News Article | May 17, 2017
The 1.3-kb GCaMP6 coding region was PCR amplified from the pGP-CMV-GCaMP6s plasmid (Addgene)33. The amplified DNA was then inserted into the plant expression vector (the HBT-HA-NOS plasmid)45 to generate the HBT-GCaMP6-HA construct. The HBT-GCaMP6-HA construct was inserted into the binary vector pCB302 (ref. 46) to generate the HBT-GCaMP6-HA transgenic plants using the Agrobacterium (GV3101)-mediated floral-dip method47. Transgenic plants were selected by spraying with the herbicide BASTA. The construct expressing HY5–mCherry was used as a control for protoplast co-transfection and nucleus labelling, and was obtained from J.-G. Chen48. NLS-Td-Tomato was used as a control for protoplast co-transfection and nucleus labelling, and was obtained from X. Liu. NIR-LUC was constructed as described previously11. UBQ10-GUS is a control for protoplast co-transfection and internal control; all HBT-CPKac-Flag-NOS expression plasmids have been described previously23. To construct HBT-CPK-GFP-NOS, the coding regions of the CPK10, CPK30 and CPK32 cDNA were amplified and then cloned into the HBT-GFP-NOS plasmid23. HBT-CPK10(M141G)-Flag was generated by site-directed mutagenesis of the HBT-CPK10-Flag construct. To complement the cpk10 cpk30/+ mutant, a 5.5-kb DNA fragment including the promoter region (3 kb) and the coding region of CPK10 was amplified from genomic DNA, which was then cloned into the plasmid HBT-HA-NOS and mutagenized to generate pCPK10-CPK10(M141G)-HA-NOS. The pCPK10-CPK10(M141G)-HA-NOS construct was inserted into pCB302 and transformed into cpk10 cpk30/+ mutant plants using the Agrobacterium (GV3101)-mediated floral-dip method47. At the T generation, homozygous single-copy insertion lines were screened for the cpk10 cpk30 double mutant carrying pCPK10-CPK10(M141G)-HA-NOS to obtain the 3MBiP-inducible icpk10,30 double mutant, which rescued the embryo lethality of the cpk10 cpk30 double mutant. The 3MBiP-inducible icpk10,30,32 triple mutant expressing CPK10(M141G)-HA (designated icpk) was generated by genetic cross to cpk32 and confirmed by molecular analyses. To construct 35SΩ-NLP6-MYC or 35SΩ-NLP7-MYC in the pCB302 binary plasmid with hygromycin B selection, the β-glucuronidase (GUS) gene in the 35SΩ-GUS plasmid49 was replaced with the DNA fragment encoding the full-length NLP6 or NLP7 fused to 6 copies of the MYC epitope tag in the HBT-NLP6-MYC or HBT-NLP7-MYC plasmid12. The NLP6-MYC and NLP7-MYC transgenic plants were generated by Agrobacterium (GV3101)-mediated transformation by floral dip and hygromycin B resistance selection. To construct HBT-NLP7-HA and HBT-NLP7-GFP, the 2.9 kb coding region of the NLP7 cDNA was amplified and then cloned into the HBT-NOS plasmid. HBT-NLP7(S205A)-HA and HBT-NLP7(S205A)-GFP were generated by site-directed mutagenesis. A 7.9-kb genomic DNA fragment of NLP7 was cloned into the pUC plasmid and fused with GFP at the C terminus to generate pNLP7-NLP7-GFP. The pNLP7-NLP7(S205A)-GFP construct was generated by site-directed mutagenesis. pNLP7-NLP7-GFP or pNLP7-NLP7(S205A)-GFP was then inserted into pCB302 and introduced into nlp7-1 mutant plants using the Agrobacterium (GV3101)-mediated floral-dip47 method for complementation analyses. To construct UBQ10-CPK10KM-YN and UBQ10-NLP7-YC, the coding regions of CPK10(KM), NLP7, YFP-N terminus and YFP-C terminus were amplified by PCR and cloned into the UBQ10-GUS plasmid. To construct pET14-NLP7-N(1-581)-HIS and pET14-NLP7-N(S205A)-HIS for protein expression, the N-terminal coding region of NLP7 and NLP7(S205A) were amplified from HBT-NLP7-HA and HBT-NLP7(S205A)-HA. All constructs were verified by sequencing. The primers used for