that the treatment of transformed hematopoietic cells with epigenetic drugs belonging to the class of histone deacetylase inhibitors (HDACi) leads to the cleavage of STAT1 at multiple sites by caspase-3 and caspase-6. This process does not occur in solid tumor cells, normal hematopoietic cells, and leukemic cells that underwent granulocytic or monocytic differentiation. STAT1 cleavage was studied under cell free conditions with purified STAT1 and a set of candidate caspases as well as with mass spectrometry. These assays indicate that unmodified STAT1 is cleaved at multiple sites by caspase-3 and caspase-6. Our study shows that STAT1 is targeted by caspases in malignant undifferentiated hematopoietic cells. This observation may provide an explanation for the selective toxicity of HDACi against rapidly proliferating leukemic cells.


Keller A.-A.,Friedrich - Schiller University of Jena | Mussbach F.,Friedrich - Schiller University of Jena | Breitling R.,Jena Bioscience GmbH | Hemmerich P.,Leibniz Institute for Age Research | And 3 more authors.
Pharmaceuticals | Year: 2013

Modulating signaling pathways for research and therapy requires either suppression or expression of selected genes or internalization of proteins such as enzymes, antibodies, nucleotide binding proteins or substrates including nucleoside phosphates and enzyme inhibitors. Peptides, proteins and nucleotides are transported by fusing or conjugating them to cell penetrating peptides or by formation of non-covalent complexes. The latter is often preferred because of easy handling, uptake efficiency and auto-release of cargo into the live cell. In our studies complexes are formed with labeled or readily detectable cargoes for qualitative and quantitative estimation of their internalization. Properties and behavior of adhesion and suspension vertebrate cells as well as the protozoa Leishmania tarentolae are investigated with respect to proteolytic activity, uptake efficiency, intracellular localization and cytotoxicity. Our results show that peptide stability to membrane-bound, secreted or intracellular proteases varies between different CPPs and that the suitability of individual CPPs for a particular cargo in complex formation by non-covalent interactions requires detailed studies. Cells vary in their sensitivity to increasing concentrations of CPPs. Thus, most cells can be efficiently transduced with peptides, proteins and nucleotides with intracellular concentrations in the low micromole range. For each cargo, cell type and CPP the optimal conditions must be determined separately. © 2013 by the authors; licensee MDPI, Basel, Switzerland.


News Article | December 9, 2015
Site: www.nature.com

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.


No statistical methods were used to predetermine sample size. The experiments were not randomized and the investigators were not blinded to allocation during experiments and outcome assessment. The deletion allele Ime4∆22-3 was obtained from imprecise excision of the transposon P{SUPor-P}KrT95D and mapped by primers 5933 F1 (CTCGCTCTATTTCTCTTCAGCACTCG) and 5933 R9 (CCTCCGCAACGATCACATCGCAATCGAG). To obtain a viable line of Ime4null, the genetic background was cleaned by out-crossing to Df(3R)Exel6197. Flight ability was scored as the number of flies capable of flying out of a Petri dish within 30 s for groups of 15–20 flies for indicated genotypes. Viability was calculated from the numbers of females compared to males of the correct genotype and statistical significance was determined by a χ2 test (GraphPad Prism). Unfertilized eggs were generated by expressing sex-peptide in virgin females as described30. The genomic rescue construct was retrieved by recombineering (Genebridges) from BAC clone CH321-79E18 by first cloning homology arms with SpeI and Acc65I into pUC3GLA separated by an EcoRV site for linearization (CTCCGCCGCCGGAACCGCCGCCTCCTCCGCCACTTTGCAGGTTGAGCGGACCGCCTCCA GGGCCGCTGCCGCCGGTGCCGCTGATATCCCAGCATGGTAGCTGCGGCCACTCCTAGTC CCGCCTTTAACCACAGCTTGGGGTCCTCCGTCATCAGGCCGAATTGCCTCGAG). An HA-tag was then fused to the end of the ORF using two PCR amplicons and SacI and XhoI restriction sites. This construct was the inserted into PBac{y+-attB-3B}VK00002 at 76A as described31. The Ime4 UAS construct was generated by cloning the ORF from fly cDNA into a modified pUAST with primers Adh dMT-A70 F1 EI (GCAGAATTCGAGATCtAAAGAGCCTGCTAAAGCAAAAAAGAAGTCACCATGGCAGATGC GTGGGACATAAAATCAC) and dMT-A70 HA R1 Spe (GGTAACTAGTCTTTTGTATTCCATTGATCGACGCCGCATTGG) by adding a translation initiation site from the Adh gene and two copies of an HA tag to the end of the ORF. This construct was then also inserted into PBac{y+-attB-3B}VK00002 at 76A. For transient transfection in S2 cells, YT52B-1 and CG6422 ORFs were amplified from fly cDNA by a combination of nested and fusion PCR incorporating a translation initiation site from the Adh gene using primers CG6422 Adh F1 (GCCTGCTAAAGCAAAAAAGAAGTCACCACATGTCAGGCGTGGATCAGATGAAAATACCAG), pACT Adh CG6422 F1 (CCAGAGACCCCGGATCCAGATATCAAAGAGCCTGCTAAAGCAAAAAAGAAGTCACCAC), CG6422 Adh R1, (GATTCCTGCGAACAGGTCCCGTGGGCGAAAC) and CG6422 3′ F1 (CCCACGGGACCTGTTCGCAGGAATCTAG), CG6422 3′ R1 (CATTGCTTCGCATTTTATCCTTGTCCGTGTCCTTAAAGCGCACGCCGATTTTAATTTG), pACT CG6422 3×HA R1 (GTGGAGATCCATGGTGGCGGAGCTCGAGGAATATTCATTGCTTCGCATTTTATCCTTGTC) for CG6422 and primers YT521 Adh F1, (AAGCAAAAAAGAAGTCACATGCCAAGAGCAGCCCGTAAACAAACGCTGCCGATGCGCGAG), pACT Adh YT521 F1 (CCAGAGACCCCGGATCCAGATATCAAAGAGCCTGCTAAAGCAAAAAAGAAGTCACATGCC), YT521 Adh R1 (TGCCATCCGGGCGAATCCTGCAAATTTACCACTCTCGTTGACCGAGAAAATGAGCAGGAC) and YT521 3′ F1(GCAGGATTCGCCCGGATGGCAGCCCCCTCAC), pact YT521 R1 (GGTGGAGATCCATGGTGGCGGAGCTCGAGCGCCTGTTGTCCCGATAGCTTCGCTG) for YT521-B, and cloned into a modified pACT using Gibson Assembly (NEB) also incorporating HA epitope tags at the C terminus. Constructs were verified by Sanger sequencing. The Sxl-HA expression vector was a gift from N. Perrimon32. The YT521-B UAS construct was generated by sub-cloning the ORF from the pACT vector into a modified pUAST with primers YT521 Adh F1 (AAGCAAAAAAGAAGTCACATGCCAAGAGCAGCCCGTAAACAAACGCTGCCGATGCGCGAG), YT521 Adh F2 (TAGGGAATTGGGAATTCGAGATCTAAAGAGCCTGCTAAAGCAAAAAAGAAGTCACATGCC) and YT521 3′ R1 (GGGCACGTCGTAGGGGTACAGACTAGTCTCGAGGCGCCTGTTGTCCCGATAGCTTCGCTG) by adding a translation initiation site from the Adh gene and two copies of an HA tag to the end of the ORF. This construct was then also inserted into PBac{y+-attB-3B}VK00002 at 76A. Essential parts of all DNA constructs were sequence-verified. S2 cells (ATCC) were cultured in Insect Express medium (Lonza) with 10% heat-inactivated FBS and 1% penicillin/streptomycin. The Drosophila S2 cell line was verified to be male by analysing Sxl alternative splicing using species-specific primers Sxl F2 (ATGTACGGCAACAATAATCCGGGTAG) and Sxl R2 (CATTGTAACCACGACGCGACGATG) to confirm species and gender (Extended Data Fig. 8). Transient transfections were done with Mirus Reagent (Bioline) according to the manufacturer’s instruction and cells were assayed 48 h after transfection for protein expression or RNA binding of expressed proteins. To adhere S2 cells to a solid support, Concanavalin A (Sigma) coated glass slides (in 0.5 mg ml−1) were added 1 day before transfection, and cells were stained 48 h after transfection with antibodies as described. Transfections and follow up experiments were repeated at least once. Total RNA was extracted using Tri-reagent (SIGMA) and reverse transcription was done with Superscript II (Invitrogen) according to the manufacturer’s instructions using an oligodT17V primer. PCR for Sxl, tra, msl2 and ewg was done for 30 cycles with 1 μl of cDNA with primers Sxl F2, Sxl R2 or Sxl NP R3 (GAGAATGGGACATCCCAAATCCACG), Sxl M F1 (GCCCAGAAAGAAGCAGCCACCATTATCAC), Sxl M R1 (GCGTTTCGTTGGCGAGGAGACCATGGG), Tra FOR (GGATGCCGACAGCAGTGGAAC), Tra REV (GATCTGGAGCGAGTGCGTCTG), Msl-2 F1 (CACTGCGGTCACACTGGCTTCGCTCAG), Msl-2 R1 (CTCCTGGGCTAGTTACCTGCAATTCCTC), Ewg 4F and Ewg 5R and quantified with ImageQuant (BioRad)22. Experiments included at least three biological replicates. For qPCR, reverse transcription was carried out on input and pull-down samples spiked with yeast RNA using ProtoScript II reverse transcriptase and random nanomers (NEB). Quantitative PCR was carried out using 2× SensiMix Plus SYBR Low ROX master mix (Quantace) using normalizer primers ACT1 F1 (TTACGTCGCCTTGGACTTCG) and ACT1 R1 (TACCGGCAGATTCCAAACCC) and for Sxl, Sxl ZB F1 (CACCACAATGGCAGCAGTAG) and Sxl ZB R1 (GGGGTTGCTGTTTGTTGAGT). Samples were run in triplicate for technical repeats and duplicate for biological repeats. Relative enrichment levels were determined by comparison with yeast ACT1, using the method33. For immunoprecipitations of Sxl RNA bound to Sxl or YTH proteins, S2 cells were fixed in PBS containing 1% formaldehyde for 15 min, quenched in 100 mM glycine and disrupted in IP-Buffer (150 mM NaCl, 50 mM Tris–HCL, pH 7.5, 1% NP-40, 5% glycerol). After IP with anti-HA beads (Sigma) for 2 h in the presence of Complete Protein Inhibitor (Roche) and 40 U RNase inhibitors (Roche), IP precipitates were processed for Sxl RT–PCR using gene-specifc RT primer SP NP2 (CATTCCGGATGGCAGAGAATGGGAC) and PCR primers Sxl NP intF (GAGGGTCAGTCTAAGTTATATTCG) and Sxl NP R3 as described31. Western blots were done as described using rat anti-HA (1:50, clone 3F10, Roche) and HRP-coupled secondary goat anti-rat antibodies (Molecular Probes)34. All experiments were repeated at least once from biological samples. Poly(A) mRNA from at least two rounds of oligo dT selection was prepared according to the manufacturer (Promega). For each sample, 10–50 ng of mRNA was digested with 1 μl of Ribonuclease T1 (1,000 U μl−1; Fermentas) in a final volume of 10 μl in polynucleotide kinase buffer (PNK, NEB) for 1 h at 37 °C. The 5′ end of the T1-digested mRNA fragments were then labelled using 10 U T4 PNK (NEB) and 1 μl [γ-32P]-ATP (6,000 Ci mmol−1; Perkin-Elmer). The labelled RNA was precipitated, resuspended in 10 μl of 50 mM sodium acetate buffer (pH 5.5), and digested with P1 nuclease (Sigma-Aldrich) for 1 h at 37 °C. Two microlitres of each sample was loaded on cellulose TLC plates (20 × 20 cm; Fluka) and run in a solvent system of isobutyric acid: 0.5 M NH OH (5:3, v/v), as the first dimension, and isopropanol:HCl:water (70:15:15, v/v/v), as the second dimension. TLCs were repeated from biological replicates. The identification of the nucleotide spots was carried out using m6A-containing synthetic RNA. Quantification of 32P was done by scintillation counting (Packard Tri-Carb 2300TR). For the quantification of spot intensities on TLCs or gels, a storage phosphor screen (K-Screen; Kodak) and Molecular Imager FX in combination with QuantityOne software (BioRad) were used. For immunoprecipitation of m6A mRNA, poly(A) mRNA was digested with RNase T1 and 5′ labelled. The volume was then increased to 500 μl with IP buffer (150 mM NaCl, 50 mM Tris–HCL, pH 7.5, 0.05% NP-40). IPs were then done with 2 μl of affinity-purified polyclonal rabbit m6A antibody (Synaptic Systems) and protein A/G beads (SantaCruz). Whole-fly extracts were prepared from 20–30 adult Drosophila previously frozen in liquid N and ground into fine powder in liquid N . Cells were then lysed in 0.5 ml lysis buffer (0.3 M NaCl, 15 mM MgCl , 15 mM Tris-HCl pH 7.5, cycloheximide 100 μg ml−1, heparin (sodium salt) 1 mg ml−1, 1% Triton X-100). Lysates were loaded on 12 ml sucrose gradients and spun for 2 h at 38,000 r.p.m. at 4 °C. After the gradient centrifugation 1-ml fractions were collected and precipitated in equal volume of isopropanol. After several washes with 80% ethanol the samples were resuspended