plasmid construction and site-directed mutagenesis are listed in Supplementary Table 3. Arabidopsis ecotype Columbia (Col-0) was used as the wild type. The cpk mutants were obtained from Arabidopsis Biological Resource Centre (ABRC)50. Homozygous T-DNA lines were identified using CPK gene-specific primers and T-DNA left-border primers. The gene-specific primers used are listed in the Supplementary Table 4. Double mutants were obtained by genetic crosses between cpk10-1, cpk30-1 and cpk32-1, and confirmed by PCR. For RT–PCR analysis of cpk single mutants, around 30 plants were grown on the Petri dish (150 mm × 15 mm) containing 100 ml of 1/2 × MS medium salt, 0.1% MES, 0.5% sucrose, 0.7% phytoagar under constant light (150 μmol m−2 s−1) at 23 °C for 7 days. Samples were collected for RT–PCR analysis. To generate icpk, cpk32-1 was crossed to icpk10,30. F plants were first screened for resistance to BASTA and then confirmed by genotyping (primers listed in Supplementary Table 4) for the homozygous cpk10 cpk30 cpk32 triple mutants. The homozygous icpk plants were isolated with no segregation for BASTA resistance in F plants. To demonstrate embryo lethality in cpk10 cpk30 mutants, cpk10 cpk30/+ plants were grown at a photoperiod of 16 h (light)/8 h (dark) (100 μmol m−2 s−1) at 23 °C/20 °C. Siliques were opened using forceps and needles under a dissecting microscope (Leica MZ 16F). Images were acquired and processed using IM software and Adobe Photoshop (Adobe). To obtain nitrate-free mesophyll protoplasts, around 16–20 plants were grown on a Petri dish (150 mm × 15 mm) containing 100 ml of nitrogen-free 1 × MS medium salt, 0.1% MES, 1% sucrose, 0.7% phytoagar, 2.5 mM ammonium succinate and 0.5 mM glutamine, pH 6 under a photoperiod of 12 h (light)/12 h (dark) (75 μmol m−2 s−1) at 23 °C/20 °C for 23–28 days. Mesophyll protoplasts were isolated from the second and the third pair of true leaves following the mesophyll protoplast isolation protocol46. To monitor plant growth without exogenous nitrogen source after germination, 30 seedlings were germinated and grown on a basal medium11 (10 mM KH PO /KH PO , 1 mM MgSO , 1 mM CaCl , 0.1 mM FeSO -EDTA, 50 μM H BO , 12 μM MnSO ·H O, 1 μM ZnCl , 1 μM CuSO ·5H O, 0.2 μM Na MoO ·2H O, 0.1% MES and 0.5% sucrose, pH 5.8) with 1% phytoagar under constant light (150 μmol m−2 s−1) at 23 °C for 4 days. Photos were taken at different days (days 1–4) using a dissecting microscope (Leica MZ 16F) with IM software. To analyse the specific plant growth programs in response to different exogenous nitrogen sources at different concentrations, seedlings were germinated and grown on basal medium for 4 days as described above, and then transferred to the basal medium with 0.1, 0.5, 1, 5 or 10 mM KNO , NH Cl, glutamine or KCl for an additional 1–7 days. For gene expression analyses with RT–qPCR and RNA-seq, 10 seedlings were germinated in one well of the 6-well tissue culture plate (Falcon) with 1 ml of the basal medium supplemented with 2.5 mM ammonium succinate as the sole nitrogen source. Plates were sealed with parafilm and placed on the shaker at 70 r.p.m. under constant light (45 μmol m−2 s−1) at 23 °C for 7 days. Before nitrate induction, seedlings were washed three times with 1 ml basal medium. Seedlings were treated in 1 ml of basal medium with KCl or KNO for 15 min. Seedlings were then harvested for RNA extraction with TRIzol (Thermo Fisher Scientific). To block the kinase activity of CPK10(M141G), seedlings were pre-treated with 10 μM 3MBiP in the basal medium for 2 min, and then treated with KCl or KNO for 15 min. For Ca2+ channel blockers and Ca2+ sensor inhibitors assays, seedlings were pre-treated with 2 mM