in water and processed. Experiments were done in duplicate. Drosophila nuclear extracts were prepared from Kc cells as described35. Templates for in vitro transcripts were amplified from genomic DNA using the primers listed below and in vitro transcribed with T7 polymerase in the presence of [α-32P]-ATP. DNA templates and free nucleotides were removed by DNase I digestion and Probequant G-50 spin columns (GE Healthcare), respectively. Markers were generated by using in vitro transcripts with or without m6ATP (Jena Bioscience), which were then digested with RNase T1, kinased with PNK in the presence of [γ-32P]-ATP. After phenol extraction and ethanol precipitation, transcripts were digested to single nucleotides with P1 nuclease as above. For in vitro methylation, transcripts (0.5–1 × 106 c.p.m.) were incubated for 45 min at 27 °C in 10 μl containing 20 mM potassium glutamate, 2 mM MgCl , 1 mM DTT, 1 mM ATP, 0.5 mM S-adenosylmethionine disulfate tosylate (Abcam), 7.5% PEG 8000, 20 U RNase protector (Roche) and 40% nuclear extract. After phenol extraction and ethanol precipitation, transcripts were digested to single nucleotides with P1 nuclease as above, and then separated on cellulose F TLC plates (Merck) in 70% ethanol, previously soaked in 0.4 M MgSO and dried36. In vitro methylation assays were done from biological replicates at least in duplicates. Primers to amplify parts of the Sxl alternatively spliced intron from genomic DNA for in vitro transcription with T7 polymerase were Sxl A T7 F (GGAGCTAATACGACTCACTATAGGGAGAGGATATGTACGGCAACAATAATCCGGGTAG) and Sxl A R (CGCAGACGACGATCAGCTGATTCAAAGTGAAAG), Sxl B T7 F (GGAGCTAATACGACTCACTATAGGGAGAGCGCTCGCATTTATCCCACAGTCGCAC) and Sxl B R (GGGTGCCCTCTGTGGCTGCTCTGTTTAC), Sxl C T7 F (GGAGCTAATACGACTCACTATAGGGGTCGTATAATTTATGGCACATTATTCAG) and Sxl C R (GGGAGTTTTGGTTCTTGTTTATGAGTTGGGTG), Sxl D T7 F (GGAGCTAATACGACTCACTATAGGGAGAAAACTTCCAGCCCACACAACACACAC) and Sxl D R (GCATATCATATTCGGTTCATACATTTAGGTCTAAG), Sxl E T7 F (GGAGCTAATACGACTCACTATAGGGAGAGGGGAAGCAGCTCGTTGTAAAATAC) and Sxl E R (GATGTGACGATTTTGCAGTTTCTCGACG), Sxl F T7 F (GGAGCTAATACGACTCACTATAGGGAGAGGGGGATCGTTTTGAGGGTCAGTCTAAG) and Sxl NP2, Sxl C T7 F and Sxl C1 R (GTAGTTTTGCTCGGCATTTTATGACCTTGAGC), Sxl C2 F (GGAGCTAATACGACTCACTATAGGGAGACTCTCATTCTCTATATCCCTGTGCTGACC) and Sxl C2 R (CTAATTTCGTGAGCTTGATTTCATTTTGCACAG), Sxl C3 F (GGAGCTAATACGACTCACTATAGGGAGACTGTGCAAAATGAAATCAAGCTCACGAAATTAG) and Sxl C R, Sxl E T7 F and Sxl E1 R (AAAAAAATCAAAAAAATAATCACTTTTGGCACTTTTTCATCAC), Sxl E2 F (GGAGCTAATACGACTCACTATAGGGAGATGAAAAAGTGCCAAAAGTGATTATTTTTTTG), Sxl E2 R (AAAAGCATGATGTATTTTTTTTTTTTTGTACTTTCGAATCACCG), Sxl E3 F (GGAGCTAATACGACTCACTATAGGGAGACGGTGATTCGAAAGTACAAAAAAAAAAAAATAC) and Sxl E R, Sxl C4 F (GAGCTAATACGACTCACTATAGGGAGAAATACTAAAACATCAAACCGCAAGCAGAGCAGC) and Sxl C4 R (GAGTGCCACTTCAAAATCTCAGATATGC), Sxl C5 F (CTAATACGACTCACTATAGGGAGACTCTTTTTTTTTTTCTTTTTTTTACTGTGCAAAATG) and Sxl C5 R (AAAAAAATATGCAAAAAAAAAAAGGTAGGGCACAAAGTTCTCAATTAC), Sxl C6 F (GAGCTAATACGACTCACTATAGGGAGACTGTGCAAAATGAAATCAAGCTCACGAAATTAG) and Sxl C6 R (CAATTTCACTATATGTACGAAAACAAAAGTGAG), Sxl E4 F (GGAGCTAATACGACTCACTATAGGGAGAACCAAAATTCGACGTGGGAAGAAAC) and Sxl E4 R (TAATCACTTTTGGCACTTTTTCATCACATTAAC), Sxl E5 F (GGCTAATACGACTCACTATAGGGAGATTTTTTTGATTTTTTTAAAGTGAAAATGTGCTCC) and Sxl E5 R (CACCGAAAAAAAATAAAAAAAAATAATCATGGGACTATACTAG), Sxl E6 F (GGCTAATACGACTCACTATAGGGAGACTTAAGTGCCAATATTTAAAGTGAAACCAATTG) and Sxl E6 R (CCCCCAGTTATATTCAACCGTGAAATTCTGC). Total RNA was extracted from 15 pulverized head/thoraces previously flash-frozen in liquid nitrogen, using TRIzol reagent from white (w) control and w;Ime4∆22-3 females that have been outcrossed for several generations to w;Df(3R)Exel6197 to equilibrate genetic background. Total RNA was treated with DNase I (Ambion) and stranded libraries for Illumina sequencing were prepared after poly(A) selection from total RNA (1 μg) with the TruSeq stranded mRNA kit (Illumina) using random primers for reverse transcription according to the manufacturer’s instructions. Pooled indexed libraries were sequenced on an Illumina HiSeq2500 to yield 40–46 million paired-end 100 bp reads, and in a second experiment 14–19 million single-end 125-bp reads for three controls and mutants each. After demultiplexing, sequence reads were aligned to the Drosophila genome (dmel-r6.02) using Tophat2.0.6 (ref. 37). Differential gene expression was determined by Cufflinks-Cuffdiff and the FDR-correction for multiple tests to raw P values with q < 0.05 considered significant38. alternative splicing was analysed by SPANKI39 and validated for selected genes based on length differences detectable on agarose gels. Illumina sequencing, differential gene expression and alternative splicing analysis was done by Fasteris (Switzerland). For dosage compensation analysis, differential expression analysis of X-linked genes versus autosomal genes in Ime4null mutant was done by filtering Cuffdiff data by a P value expression difference significance of P < 0.05, which corresponds to a false discovery rate of 0.167 to detect subtle differences in expression consistent with dosage compensation. Visualization of sequence reads on gene models and splice junctions reads in Sashimi plots was done using Integrated Genome Viewer40. For validation of alternative splicing by RT–PCR as described above, the following primers were used: Gprk2 F1 (CCAACCAGCCGAAACTCACAGTGAAGC) and Gprk2 R1 (CAGGGTCTCGGTTTCAGACACAGGCGTC), fl(2)d F1 (GCAGCAAACGAGAAATCAGCTCGCAGCGCAG) and fl(2)d R1 (CACATAGTCCTGGAATTCTTGCTCCTTG), A2bp1 F3 (CTGTGGGGCTCAGGGGCATTTTTCCTTCCTC) and A2bp1 R1 (CTCCTCTCCCGTGTGTCTTGCCACTCAAC), cv.-c F1 (GGGTTTCCACCTCGACCGGGAAAAGTCG) and cv.-c R1 (GCGTTTGCGGTTGCTGCTCGCGAAGAGAG), CG8312 F1 (GCGCGTGGCCTCCTTCTTATCGGCAGTC) and CG8312 R1 (GCGTGGCCACTATAAAGTCCACCTCATC), Chas F2 (CCGATTCGATTCGATTCGATCCTCTCTTC) and Chas R1 (GTCGGTGTCCTCGGTGGTGTTGGTGGAG). GO enrichment analysis was done with FlyMine. For the analysis of uATGs, an R script was used to count the uATGs in 5′ UTRs in all ENSEMBL isoforms of those genes which are differentially spliced in Ime4 mutants, that were then compared to the mean number of ATGs in all Drosophila ENSEMBL 5′ UTRs using a t-test. Gene expression data were obtained from flybase. >pattern <-”atg” # the pattern to look for >dict <-PDict(pattern, max.mismatch = 0)#make a dictionary of the pattern to look for >seq <- DNAStringSet(unlist(fasta_file)[1:638])#make the DNAstringset from the DNAsequences that is, all 638 UTRs related to the 156 genes identified in spanki >result <-vcountPDict(dict,seq)#count the pattern in each of the sequences >pattern <-”atg” # the pattern to look for >dict <-PDict(pattern, max.mismatch = 0)#make a dictionary of the pattern to look for >seq <- DNAStringSet(unlist(fasta_file)[1:29822])#make the DNAstringset from the DNAsequences that is, all UTRs >result <-vcountPDict(dict,seq)#count the pattern in each of the sequences Ime4 or YT521-B were expressed in salivary glands with C155-GAL4 from a UAS transgene. Larvae were grown at 18 °C under non-crowded conditions. Salivary glands were dissected in PBS containing 4% formaldehyde and 1% Triton X-100, and fixed for 5 min, and then for another 2 min in 50% acetic acid containing 4% formaldehyde, before placing them in lactoacetic acid (lactic acid:water:acetic acid, 1:2:3). Chromosomes were then spread under a siliconized cover slip and the cover slip removed after freezing. Chromosome were blocked in PBT containing 0.2% BSA and 5% goat serum and sequentially incubated with primary antibodies (mouse anti-PolII H5, 1:1000, Abcam, or rabbit anti-histone H4, 1:200, Santa-Cruz, and rat anti-HA monoclonal antibody 3F10, 1:50, Roche) followed by incubation with Alexa488- and/or Alexa647-coupled secondary antibodies (Molecular Probes) including DAPI (1 μg ml−1, Sigma). RNase A treatment (4 and 200 μg ml−1) was done before fixation for 5 min. Ovaries were analysed as previously described41. The YTH domain (amino acids 207–423) was PCR-amplified with oligos YTHdom F1 (CAGGGGCCCCTGTCGACTAGTCCCGGGAATGGTGGCGGCAACGGCCG) and R1 (CACGATGAATTGCGGCCGCTCTAGATTACTTGTAGATCACGTGTATACCTTTTTCTCGC) and cloned with Gibson assembly (NEB) into a modified pGEX expression vector to express a GST-tagged fusion protein. The YTH domain was cleaved while GST was bound to beads using Precession protease. Electrophoretic mobility shift assays and UV cross-linking assays were performed as described35, 42. Quantification was done using ImageQuant (BioRad) by measuring free RNA substrate to calculate bound RNA from input. All binding assays were done at least in triplicates. RNA-seq data that support the findings of this study have been deposited at GEO under the accession number GSE79000, combining the single-end (GSE78999) and paired-end (GSE78992) experiments. All other data generated or analysed during this study are included in this published article and its Supplementary Information.