LaCl , 2 mM GdCl , 250 μM W5 or 250 μM W7 in 1 ml of basal medium for 20 min, and then induced by 0.5 mM KCl or KNO for 15 min. To monitor root morphology, seedlings were germinated and grown on a basal medium supplemented with 2.5 mM ammonium succinate and 1% phytoagar under constant light (150 μmol m−2 s−1) at 23 °C for 3 days. Plants were then transferred to the basal medium supplemented with 1 μM 3MBiP and 5 mM KNO , 2.5 mM ammonium succinate, 5 mM KCl or 1 mM glutamine and grown for 5–8 days. After seedling transfer, 1 ml of 1 μM 3MBiP was added to the medium every 2 days. To monitor lateral root developmental stages, seedlings were monitored using a microscope (Leica DM5000B) with a 20× objective lens according to the protocol described previously41. To measure the primary and lateral root length, pictures were taken using a dissecting microscope (Leica MZ 16 F) with IM software and analysed by ImageJ. To compare the shoot phenotype, 8-day-old seedlings were cut above the root–shoot junction to measure the shoot fresh weight and acquire images. To analyse the cpk single-mutant phenotype, plants were germinated and grown on ammonium succinate medium for 3 days and then transferred to basal medium plates supplemented with 5 mM KNO for 6 days. To analyse double mutants in response to 3MBiP, plants were transferred to basal medium plates supplemented with 5 mM KNO and 1 μM 3MBiP for 6 days, and 3MBiP was reapplied every 2 days. Individual 9-day-old seedlings (n = 12) were collected to measure fresh weight and acquire images. To characterize the shoot phenotype of nlp7-1 and the complementation lines, around 20 seeds were germinated on the Petri dish (150 mm × 15 mm) containing 100 ml of nitrogen-free 1 × MS medium salt (Caisson), 0.1% MES, 1% sucrose, 0.7% phytoagar and 25 mM KNO medium pH 5.8 under a 16 h (light)/8 h (dark) photoperiod (100 μmol m−2 s−1) at 18 °C and grown for 21 days. The shoots were collected for measurement of fresh weight and acquisition of images. For analyses of the shoot phenotype in icpk, seeds were germinated and grown on the ammonium succinate basal medium plate for 3 days and then transferred to the same medium supplemented with 1 μM 3MBiP. The inhibitor 3MBiP (5 ml of 1 μM) was reapplied on the medium twice during the growth. Two transgenic seedlings expressing apoaequorin22 were germinated and grown in one well of a 12-well tissue culture plate (Falcon) with 0.5 ml of the basal medium supplemented with 2.5 mM ammonium succinate for 6 days. Individual plants were transferred to a luminometer cuvette filled with 100 μl of the reconstitution buffer (2 mM MES pH 5.7, 10 mM CaCl , and 10 μM native coelenterazine from NanoLight Technology) and incubated at room temperature in the dark overnight. The emission of photons was detected every second using the luminometer BD Monolight 3010. The measurement was initiated by injection of 100 μl 20 mM KCl, 20 mM KNO , 200 nM flg22 or ultrapure water into the cuvettes. Luminescence values were exported and processed using Microsoft Excel software. For Ca2+ imaging in protoplasts, mesophyll protoplasts (2 × 105) in 1 ml buffer were co-transfected with 70 μg HBT-GCaMP6 and 50 μg HBT-HY5-mCherry plasmid DNA. Transfected protoplasts were incubated in 5 ml of WI buffer45 for 4 h. Before time-lapse recording, a coverslip was placed on a 10-well chamber slide covering three-quarters of a well, and placed on the microscope stage. Mesophyll protoplasts co-expressing GCaMP6 and HY5–mCherry (2 × 104 protoplast cells) were spun down for 1 min at 100g. WI-Ca2+ buffer (WI buffer plus 4 mM CaCl ) (0.5 μl) with different stimuli (40 mM KCl, 40 mM KNO or 40 mM NH Cl) or 80 mM Ca2+ chelator (EGTA) were added into 1.5 μl of concentrated mesophyll protoplasts in WI buffer. The final concentration of each stimulus was 10 mM KCl, 10 mM KNO , 10 mM NH Cl or 20 mM EGTA in the solution. The stimulated protoplasts were immediately loaded onto the slide and imaged via the Leica AF software on a Leica DM5000B microscope with the 20× objective lens. The exposure time for GCaMP6 was set at 1 s and recorded every 2 s to generate 199 frames. The exposure time was set at 45 ms for the bright field and 1 s for the mCherry signal. The fluorescence intensity was determined with the region of interest (ROI) function for each protoplast. The intensity data were exported and processed using Microsoft Excel software. The images were exported and processed using Adobe Photoshop software. To make a video, individual images were cropped using Adobe Photoshop software and saved in JPEG format. The videos were generated using ImageJ with the cropped images. For Ca2+ imaging with the GCaMP6 transgenic seedling cotyledons, 5 seedlings were germinated in 1 well of a 6-well tissue culture plate (Falcon) with 1 ml of the basal medium supplemented with 2.5 mM ammonium succinate for 7 days. A chamber was made on microscope slides between two strips of the invisible tape (0.5 cm × 3 cm) and filled with 150 μl of the basal medium. A cotyledon of the 7-day-old seedling was cut in half using a razor blade and embedded in the medium. A thin layer of cotton was placed on top of the cotyledon to prevent moving. The coverslip was placed on the sample and fixed by another two strips of the invisible tape. The cotyledon was allowed to recover on the slide for 10 min. Confocal imaging was acquired using the Leica laser scanning confocal system (Leica TCS NT confocal microscope, SP1). The mesophyll cells in the cotyledon were targeted for Ca2+ imaging at the focal point. Basal medium (200 μl) with 10 mM KCl, 10 mM KNO or 20 mM EGTA was loaded along one edge of the coverslip. A Kimwipes tissue on the opposite edge was used to draw the buffer into the chamber. To record fluorescence images, the excitation was provided at 488 nm and images were collected at emission 515–550 nm. The scanning resolution was set at 1,024 × 1,024 pixels. Images were captured every 10 s and averaged from two frames. In total, 80 images were collected and processed using Adobe Photoshop software. A video was generated with collected images using the method described above. For Ca2+ imaging with the GCaMP6 transgenic seedling at the root tip and the elongated region of roots (around the middle region of the root), 10 seedlings were germinated and grown on the tissue culture plate (Falcon) with the basal medium and 1% phytoagar under constant light (150 μmol m−2 s−1) at 23 °C for 4 days. The images were obtained using Leica laser scanning confocal system as described above for cotyledon Ca2+ imaging. In total, 33 images were collected and processed using Adobe Photoshop software. A video was generated with collected images using the method described above. Time-course, specificity and dosage analyses of NIR-LUC activity in response to nitrate induction was carried out in mesophyll protoplasts (2 × 104 protoplasts in 100 μl) co-transfected with 10 μg NIR-LUC and 2 μg UBQ10-GUS (as the internal control) and incubated in WI buffer45 for 4 h, and then induced by 0.5 mM KCl, KNO , NH + or Gln or different concentrations of KNO for 2 h. For time-course analysis, the fold change is calculated relative to the value of KCl treatment at each time point. For the nitrate-sensitized functional genomic screen, nitrate-free mesophyll protoplasts (2 × 104 protoplasts in 100 μl) were co-transfected with 