Wild-type male C57BL/6 mice and B6.129S4–PDGFRαtm11(EGFP)Sor/J mice (Jackson strain number 007669), which contain an H2B–eGFP fusion protein knocked into the Pdgfra locus, were obtained from Jackson Laboratories. Young adult mice were 6–8 weeks of age; aged mice were 22–24 months of age. Mice were housed and maintained in the Veterinary Medical Unit at the Veterans Affairs Palo Alto Health Care System. Animal protocols were performed in accordance with the policies of the Administrative Panel on Laboratory Animal Care of Stanford University. Mice were anaesthetized using isoflurane. To assess muscle regeneration, 50 μl of a 1.2% barium chloride (BaCl ) solution (Sigma-Aldrich) was injected into tibialis anterior muscles as described previously5. To isolate activated FAPs for western blot analysis and FACS analysis, 50 μl of 1.2% BaCl or 50% (v/v) glycerol/water was injected throughout the lower hindlimb muscles. For induction of fibrosis, 30 μl of 50% (v/v) glycerol or 30 μl 1.2% BaCl solution was injected into tibialis anterior muscles. Muscles were dissected from mice and dissociated mechanically. All hindlimb muscles were used except in experiments where FAPs were isolated from VMOs injected into tibialis anterior muscles. In this case, only the tibialis anterior muscle was dissected. The muscle suspension was digested using collagenase II (760 U ml−1; Worthington Biochemical Corporation) in Ham’s F10 medium (Invitrogen) with 10% horse serum (Invitrogen) for 90 min at 37 °C with agitation. The suspension was then washed and digested in collagenase II (152 U ml−1; Worthington Biochemical Corporation) and dispase (2 U ml−1; Invitrogen) for 30 min at 37 °C with agitation. The resultant mononuclear cells were then stained with the following antibodies: VCAM-1-biotin (clone 429; BioLegend, 105704), CD31-APC (clone MEC 13.3; BioLegend, 102510), CD45-APC (clone 30-F11; BioLegend, 103112) and Sca-1-Pacific Blue (clone D7; BioLegend, 108120) at 1:75. Streptavidin-PE-Cy7 (BioLegend, 405206) at 1:75 was used to amplify the VCAM-1 signal. FAPs were collected according to the following sorting criteria: CD31−CD45−Sca-1+. FACS was performed using BD-FACS Aria II and BD-FACS Aria III cell sorters equipped with 488 nm, 633 nm and 405 nm lasers. The cell sorters were carefully optimized for purity and viability and sorted cells were subjected to FACS analysis immediately after sorting to confirm FAP purity. FAPs were isolated from uninjured C57BL/6 mice as described above and lysed. RNA was prepared with the RNeasy Mini Kit as per the manufacturer’s instructions (Qiagen). A 3′ blocking reaction was performed using a poly(A) tailing kit (Ambion) and 3′-dATP (Jena Bioscience) and the reaction mixture was incubated at 37 °C for 30 min. RNA was hybridized to flow cell surfaces for direct RNA sequencing as previously described18. Raw direct RNA sequencing reads were filtered using the Helicos-developed pipeline, Helisphere, to eliminate reads less than 25 nucleotides long or of low quality. These reads were then mapped to the mouse genome (NCBI37/mm9) using an IndexDPgenomic module and reads with a score above 4.3 were allowed. To avoid artefacts from mispriming, reads mapping to regions in the genome where more than four consecutive adenines were coded immediately 3′ to the mapping sequence were excluded from further analysis. Reads were viewed using the Integrative Genomics Viewer32, 33. Total RNA was extracted from FAPs isolated from uninjured C57BL/6 mice using TRIzol (Invitrogen) as per the manufacturer’s instructions. To identify the polyadenylation sites, the sample was reverse transcribed using the SMARTer RACE cDNA amplification kit (Clontech) according to the manufacturer’s instructions using the primers listed in Extended Data Table 1. The amplified fragments were subcloned into pGEM-T-Easy (Promega) and sequenced. Sequencing data were visualized with 4Peaks. To assess levels of the intronic variant and UTR variants, primers were designed to span the Pdgfra transcript (Extended Data Table 2). Variant expression was normalized to Gapdh using the comparative C method27 and reported relative to the average of control-treated samples. A construct corresponding to In-PDGFRα (DNAFORM, AK035501, RIKEN clone 9530057A20) was obtained. This construct was subcloned into the pMXs-IRES-GFP retroviral backbone (Cell BioLabs, Inc.) to generate pMXs-I-Pα. Replication-incompetent retroviral particles were generated by transfection of the 293T human embryonic kidney cell-derived Phoenix helper cell line (gift from G. Nolan). Viral supernatant was filtered through 0.45-μm polyethersulfone filters, concentrated using PEG precipitation and stored at −80 °C. FAPs were plated in 6-well plates and grown in DMEM supplemented with 10% fetal bovine serum (FBS). When cells reached 70% confluency, viral supernatant and polybrene (at a final concentration of 4 μg ml−1) were added to the medium. For overexpression experiments, FAPs were incubated with the viral supernatant for 48 h before analysis. For signalling assays, FAPs were incubated with the viral supernatant for 24 h. Afterwards the medium was changed to serum-free DMEM containing viral supernatant and the cells were incubated for an additional 24 h. The FAPs were then treated with 1 ng ml−1 PDGF-AA for 15 min, after which the cells were used for western blot analysis. A peptide with the sequence GKSAHAHSGKYDLSVV, which represents the unique C-terminal region of In-PDGFRα protein, was generated (Thermo Scientific Pierce, OE0726). To generate In-PDGFRα rabbit polyclonal antibodies directed against In-PDGFRα, New Zealand white rabbits that were specific pathogen free were immunized with 0.25 mg of the peptide in Complete Freund’s Adjuvant. The rabbits received three boosters of antigen consisting of 0.10 mg in Incomplete Freund’s Adjuvant at days 14, 42 and 56 after immunization. Serum was collected at days 70 and 72 (Thermo Scientific Pierce). Cells and homogenized tissues were lysed with RIPA lysis buffer supplemented with protease and phosphatase inhibitors (Roche). The lysates were run on Criterion SDS–PAGE gels (Bio-Rad), transferred to nitrocellulose membranes (Fisher Scientific), and analysed by western blot using the following rabbit antibodies: PDGFRα polyclonal (1:1,000, Cell Signaling, 3174), PDGFRα centre (1:100, Abgent, AP14254c), In-PDGFRα custom (1:1,000), pPDGFRαTyr754 polyclonal (1:1,000, Cell Signaling, 4547), Akt polyclonal (1:1,000, Cell Signaling, 9272), pAkt polyclonal (1:1,000, Cell Signaling, 9271), PLCγ polyclonal (1:1,000, Cell Signaling, 5690), pPLCγ polyclonal (1:1,000, Cell Signaling, 2821), ERK polyclonal (1:2,000, Cell Signaling, 4695), pERK polyclonal (1:2,000, Cell Signaling, 4370), SMAD2/3 monoclonal (1:1,000, Cell Signaling, 8685), and pSMAD2Ser465/Ser467/SMAD3Ser423/Ser425 monoclonal (1:1,000, Cell Signaling, 8828). Membranes were incubated in horseradish-peroxidase-labelled secondary antibodies and bands were visualized with enhanced chemiluminescence (Advansta). siRNAs were designed using the Dharmacon siDESIGN Center for knockdown of In-PDGFRα and FL-PDGFRα (Extended Data Table 2). To knockdown either In-PDGFRα or FL-PDGFRα in FAPs, approximately 8 × 104 cells were plated in a 12-well plate containing DMEM supplemented with 10% FBS and grown to 70–80% confluence. Cells were incubated in 200 nM of either PDGFRα or control siRNAs using Lipofectamine 2000 (Invitrogen). To assess knockdown, cells were collected at 24 h for qPCR analysis. For western blot analyses, 3 × 105 cells were plated in 6-well plates and incubated in Ham’s F10 medium (Invitrogen) supplemented with 10% horse serum (Invitrogen) for 24 h. The medium was then replaced with serum-free Ham’s F10 (Invitrogen) supplemented with 200 nM siRNA and incubated for an additional 24 h. Morpholinos were designed to target two polyadenylation sites on the intronic variant (pA : 5′-TGATTACATTATATCTGTCTTTATT-3′ and pA : 5′-AGCAAAGACCATCATAGCAGAATGA-3′) and the upstream 5′ splice site of the intron (5′ss: 5′-ATGGGCACTTTTACCTAGCATGGAT-3′) (Gene Tools, LLC). For in vitro treatment, cells were grown to 70–80% confluency in DMEM (Invitrogen) supplemented with 10% FBS (Atlanta Biologicals). Cells were incubated in 10 μM of the indicated morpholino using the Endo-Porter transfection reagent (Gene Tools, LLC). Cells were collected at 24 h for qPCR analysis with RNA isolated using the RNeasy Plus Mini kit with on-column DNase digestion as per manufacturer’s instructions (Qiagen). For western blot analysis, cells were transfected for 24 h in Ham’s F10 medium (Invitrogen) supplemented with 10% horse serum (Invitrogen). The medium was then replaced with serum-free Ham’s F10 (Invitrogen) and incubated for an additional 24 h. For signalling assays, cells were then incubated for 15 min with PDGF-AA (Peprotech) at 0.1 ng ml−1 or 20 ng ml−1 for cells treated with pA-AMOs or 5′ss-AMO, respectively, and lysed for western blot analysis as described above. For AMO treatment, FAPs were isolated from the uninjured hindlimb muscles of C57BL/6 mice and seeded at 1 × 105 cells per well in poly-d-lysine-coated 8-well chamber slides (BD Biosciences) coated with ECM gel (Sigma-Aldrich). Cells were transfected with 10 μM AMO using Endoporter (Gene Tools) and expanded for 2 days in Ham’s F10 (Invitrogen) supplemented with 10% horse serum (Invitrogen). The medium was then replaced with Opti-MEM supplemented with 2 ng ml−1 PDGF-AA ligand and 10 μm EdU (Invitrogen). Cells were fixed in 4% paraformaldehyde (Sigma-Aldrich) after 24 h. For siRNA treatment, FAPs were isolated from the uninjured hindlimb muscles of C57BL/6 mice and seeded at 2 × 105 cells per well in poly-d-lysine coated 8-well chamber slides (BD Biosciences) coated with ECM gel (Sigma-Aldrich). The medium was supplemented with 200 nM siRNA and transfected using Lipofectamine 2000 (Invitrogen). After 24 h, the medium was replaced with Opti-Mem and the cells were re-transfected with 200 nM siRNA and 50 ng ml−1 PDGF-AA. In siRNA-treated samples, EdU was not included in this medium. Rather, after 20 h the medium was replaced with Opti-Mem containing 10 μm EdU (Invitrogen). Cells were fixed 4 h later. For retroviral overexpression of In-PDGFRα, FAPs were isolated from uninjured hindlimbs of C57BL/6 mice and seeded at 2 × 105 cells per well in poly-d-lysine coated 8-well chamber slides (BD Biosciences) coated with ECM gel (Sigma-Aldrich). FAPs were cultured in DMEM supplemented with 10% FBS along with viral supernatant and 4 μg ml−1 polybrene. After 24 h, the medium was replaced with serum-free DMEM containing viral supernatant and 20 ng ml−1 PDGF-AA. Twenty hours later, the medium was replaced with Opti-MEM containing 10 μM EdU. Cells were fixed after 4 h. For EdU incorporation experiments, cells were stained using the Click-iT EdU Imaging Kit (Invitrogen). Cells were analysed on a Zeiss Observer Z1 fluorescent microscope (Carl Zeiss) equipped with a Hamamatsu Orca-ER camera (Hamamatsu) and Improvision Volocity software (Perkin Elmer). Cells isolated by FACS from uninjured hindlimb muscles were seeded at a density of 3.5 × 104 cells per well in 96-well plates in Ham’s F10 medium supplemented with 2% horse serum. After 48 h, cells were nearly confluent and the medium was changed to Ham’s F10 with 2% horse serum and 20 ng ml−1 PDGF-AA. A wound was made by scratching a 200-μl pipette tip across the monolayer of cells. The initial scratch area was determined immediately and set to 100%. Images were taken at regular intervals and the scratch area at each time point was measured and calculated as a percentage of the initial scratch area. Scratch closure is defined as the inverse of the cell-free area as a percentage of total area. For in vitro microarray analysis, FAPs were isolated from the uninjured hindlimb muscles of C57BL/6 mice. Cells were plated at 1 × 106 cells per well in 12-well plates. Cells were grown for 2.5 days in DMEM supplemented with 10% FBS. The medium was switched to Ham’s F10 supplemented with 10% horse serum and transfected with 10 μM AMO as indicated for 48 h. The medium was then replaced with Opti-Mem and cells were re-transfected with 10 μM AMO. After 48 h, the cells were lysed and RNA was prepared with the RNeasy Mini Kit as per the manufacturer’s instructions (Qiagen). For in vivo microarray analysis, tibialis anterior muscles were injured with 30 μl of glycerol each and injected with the indicated VMO after 3 days. FAPs were then isolated from the muscles 2 days after VMO injection. Cells were pelleted and RNA prepared from samples as indicated above. The microarray data were obtained using Affymetrix Mouse 1.0 ST. For gene set enrichment analysis (GSEA), the samples were normalized and processed using GenePattern ExpressionFileCreator and PreProcessData set modules. Expression data were analysed and visualized with GSEA28 and GENE-E (http://www.broadinstitute.org/cancer/software/GENE-E/). For ingenuity pathway analysis, including causal network analysis, the samples were normalized using Affymetrix Expression Console Software and analysed for enrichment using IPA (Ingenuity Systems, http://www.ingenuity.com). Array data were deposited into Gene Expression Omnibus (Accessions GSE60099 and GSE81744). Vivo-morpholinos were designed to target two polyadenylation sites on the intronic variant (pA -VMO: 5′-TGATTACATTATATCTGTCTTTATT-3′ and pA -VMO: 5′-AGCAAAGACCATCATAGCAGAATGA-3′) and the upstream 5′ splice site of the intron (5′ss-VMO: 5′-ATGGGCACTTTTACCTAGCATGGAT-3′) (Gene Tools, LLC). For treatment in vitro, cells were isolated from hindlimb muscles of C57BL/6 mice and grown to 70–80% confluency in DMEM (Invitrogen) supplemented with 10% FBS (Atlanta Biologicals). Cells were incubated in the 10 μM of the indicated morpholino (Gene Tools, LLC). Cells were collected at 24 h for qPCR analysis. For in vivo qPCR analysis, tibialis anterior muscles were injured with glycerol as described above and injected with 250 ng of the indicated VMO at the site of injury 3 days later. FAPs were sorted by FACS 7 days after VMO injection for qPCR analysis. For ex vivo proliferation and scratch assays, tibialis anterior muscles were injured with glycerol and injected with 250 ng of the indicated VMO 3 days after injury. FAPs were isolated 2 days later by FACS. In EdU incorporation studies, cells were seeded at 4 × 104 cells per well in poly-d-lysine-coated 8-well chamber slides (BD Biosciences) coated with ECM gel (Sigma-Aldrich). Cells were incubated in 10 ng ml−1 PDGF-AA (Peprotech) and 10 μM EdU (Invitrogen) for 24 h. The cells were fixed and stained. In the ex vivo proliferation studies as well as the in vivo proliferation studies described below, the proliferation index was used to denote the percentage EdU incorporation normalized to control. In the scratch assays, cells were seeded and treated as described above. For in vivo proliferation studies, tibialis anterior muscles were injected with 150 ng of the indicated VMO at 0 and 24 h. FAPs were isolated at 48 h via FACS. To assess in vivo proliferation, the cells were exposed to 10 μM EdU immediately after muscle isolation and incubated in 10 μM EdU ex vivo during the collagenase, collagenase/dispase, and antibody incubations as described above. The cells were plated in poly-d-lysine-coated 8-well chamber slides (BD Biosciences) coated with ECM gel (Sigma-Aldrich), fixed 1 h after plating, and stained using the Click-iT EdU Imaging Kit (Invitrogen). For histological analysis, tibialis anterior muscles were injured with glycerol or BaCl and injected at the site of injury with 250 ng of the indicated VMO. After 7 days, the muscles were snap frozen in isopentane cooled in liquid nitrogen immediately after dissection. Muscles sections were stained with Gomori-trichrome (Richard-Allan Scientific) per manufacturer’s instructions or oil red O (Sigma-Aldrich) as previously described29. The fibrotic index was calculated as the area of fibrosis divided by total area of muscle normalized to control-treated muscle. The fibro–adipose index was defined as the area of fibrosis plus the area of adiposis (as detected by oil red O staining) divided by total area of muscle, normalized to control. Major factors in determining sample size included the level of the effect and the inherent variability in measurements obtained. No statistical methods were used to predetermine sample size. Animals were excluded from the study only if their health status was compromised, such as occurred when animals had visible wounds from fighting. Samples were not specifically randomized or blinded. However, mouse identifiers were used when possible to blind evaluators to experimental conditions, and all samples within experiments were processed identically for measurement quantification using automated tools as specified. The sequencing data were deposited into the NCBI Sequence Read Archive (accession number SRP079186). Array data were deposited into Gene Expression Omnibus (accession numbers GSE60099 and GSE81744).