8 μg HBT-CPKac (constitutively active CPK) or a control vector, 10 μg NIR-LUC and 2 μg UBQ10-GUS plasmid DNA, and incubated for 4 h to allow CPKac protein expression. To investigate the functional relationship between CPK10ac and NLP7 in nitrate signalling, nitrate-free mesophyll protoplasts (4 × 104 protoplasts in 200 μl) were co-transfected with 8 μg NIR-LUC and 2 μg UBQ10-GUS plasmid DNA, as well as 5 μg HBT-CPK10ac, HBT-CPK10ac(KM) or a control vector, or HBT-NLP7 or HBT-NLP7(S205A) in different combinations supplemented with 5 μg control vector to reach a total of 20 μg per transfection reaction, and incubated for 4 h for protein expression. Protoplasts were then induced with 0.5 mM KCl or KNO for 2 h. The luciferase and GUS assay were carried out as described before45. The expression levels of NLP7–HA and CPK–Flag or CPK10ac–Flag in protoplasts were monitored by immunoblot with anti-HA-peroxidase (Roche, 11667475001; 1:2,000) and anti-Flag-HRP (Sigma, A8592; 1:2,000) antibodies, respectively. Expression vectors were transformed into Rosetta 2 (DE3) pLysS Competent Cells (Novagen). Cells were induced by 1 mM of IPTG when OD reached 0.6, and proteins were expressed at 18 °C for 18 h. Affinity purification was carried out using HisTrap columns (GE Healthcare) and the ÄKTA FPLC system. Purified proteins were buffer exchanged into PBS using PD-10 Desalting Columns (GE Healthcare), and then concentrated by Amicon Ultra-4 Centrifugal Filter Unit with Ultracel-10 membrane (EMD Millipore). Around 106 protoplasts were incubated in WI buffer (5 ml) in Petri dishes (9 × 9 cm) for 4 h before induction with 10 mM KCl or KNO for 10 min. Protoplasts were harvested and lysed in 200 μl of extraction buffer: 150 mM NaCl, 50 mM Tris-HCl pH 7.5, 5 mM EDTA, 1% Triton X-100, 1× protease inhibitor cocktail (Complete mini, Roche) and 1 mM DTT. The protein extract supernatant was obtained after centrifugation at 18,000g for 10 min at 4 °C. Total proteins (20 μg) were loaded on 8% SDS–PAGE embedded with or without 0.5 mg ml−1 histone type III-S (Sigma) as a general CPK phosphorylation substrate23. The gel was washed three times with washing buffer (25 mM Tris-HCl pH 7.5, 0.5 mM DTT, 5 mM NaF, 0.1 mM Na VO , 0.5 mg ml−1 BSA and 0.1% Triton X-100), and incubated for 20 h with three changes in the renaturation buffer (25 mM Tris-HCl pH 7.5, 0.5 mM DTT, 5 mM NaF and 0.1 mM Na VO ) at 4 °C. The gel was then incubated in the reaction buffer (25 mM Tris-HCl pH 7.5, 2 mM EDTA, 12 mM MgCl , 1 mM CaCl , 1 mM MnCl , 1 mM DTT and 0.1 mM Na VO ) with or without 20 mM EGTA at room temperature for 30 min. The kinase reaction was performed for 1 h in the reaction buffer supplemented with 25 μM cold ATP and 50 μCi [γ-32P]ATP with or without 20 mM EGTA. The reaction was stopped by extensive washes in the washing buffer (5% trichloroacetic acid and 1% sodium pyrophosphate) for 6 h. The protein kinase activity was detected on the dried gel using the Typhoon imaging system (GE Healthcare). 1-Isopropyl-3-(3-methylbenzyl)-1H-pyrazolo[3,4-d]pyrimidin-4-amine (3MBiP) was synthesized using the same procedures as those for a close structural analogue, 3MB-PP1 (ref. 39), with comparable yields, except that iso-propylhydrazine was substituted for tert-butylhydrazine. 1H NMR (400 MHz, DMSO-d ) δ 8.12 (s, 1H), 7.15 (d, J = 7.6 Hz, 1H), 7.08 (s, 1H), 7.00 (t, J = 7.5 Hz, 2H), 4.96 (p, J = 6.7 Hz, 1H), 4.31 (s, 2H), 2.24 (s, 3H), 1.44 (d, J = 6.7 Hz, 6H). 13C NMR (100 MHz, DMSO-d ) δ 158.41, 155.78, 153.69, 143.12, 139.56, 137.85, 129.51, 128.79, 127.30, 125.87, 98.92, 48.18, 40.10, 33.70, 22.23, 21.54. ESI–MS calculated for C H N [M + H]+ is 282.2, found 282.7. For in vitro kinase assay with CPK10(M141G)–Flag or CPK10–Flag, 4 × 104 protoplasts expressing CPK10(M141G)–Flag or CPK10–Flag were lysed in 200 μl immunoprecipitation buffer that contained 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 5 mM EDTA, 1 mM DTT, 2 mM NaF, 2 mM Na VO , 1% Triton X-100 and 1× protease inhibitor cocktail (Complete mini, Roche). Protein extracts were incubated with 0.5 μg anti-Flag antibody (Sigma, F1804) at 4 °C for 2 h and an additional 1 h with protein G Sepharose beads (GE Healthcare). The immunoprecipitated kinase protein was washed three times with immunoprecipitation buffer and once with kinase buffer (20 mM Tris-HCl pH 7.5, 15 mM MgCl , 1 mM CaCl and 1 mM DTT). Kinase reactions were performed for 1 h in 25 μl kinase buffer containing 1 μg histone (Sigma H5505 or H4524), 50 μM cold ATP and 2 μCi [γ-32P]ATP. To block the CPK10(M141G)–Flag kinase activity, 1 μM 3MBiP or DMSO as a control was added in the 25 μl kinase buffer for 2 min before performing the kinase reaction. The reaction was stopped by adding SDS–PAGE loading buffer. After separation on a 12% SDS–PAGE gel, the protein kinase activity was detected on the dried gel using the Typhoon imaging system. For the in vitro kinase assay with CPK10(M141G)–HA isolated from icpk10,30 seedlings, 12 7-day-old seedlings grown in 2 wells of a 6-well-plate with 1 ml medium (0.5 × MS, 0.5% sucrose and 0.1% MES pH 5.7) were grounded in liquid nitrogen into powder and lysed in 200 μl of immunoprecipitation buffer. The CPK10(M141G)–HA protein was immunoprecipitated with the anti-HA antibody (Roche, 11666606001) and protein G Sepharose beads. In vitro kinase assay with CPK10(M141G)–HA proteins was carried out as described above. For the in vitro kinase assay with the subgroup III CPKs, Flag-tagged CPK7, CPK8, CPK10, CPK10(KM) (K92M, a kinase-dead mutation in the conserved ATP binding domain), CPK13, CPK30 and CPK32 were expressed in 105 protoplasts and purified with 1 μg anti-Flag antibody conjugated to protein G Sepharose beads as described above. CPK11–Flag from subgroup I was used as a negative control to demonstrate the specificity of NLP7 as a substrate for only subgroup III CPKs. NLP7–HIS (~1 μg) purified from Escherichia coli or histone type III-S (2 μg) was used as substrate in the in vitro kinase assay. Kinase reactions were performed for 1 h at 28 °C in 25 μl kinase buffer containing 5 μM cold ATP and 6 μCi [γ-32P]ATP, which greatly enhanced the CPK activity. To reduce the background caused by free [γ-32P]ATP in the gel, 50 μM cold ATP was added to the kinase reaction before sample loading in 10% (NLP7–HIS) or 12% (HIS) SDS–PAGE gel. To demonstrate that the kinase activities of CPK10, CPK30 and CPK32 were Ca2+-dependent, 4 × 104 (CPK10 or CPK10ac) or 105 (CPK30, CPK32, CPK30ac or CPK32ac) protoplasts expressing CPKs for 12 h instead of 6 h (to increase the yield of CPK proteins) were lysed in 200 μl (CPK10) or 400 μl (CPK30 or CPK32) of immunoprecipitation buffer. The CPK proteins were immunoprecipitated with anti-Flag antibody (0.5 μg for CPK10 or CPK10ac, and 2 μg for CPK30, CPK32, CPK30ac or CPK32ac) conjugated to protein G Sepharose beads. The immunoprecipitated CPKs were washed three times with immunoprecipitation buffer and twice with EGTA kinase buffer (20 mM Tris-HCl pH 7.5, 15 mM MgCl , 15 mM EGTA and 1 mM DTT). Kinase reactions were performed for 1 h at 28 °C in 25 μl kinase buffer or EGTA kinase buffer containing 5 μM cold ATP and 6 μCi [γ-32P]ATP and purified NLP7–HIS (~1 μg), NLP7-N (1–581 amino acids) (~0.8 μg), NLP7-N(S205A) (~0.8 μg), or histone type III-S (2 μg). After performing the kinase reaction, 50 μM cold ATP was added to reduce the background caused