Reissmann S.,Friedrich - Schiller University of Jena | Reissmann S.,Jena Bioscience GmbH
Journal of Peptide Science | Year: 2014

The penetration of polar or badly soluble compounds through a cell membrane into live cells requires mechanical support or chemical helpers. Cell-penetrating peptides (CPPs) are very promising chemical helpers. Because of their low cytotoxicity and final degradation to amino acids, they are particularly favored in in vivo studies and for clinical applications. Clearly, the future of CPP research is bright; however, the required optimization studies for each drug require considerable individualized attention. Thus, CPPs are not the philosopher's stone. As of today, a large number of such transporter peptides with very different sequences have been identified. These have different uptake mechanisms and can transport different cargos. Intracellular concentrations of cargos can reach a low micromole range and are able to influence intracellular reactions. Internalized ribonucleic acids such as small interfering RNA (siRNA) and mimics of RNA such as peptide nucleic acids, morpholino nucleic acids, and triesters of oligonucleotides can influence transcription and translation. Despite the highly efficient internalization of antibodies, enzymes, and other protein factors, as well as siRNA and RNA mimics, the uptake and stabile insertion of DNA into the genome of the host cells remain substantially challenging. This review describes a wide array of differing CPPs, cargos, cell lines, and tissues. The application of CPPs is compared with electroporation, magnetofection, lipofection, viral vectors, dendrimers, and nanoparticles, including commercially available products. The limitations of CPPs include low cell and tissue selectivity of the first generation and the necessity for formation of fusion proteins, conjugates, or noncovalent complexes to different cargos and of cargo release from intracellular vesicles. Furthermore, the noncovalent complexes require a strong molar excess of CPPs, and extensive experimentation is required to determine the most optimal CPP for any given cargo and cell type. Yet to predict which CPP is optimal for any given target remains a complex question. More recently, there have been promising developments: the enhancement of cell specificity using activatable CPPs, specific transport into cell organelles by insertion of corresponding localization sequences, and the transport of drugs through blood-brain barriers, through the conjunctiva of eyes, skin, and into nerve cells. Proteins, siRNA, and mimics of oligonucleotides can be efficiently transported into cells and have been tested for treatment of certain diseases. The recent state of the art in CPP research is discussed together with the overall scope, limitations, and some recommendations for future research directions. Copyright © 2014 European Peptide Society and John Wiley & Sons, Ltd.


Kushnir S.,Max Planck Institute of Molecular Physiology | Cirstea I.C.,Max Planck Institute of Molecular Physiology | Basiliya L.,Max Planck Institute of Molecular Physiology | Lupilova N.,Max Planck Institute of Molecular Physiology | And 2 more authors.
Molecular and Biochemical Parasitology | Year: 2011

The trypanosomatid protozoon Leishmania tarentolae is a well-established model organism for studying causative agents of several tropical diseases that was more recently developed as a host for recombinant protein production. Although several expression architectures based on foreign RNA polymerases have been established for this organism, all of them rely on integration of the expression cassette into the genome. Here, we exploit a new type of expression architecture based on linear elements. These expression vectors were propagated in Escherichia coli as circular plasmids and converted into linear episomes with telomere-like structures prior to transfection of L. tarentolae. Overexpression of recombinant proteins in transgenic organisms exceeding 10% of total cellular protein, one of the highest overexpression levels obtained in a eukaryotic organism for a cytosolic protein. We show that the linear elements are stably propagated in L. tarentolae cells over long periods of time (>90 generations) without major changes in structure or expression yields. Overexpressing cultures can be obtained without clonal selection of the transfected cells. To establish the utility of the developed system for protein production in a parallelized format, we expressed 37 cytosolic, peripheral, and membrane proteins as fusions with EGFP in L. tarentolae using linear vectors. We detected the expression of 30 of these targets and describe the preparative purification of two arbitrarily selected proteins. © 2011 Elsevier B.V. All rights reserved.


Mussbach F.,Jena Bioscience GmbH | Mussbach F.,Friedrich - Schiller University of Jena | Franke M.,Friedrich - Schiller University of Jena | Zoch A.,Friedrich - Schiller University of Jena | And 3 more authors.
Journal of Cellular Biochemistry | Year: 2011

Internalization of peptides and proteins into live cells is an essential prerequisite for studies on intracellular signal pathways, for treatment of certain microbial diseases and for signal transduction therapy, especially for cancer treatment. Cell penetrating peptides (CPPs) facilitate the transport of cargo-proteins through the cell membrane into live cells. CPPs which allow formation of non-covalent complexes with the cargo are used primarily in this study due to the relatively easy handling procedure. Efficiency of the protein uptake is estimated qualitatively by fluorescence microscopy and quantitatively by SDS-PAGE. Using the CPP cocktail JBS-Proteoducin, the intracellular concentrations of a secondary antibody and bovine serum albumin can reach the micromolar range. Internalization of antibodies allows mediation of intracellular pathways including knock down of signal transduction. The high specificity and affinity of antibodies makes them potentially more powerful than siRNA. Thus, CPPs represent a significant new possibility to study signal transduction processes in competition or in comparison to the commonly used other techniques. To estimate the highest attainable intracellular concentrations of cargo proteins, the CPPs are tested for cytotoxicity. Cell viability and membrane integrity relative to concentration of CPPs are investigated. Viability as estimated by the reductive activity of mitochondria (MTT-test) is more sensitive to higher concentrations of CPPs versus membrane integrity, as measured by the release of dead cell protease. Distinct differences in uptake efficiency and cytotoxic effects are found using six different CPPs and six different adhesion and suspension cell lines. © 2011 Wiley Periodicals, Inc.


News Article | October 23, 2015
Site: www.nature.com

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. Cloning of the pET15b-CarH construct encoding for Thermus thermophilus CarH with an N-terminal His -tag was described previously8. The H132A mutation was introduced into pET15b-CarH using QuikChange PCR mutagenesis (Stratagene) with Pfu Turbo DNA polymerase. All other mutants were obtained by gene synthesis (Genscript) with appropriate 5′ and 3′ restriction sites for cloning into the pET15b expression vector. WT CarH and mutants were purified as described previously8. A slightly modified protocol was used for His -tagged CarH for crystallization. After expression and affinity chromatography, performed as described previously8, a threefold molar excess of AdoCbl (Sigma) was added and the mixture was incubated on ice for 1 h. All subsequent handling was performed in a dark room under red light. The protein solution was applied to a HiLoad 26/60 Superdex 200 size exclusion column (GE Healthcare) pre-equilibrated with CarH buffer (0.1 M NaCl, 0.05 M Tri⋅HCl, pH 8). Under these conditions, tetrameric AdoCbl-bound CarH eluted as a single peak, separate from residual amounts of monomeric CarH. Fractions containing AdoCbl-bound CarH were combined and concentrated to about 8 mg ml−1, as judged by the absorbance at 280 nm using the combined ε for AdoCbl (22.5 mM–1 cm–1, determined spectroscopically on the basis of published extinction coefficients at 260 nm, 288 nm, and 522 nm (refs 41, 42, 43)) and for CarH (37.9 mM–1 cm–1, calculated from the protein sequence using ProtParam at http://web.expasy.org/protparam). Purified native and mutant protein identities were verified before use by high-performance liquid chromatography (HPLC) coupled to electrospray ionization–time-of-flight (ESI–TOF) or ion-trap mass spectrometry using an Agilent 1100 Series HPLC equipped with a µ-well plate autosampler and a capillary pump and connected to an Agilent Ion Trap XCT Plus Mass Spectrometer with an ESI interface (Agilent Technologies). Samples were injected into a Zorbax Poroshell 300 SB-C18 HPLC column (Agilent Technologies) that was coupled online to the mass spectrometer using an electrospray interface. Samples were separated at 60 °C at a flow rate of 0.2 ml min–1 using a linear gradient of buffer A (water/acetonitrile/formic acid, 95:4.9:0.1) to 90% buffer B (water/acetonitrile/formic acid, 10:89.9:0.1) over 30 min and protein elution was monitored at 210 nm and 280 nm. Mass spectra were acquired in the positive ion mode in an m/z range from 100 to 2,200. The integrity of the AdoCbl Co–C bond was assessed by UV–vis spectroscopy (described below). Protein containing intact AdoCbl was flash-frozen in liquid nitrogen until further use. CarH containing photolysed AdoCbl was generated by exposing the protein solution to ambient light for 30 min at 4 °C. Complete photolysis was assessed by UV–vis spectroscopy (described below). Light-exposed CarH was used for crystallization experiments immediately. CarH–DNA complexes for crystallization were generated by mixing protein and DNA at the desired ratio and incubating the mixture for 1 h on ice in the dark before crystallization experiments. HPLC-purified single-stranded DNA oligonucleotides without heavy atom labels (Integrated DNA Technologies) or containing a single 5-iodo-deoxycytidine (Jena Bioscience) were dissolved to 1 mM in CarH buffer. Equimolar amounts of complementary oligonucleotides were mixed, heated to 95 °C for 10 min, and then slowly left to cool down to 4 °C in a thermocycler over the course of 1 h for annealing. Final double-stranded DNA concentrations were assessed by the absorbance at 260 nm using the calculated sequence-specific ε (http://biophysics.idtdna.com/UVSpectrum.html). Purified AdoCbl-bound CarH was crystallized in three different crystal forms. All crystallization procedures for AdoCbl-bound CarH were performed in a dark room under red light. Crystals of AdoCbl-bound CarH in crystal form 1 were obtained by the hanging drop vapour diffusion technique at 25 °C. An aliquot (1 µl) of a protein solution (7 mg ml−1 AdoCbl-bound CarH in CarH buffer) was mixed with 1 µl of a precipitant solution (10% (w/v) PEG 8000, 10% (v/v) glycerol, 0.04 M KH PO ) on a glass cover slip. The cover slip was sealed with grease over a reservoir containing 500 µl of the precipitant solution. Octahedral crystals appeared within 3 days and grew to maximum size within 7 days. Under these conditions, the protein underwent proteolysis at the linker region between the DNA-binding domain and the four-helix bundle, as judged by SDS–polyacrylamide gel electrophoresis. The crystals consisted only of the C-terminal light-sensing domains. Crystals were transferred in two steps of increasing glycerol concentration into a cryogenic solution containing 10% (w/v) PEG 8000, 20% (v/v) glycerol, 0.04 M KH PO , 0.05 M Tri⋅HCl pH 8, and 0.1 M NaCl, soaked in that solution for 20 s, and then flash-frozen in liquid nitrogen. A second crystal form of AdoCbl-bound CarH was obtained by the sitting drop vapour diffusion technique at 25 °C. An aliquot (0.15 µl) of a protein solution (5.9 mg ml−1 AdoCbl-bound CarH in CarH buffer, supplemented with 70 µM of a 31-bp DNA oligonucleotide) was mixed with 0.15 µl of a precipitant solution (20% (w/v) PEG 3350, 0.2 M KCl) using a Phoenix liquid handling robot (Art Robbins Instruments). The drop was equilibrated against 70 µl of the precipitant solution. Rectangular crystals appeared within 6 months. Again, the protein underwent proteolysis and the crystals only consisted of the C-terminal light-sensing domains. Crystals were transferred in two steps of increasing glycerol concentration into a cryogenic solution containing the precipitant supplemented with 20% (v/v) glycerol, soaked in that solution for 5 s, and then flash-frozen in liquid nitrogen. A third crystal form of AdoCbl-bound CarH containing full-length protein was obtained by the sitting drop vapour diffusion technique at 25 °C. An aliquot (0.15 µl) of a protein solution (6 mg ml−1 AdoCbl-bound CarH in CarH buffer, supplemented with 70 µM of a 28-bp DNA segment) was mixed with 0.15 µl of a precipitant solution (20% (w/v) PEG 3350, 0.1 M ammonium citrate tribasic pH 7) using a Phoenix liquid handling robot (Art Robbins Instruments). The drop was equilibrated against 70 µl of the precipitant solution. Rod crystals appeared within 1 month. These crystals contained full-length AdoCbl-bound CarH but no DNA. Crystals were transferred in three steps of increasing glycerol concentration into a cryogenic solution containing the precipitant supplemented with 20% (v/v) glycerol, soaked in that solution for 10 s, and then flash-frozen in liquid nitrogen. Light-exposed CarH was crystallized by the hanging drop vapour diffusion technique at 25° C. An aliquot (1 µl) of a protein solution (4.5 mg ml−1 light-exposed CarH in CarH buffer) was mixed with 1 µl of a precipitant solution (3.4 M NaCl, 0.1 M Bis-Tris pH 6) on a glass cover slip. The cover slip was sealed with grease over a reservoir containing 500 µl of the precipitant solution. Octahedral crystals appeared within 8 months. Crystals were transferred in three steps of increasing glycerol concentration into a cryogenic solution containing the precipitant supplemented with 18% (v/v) glycerol, incubated in that solution for 10 s, and then flash-frozen in liquid nitrogen. CarH bound both to AdoCbl and to a 26-bp DNA segment was crystallized by the hanging drop vapour diffusion technique at 25 °C. An aliquot (1 µl) of a protein solution (6 mg ml−1 AdoCbl-bound CarH in CarH buffer, supplemented with 67.5 µM of a 26-bp DNA segment, 1.5-fold molar excess) was mixed with 1 µl of a precipitant solution (16% PEG 3350, 0.2 M l-proline, 0.1 M HEPES pH 7.5) on a glass cover slip. The cover slip was sealed with grease over a reservoir containing 500 µl of the precipitant solution. Tetragonal bipyramidal crystals appeared within 3 weeks. Crystals were transferred in three steps of increasing PEG 400 concentration into a cryogenic solution containing the precipitant supplemented with 15% (w/v) PEG 400, incubated in that solution for 20 s, and then flash-frozen in liquid nitrogen. CarH bound to both AdoCbl and a 26-bp DNA segment containing 5-iodo-deoxycytidine (Extended Data Fig. 6b–d) in position −25 of the sense strand (Extended Data Fig. 5a) was crystallized by the hanging drop vapour diffusion technique at 25 °C. An aliquot (1 µl) of a protein solution (5 mg ml−1 AdoCbl-bound CarH in CarH buffer, supplemented with 94 µM of the iodine-labelled 26-bp DNA segment, 2.5-fold molar excess) was mixed with 1 µl of a precipitant solution (11.5% PEG 3350, 0.28 M l-proline, 0.1 M Tris pH 8.5) on a glass cover slip. The cover slip was sealed with grease over a reservoir containing 500 µl of the precipitant solution. Crystals appeared within 4 months. Crystals were transferred in five steps of increasing xylitol concentration into a cryogenic solution containing the precipitant supplemented with 25% (w/v) xylitol, incubated in that solution for 30 s, and then flash-frozen in liquid nitrogen. All data were collected at the Advanced Photon Source (Argonne, Illinois, USA) at beamline 24ID-C using a Pilatus 6M pixel detector at a temperature of 100 K. Crystals of AdoCbl-bound CarH crystal form 1 belong to space group P4 2 2. An initial AdoCbl-bound CarH crystal was used for a fluorescence scan to determine the Co peak wavelength for anomalous data collection. Another crystal was then used for collection of both native data and anomalous peak data. Native data were collected in a single wedge of 75° at a wavelength of 0.9792 Å (12,662 eV). The crystal was displaced continuously along its major macroscopic axis during data collection. Anomalous peak data were collected in a single wedge of 345° at a wavelength of 1.6039 Å (7,730 eV). The crystal was aligned using a mini-κ goniometer such that Bijvoet mates were recorded on the same frame. All other data except for iodine anomalous data and native data of light-exposed CarH (see below) were collected at a wavelength of 0.9795 Å (12,658 eV). Crystals of AdoCbl-bound CarH crystal form 2 belong to space group P2 2 2 . Data were collected in a single wedge of 100°. Crystals of AdoCbl-bound CarH crystal form 3 belong to space group P1. Data were collected in a single wedge of 270° and the crystal was displaced continuously along its major macroscopic axis during data collection. Crystals of light-exposed CarH belong to space group I4 22 and data were collected at a wavelength of 0.9791 Å (12,663 eV) in a single wedge of 150°. Crystals of DNA-bound CarH both with and without the iodine label belong to space group P2 2 2. Data for crystals with unlabelled DNA were collected in a single wedge of 180°. Data for crystals of CarH in complex with iodine-labelled DNA were collected at a wavelength of 1.7365 Å (7,140 eV) in a single wedge of 200° and the crystal was displaced continuously along its major macroscopic axis during data collection. Data for the AdoCbl-bound CarH (crystal form 1) Co peak data set were integrated in HKL2000 and scaled in Scalepack44. Data for all other data sets were integrated in XDS and scaled in XSCALE45. Data collection statistics are summarized in Extended Data Table 1. The structure of AdoCbl-bound CarH in crystal form 1 (space group P4 2 2) was determined to 2.80 Å resolution using single-wavelength anomalous diffraction. Positions of two cobalt sites, corresponding to two CarH protomers in the asymmetric unit, were located using ShelxD46 in the HKL2MAP shell47 and refined using SHARP/autoSHARP48. The initial overall figure of merit (acentric) was calculated by SHARP to be 0.43 to 5.1 Å resolution. Experimental maps from the SHARP output, solvent flattened using SOLOMON49 and extended to 3.3 Å resolution, were of sufficient quality to place two copies of the Cbl-binding domain of MetH17 (PDB accession number 1BMT, residues 745–868), eight additional helices, and AdoCbl in the electron density. This initial model was used to better define solvent boundaries in another round of solvent flattening of SOLOMON. Using the resulting electron density, loop regions were modified and side chains with visible electron density were added. A near-complete model of AdoCbl-bound CarH (containing 374 amino-acid residues and bound AdoCbl) was then used for rigid body refinement in Phenix50 against the native AdoCbl-bound CarH data set (crystal form 1) using data from 100 to 2.80 Å resolution. The resulting R-factors were 42.0% and 44.1% for the working and the free R-factor, respectively. The model was refined by manual adjustment in Coot51 until rigid body refinement in Phenix yielded R-factors of 30.8% and 34.7% for the working and the free R-factor, respectively. Subsequent cycles of refinement included positional refinement with non-crystallographic symmetry restraints and individual B-factor refinement in Phenix until the R-factors were 20.9% and 24.2% for the working and the free R-factor, respectively. This model was not refined to completion. The near-complete model was used to determine the structures of AdoCbl-bound CarH in crystal form 2 (space group P2 2 2 ) and crystal form 3 (space group P1), which are of higher resolution (crystal form 2) or contain the full-length protein (crystal form 3). The structure of AdoCbl-bound CarH in crystal form 2 was determined to 2.15 Å resolution by molecular replacement in Phaser52. The structure in crystal form 2 contains four CarH protomers in the asymmetric unit, corresponding to a tetramer. After molecular replacement, ten cycles of simulated annealing refinement were performed in Phenix to remove model bias. The model was then refined by iterative cycles of manual adjustment in Coot and refinement in Phenix. Initially, strict non-crystallographic symmetry restraints were applied for the two head-to-tail dimers in the asymmetric unit. Subsequently, these restraints were loosened for residues that are in unique environments either because of the asymmetric tetramer architecture or because of crystal contacts. In advanced stages of refinement, water molecules were added manually in Coot and refined in Phenix, with placement of additional water molecules until their number was stable. Final cycles of refinement included TLS parametrization53 with one TLS group per CarH protomer. The structure of AdoCbl-bound CarH in crystal form 3 was determined to 2.80 Å resolution using molecular replacement. First, two CarH tetramers were placed in the asymmetric unit using Phaser, accounting for all eight protomers in the asymmetric unit. Subsequently, four CarH DNA-binding domains from the structure of light-exposed CarH (see below) were placed using Phaser. After refinement in Phenix, there was clear electron density for an additional DNA-binding domain as well as fragments of the three remaining DNA-binding domains, accounting for all eight DNA-binding domains in the asymmetric unit. The model was refined by iterative cycles of manual adjustment in Coot and refinement in Phenix. Strict non-crystallographic symmetry restraints were applied for all CarH protomers in the asymmetric unit and loosened in later stages of refinement as described above. No water molecules were added to this structure. Final cycles of refinement included TLS parametrization53. For each CarH protomer, the light-sensing domain was defined as a single TLS group and, if fully present, the DNA-binding domain was defined as an additional TLS group. The structure of light-exposed CarH was determined to 2.65 Å resolution by molecular replacement in Phaser using consecutive searches for the CarH Cbl-binding domain, the four-helix bundle, and the first conformation of the NMR structure of the CarA DNA-binding domain (PDB accession number 2JML16). The structure contains one protomer in the asymmetric unit and all three domains could be placed unambiguously. Ten cycles of simulated annealing refinement were performed in Phenix. The model was then refined by iterative cycles of manual adjustment in Coot and refinement in Phenix. In advanced stages of refinement, water molecules were added manually in Coot and refined in Phenix, with placement of additional water molecules until their number was stable. Final cycles of refinement included TLS parametrization53 with one TLS group. The structure of CarH bound to AdoCbl and a 26-bp DNA segment was determined to 3.89 Å resolution by molecular replacement in Phaser using consecutive searches for two CarH tetramers without the DNA-binding domains and for two 26-bp DNA segments (models generated by the 3D-DART server54; http://haddock.science.uu.nl/services/3DDART/). After molecular replacement, there was clear electron density for six DNA-binding domains in the asymmetric unit, which were positioned manually in the electron density from the structure of light-exposed CarH. The model was refined by iterative cycles of manual adjustment in Coot and refinement in Phenix. B-factors were refined grouped by residue and positions of individual atoms were restrained using non-crystallographic symmetry restraints. Planarity and hydrogen bonding restraints were applied to DNA base pairs. Final cycles of refinement included TLS parametrization using one TLS group for each CarH protomer and each DNA segment. Anomalous difference maps, calculated from data collected on crystals that contained a DNA segment with an iodine label at position −25 (Extended Data Fig. 5a) were used to unambiguously determine the orientation of the DNA segment in the crystal structure and thus validate the sequence assignments. Maps, calculated using FFT55 in the CCP4 software suite56, revealed a strong anomalous difference density peak at one position for each of the two CarH–DNA complexes in the asymmetric unit, allowing for position −25 of the sense strand to be assigned in the structure (Extended Data Fig. 6b–d). Note that the iodine-labelled DNA segment differed slightly from the DNA segment used in the structure determination (Extended Data Fig. 5a), but both crystallize in the same space group and with the same crystal packing. Parameter files for cobalamin were provided by O. Smart at Global Phasing. Refinement restrains for the 5′-dAdo group were generated using the Grade Web Server (Global Phasing). Crystallographic refinement of all CarH structures yielded models possessing low free R-factors, excellent stereochemistry, and small root mean square deviations from ideal values for bond lengths and angles. In all models, side chains without visible electron density were truncated to the last atom with electron density, and amino acids without visible electron density were not included in the model. All refinement statistics are summarized in Extended Data Table 1. The models were validated using simulated annealing composite omit maps (AdoCbl-bound CarH, light-exposed CarH) or regular refinement composite omit maps (DNA-bound CarH) calculated in CNS57 and Phenix. Model geometry was analysed using MolProbity58 and ProCheck59. Analysis of the Ramachandran statistics using MolProbity indicated that for AdoCbl-bound CarH (crystal form 2), 98.1%, 1.9%, and 0.0% of residues are in the favoured, allowed, and disallowed regions, respectively; for AdoCbl-bound CarH (crystal form 3), 97.7%, 2.3%, and 0.0% of residues are in the favoured, allowed, and disallowed regions, respectively; for light-exposed CarH, 97.8%, 2.2%, and 0.0% of residues are in the favoured, allowed, and disallowed regions, respectively; and for AdoCbl- and DNA-bound CarH, 97.1%, 2.7%, and 0.2% of residues are in the favoured, allowed, and disallowed regions, respectively. The larger number of residues in the disallowed region of the Ramachandran plot of DNA-bound CarH is due to the modest resolution of the structure. Figures were generated using PyMOL60. Interfaces between subunits were analysed using the ‘Protein interfaces, surfaces and assemblies’ service PISA at the European Bioinformatics Institute (http://www.ebi.ac.uk/msd-srv/prot_int/pistart.html)61. Crystallography software packages were compiled by SBGrid62. All DNA binding assays were repeated three to five times for each experimental condition. EMSAs were performed in the dark as described previously8. A 177-bp DNA probe PCR-amplified using primers with one 5′-end 32P-labelled with T4 polynucleotide kinase (T4PK; Takara) before the PCR or shorter HPLC-purified synthetic probes (Biolegio) were used in the EMSAs. With the latter, one strand was 32P-labelled at the 5′-end with T4PK and then mixed with a twofold excess of the unlabelled complementary strand to ensure that all of the labelled strand was present as double-stranded probe. The strand mixture was incubated at 100 °C for 2 min and then slowly left to cool down for hybridization. For EMSAs, a 20 μl reaction volume containing the DNA probe (1.2 nM, approximately 13,000 counts per minute) and protein with a fivefold excess of AdoCbl in 0.1 M KCl, 0.025 M Tri⋅HCl, pH 8, 1 mM DTT, 10% (v/v) glycerol, 200 ng µl−1 BSA, and 1 µg of sheared salmon sperm DNA as non-specific competitor was incubated for 30 min at 65 °C (177-bp probe) or 30 °C (shorter probes). They were then loaded onto 6% native polyacrylamide gels (37.5:1 acrylamide:bisacrylamide) pre-run for 30 min in 0.5× TBE buffer (0.045 M Tris base, 0.045 M boric acid, 1 mM EDTA) and subjected to electrophoresis for 1.5 h at 200 V, 10 °C. Gels were vacuum-dried and analysed by autoradiography. Autoradiograms were scanned using an Image Scanner II imager with LabScan 5.0 software (GE Healthcare). Band intensities were quantified using ImageJ (NIH) with those of free DNA used to estimate the fraction bound, which was fitted to the three-parameter Hill equation using SigmaPlot (Systat Software) to estimate K the apparent equilibrium dissociation constant equivalent to the protein concentration for half-maximal binding, and n, the Hill coefficient. The latter, for example expected to be 2 for dimer or 4 for tetramer DNA-binding models, can vary owing to cooperativity effects, contributions from monomer-tetramer equilibria, and/or deviations from true equilibrium. DNase I and hydroxyl radical footprinting analyses were performed under solution conditions similar to EMSA using previously described protocols8, 63. A 130-bp CarH operator-promoter DNA probe (Extended Data Fig. 5a) was 32P-radiolabelled at the 5′ end of its sense or anti-sense strand by PCR using appropriately labelled primers, as described above. For DNase I footprinting, 20 μl of 32P-radiolabelled DNA probe (~20,000 counts per minute) with 800 nM CarH and fivefold excess of AdoCbl in EMSA buffer lacking glycerol and with 0.01 M MgCl were incubated for 30 min at 37 °C, then treated with 0.07 units of DNase I for 2 min and finally quenched with 0.025 M EDTA. For hydroxyl radical footprinting, samples (as for DNase I footprints but without MgCl ) were treated with 2 μl each of freshly prepared Fe(II)-EDTA solution (1 mM ammonium iron (II) sulfate, 2 mM EDTA), 0.01 M sodium ascorbate, and 0.6% hydrogen peroxide for 4 min at 25 °C. The reaction was stopped with 2 μl each of 0.1 M thiourea and 0.5 M EDTA (pH 8). Footprinting reactions were done under dim light and, after quenching, under normal light. DNA from each sample was ethanol precipitated, washed twice with 70% ethanol, dried, and resuspended in formamide loading buffer. The 5 μl samples were heated at 95 °C for 3 min and loaded onto a 6% polyacrylamide-8 M urea sequencing gel together with G + A chemical sequencing ladders. Gels were vacuum-dried and analysed by autoradiography, and the bands quantitated using GelAnalyzer 2010a (http://www.gelanalyzer.com). Each experiment was repeated at least three times. Analytical SEC for all CarH mutants except for H132A CarH was performed using an ÄKTAbasic unit and a Superdex 200 analytical SEC column (GE Healthcare)8. The calibration curve was log(M  = 7.885 − 0.221V ), where M is the apparent molecular mass and V is the elution volume. Pure protein (100 μl, 50−100 μM) was incubated with a fivefold molar excess of AdoCbl for at least 15 min and analysed by SEC in the dark or after light irradiation for 5 min with white light from fluorescent lamps at 10 W m−2. Elution at 0.4 ml min−1 flow rate was tracked by absorbance at 280 nm and 522 nm, and M was estimated from V . Each SEC experiment was performed at least three times. Analytical SEC for H132A CarH was performed using an ÄKTA FPLC unit and a Superose 6 10/300 GL column (GE Healthcare) equilibrated with CarH buffer. The calibration curve was log(M  = 9.74 − 0.30V ). WT or H132A CarH (300 µl, 20–50 µM) with stoichiometric AdoCbl with or without exposure to white light for 1 h were injected onto the column and elution was tracked by absorbance at 280 nm. For AdoCbl exchange studies, WT or H132A CarH samples were exposed to light as described and then incubated with a tenfold excess of free AdoCbl for the given periods and temperatures and analysed by SEC. Solution UV–vis spectra were recorded at 25 °C on a SpectraMax Plus 384 (Molecular Devices) using SoftMax Pro 5 software (Molecular Devices) and a 1 cm path length quartz cuvette (Starna). WT or H132A CarH in CarH buffer were transferred to the cuvette under red light or after exposure to white light for 20 min and UV–vis spectra were recorded from 250 to 800 nm. The spectrum of pure CarH buffer was used for background subtraction. No photolysis occurred on the timescale of spectrum acquisition, as repeated acquisition did not lead to spectral changes. Spectra of Cbl with increasing imidazole concentrations, similar to spectra reported previously20, were obtained with the same experimental parameters. Cbl solutions contained 50 µM OHCbl˙HCl (Sigma) in 50 mM Tris with 0 mM, 0.4 mM, or 400 mM imidazole, adjusted to a final pH of 8 to match the protein solutions. All solutions were incubated for 16 h at 25 °C to allow complete ligand exchange to take place. Single-crystal UV–vis spectra were recorded at a temperature of 100 K at Stanford Synchrotron Radiation Laboratory beamline 11-1 (Menlo Park, California, USA) using a UV–vis microspectrophotometer. The setup used a Hamamatsu light source (50 µm light spot) with deuterium and halogen lamps, UV solarization-resistant optical fibres, reflective Newport Schwardchild objectives, and an Ocean Optics QE65000 Spectrum Analyzer. Spectra were acquired as 50 averages with an integration time of 0.03 s and a boxcar width of 3. A crystal of AdoCbl-bound CarH was cryoprotected, transferred to a nylon fibre loop, and frozen in liquid nitrogen as described above. A background spectrum was acquired on a region of the fibre loop containing just cryoprotectant. A sample spectrum was then acquired on the crystal.


Persistence Market Research (PMR) delivers key insights on the global cell free protein expression market in its upcoming report titled, "Global Market Study on Cell Free Protein Expression: Continuous Flow Expression System Segment Expected to Account for a Higher Market Share Between 2016 and 2024". In terms of revenue, the global cell free protein expression market is projected to register a CAGR of 6.0% over the forecast period owing to various factors, on which PMR offers detailed insights and forecasts. The primary factors fuelling demand for cell free protein expression market are increasing R&D outsourcing by pharmaceutical and biotechnological companies, declining R&D productivity and patent cliff sales drop leading to increasing research intensity in the pharmaceutical sector and increasing expenditure on biosimilar development. Other factors driving cell free protein expression market are a growing demand for simple and efficient protein production methods, contamination free approach and increasing focus on production of mammalian cell free lysate due to drug discovery. One of the major bottleneck in the cell free protein expression market is the low protein production volumes in the process. This limitation makes cell free protein expression unsuitable for large industrial applications and is generally preferred in small R&D processes. However, efforts are being made to overcome this constraint and make it a preferred expression system in industrial settings. The market is segmented based on product type, application, expression mode, end users, and region. Based on product type, the market has been segmented into E. coli cell-free protein expression system, rabbit reticulocytes cell-free protein expression system, wheat germ cell-free protein expression system, insect cells cell-free protein expression system, mammalian cell-free protein expression system, and consumables (labelling tags, vectors). Mammalian cell-free protein expression system segment is expected to grow with the fastest CAGR over the forecast period, owing to increasing usage of mammalian cell lysate in humanized proteomic and biologics study. The segment is expected to register a significant CAGR of 6.0% during the forecast period. Wheat germ cell-free protein expression system is also anticipated to witness a CAGR of 5.8% over the forecast period. The rapidly increasing demand for simpler, rapid and efficient protein production methods is boosting the cell-free protein expression market in research and development field. The market has been segmented based on major applications such as enzyme engineering, protein labelling, protein-protein interaction, and protein purification. The protein-protein interaction segment is expected to register the highest CAGR of 6.0% over the forecast period due to increasing number of proteomic studies. The simple presentation of cell-free protein expression system makes it easier to integrate them into high throughput platforms for efficient biologics and proteomics studies. Cell-free protein expression systems allow protein screening without necessitating a gene-cloning step thus enabling an accelerated process/product development pipelines which makes up for attractive opportunity for market players. Based on end users, the market has been segmented into biotechnological companies, pharmaceutical companies, contract research organizations, and academic/research institutes.  The academic/research institutes is anticipated to account for the highest market share over the forecast period, registering a CAGR of 6.1% due to vast research applications in the protein libraries generation for functional genomic studies, customized drug development studies, and the expression of virus-like particles, among many other applications. This report assesses trends, that drive growth of each segment on the global as well as regional levels, and offers potential takeaways, that could prove substantially useful to biopharmaceutical manufacturing companies who wish to enter into the cell free protein expression market. North America and Europe are expected to dominate the cell free protein expression market with maximum market share in 2016. North America and Europe collectively, are expected to account for more than 65% of the total cell free protein expression market share in terms of value in 2016. Among emerging markets, Asia Pacific is estimated to exhibit the highest CAGR of 6.1% over the forecast period, due to increase in the research and development expenditure in the region. Some key players in the global cell free protein expression market identified in the report include Thermo Fisher Scientific, Jena Bioscience GmbH, New England Biolabs, biotechrabbit GmbH, Bioneer Corporation, Promega Corporation, Cube Biotech GmbH, Takara Bio, Inc., CellFree Sciences Co., Ltd., and GeneCopoeia, Inc. We have discussed individual strategies of these companies in terms of increasing focus on protein yield, initiatives to increase sales, and enhancing distribution base. The report has been concluded with strategic recommendations for players already present in the market and new players planning to enter the market, which could help them in the long run. Persistence Market Research (PMR) is a third-platform research firm. Our research model is a unique collaboration of data analytics and market research methodology to help businesses achieve optimal performance. To support companies in overcoming complex business challenges, we follow a multi-disciplinary approach. At PMR, we unite various data streams from multi-dimensional sources. By deploying real-time data collection, big data, and customer experience analytics, we deliver business intelligence for organizations of all sizes.

Loading Jena Bioscience GmbH collaborators
Loading Jena Bioscience GmbH collaborators