by free [γ-32P]ATP. The reaction was stopped by adding SDS–PAGE loading buffer. After separation on a 12% SDS–PAGE gel (histone type III-S) or 10% (NLP7–HIS or NLP7-N–HIS) SDS–PAGE gel, the protein kinase activity was detected on the dried gel using the Typhoon imaging system. Substrate was stained with InstantBlue Protein Stain (C.B.S. Scientific). The expression levels of CPK or CPKac proteins were monitored by immunoblot with anti-Flag-HRP (Sigma, A8592; 1:4,000) antibody. CPKac proteins without the Ca2+-binding EF-hand domains provided constitutive kinase activities that were insensitive to EGTA. The sensitivity of CPK10, CPK30 and CPK32 to EGTA in kinase assays demonstrated their functions as Ca2+ sensors in nitrate signalling, which was further supported by the lack of NLP7–HA phosphorylation and the nuclear retention of NLP7–GFP in icpk mutant cells. Importantly, NLP7(S205A) lost nitrate-induced phosphorylation, nuclear localization, NIR-LUC activation, and endogenous target gene activation in wild-type protoplasts and seedlings. RNA isolation, RT–PCR and RT–qPCR were carried out as described previously11. The primers used for RT–PCR and RT–qPCR are listed in Supplementary Table 5. TUB4 was used as a control in wild-type and cpk mutants. The relative gene expression was normalized to the expression of UBQ10. Triplicate biological samples were analysed with consistent results. We chose the early time point to minimize secondary target genes and the complexity that negative feedback would have introduced, including indirect effects from assimilation of nitrate and the subsequent activation of transcriptional repressors1, 3, 4, 8, 10, 13. Seven-day-old wild-type and icpk seedlings were pretreated with 10 μM 3MBiP for 2 min and then treated for 15 min with either 10 mM KCl or 10 mM KNO . Total RNA (0.5 μg) was used for preparing the library with the Illumina TruSeq RNA sample Prep Kit v2 according to the manufacturer’s guidelines with 9 different barcodes (triplicate biological samples). The libraries were sequenced for 50 cycles on an Illumina HiSeq 2500 rapid mode using two lanes of a flow cell. The sequencing was performed at MGH Next Generation Sequencing Core facility (Boston, USA). Fastq files, downloaded from the core facility, were used for data analysis. The quality of each sequencing library was assessed by examining fastq files with FastQC. Reads in the fastq file were first aligned to the Arabidopsis genome, TAIR10, using Tophat51. HTSeq52 was used to determine the reads per gene. Finally, DESeq2 (ref. 53) analysis was performed to determine differential expression54. For HTSeq-normalized counts in each sample, differentially expressed genes were determined for wild-type KNO versus wild-type KCl and icpk KNO versus wild-type KNO . The differential expression analysis in DESeq2 uses a generalized linear model of the form where counts K for gene i, sample j are modelled using a negative binomial (NB) distribution with fitted mean μ and a gene-specific dispersion parameter α . The fitted mean is composed of a sample-specific size factor s and a parameter q proportional to the expected true concentration of fragments for sample j. The coefficients β give the log fold changes for gene i for each column of the model matrix X. Results were imported into Microsoft Excel for filtering. To generate a list to minimize false positives of primary nitrate-responsive genes in the wild type, we applied a relatively high stringency, q ≤ 0.05 cut-off, followed by a log ≤ −1 or ≥ 1 cut-off. To generate a heatmap, we performed agglomerative hierarchical clustering on genes with Gene Cluster 3.0 (ref. 55) using Correlation (uncentred) as the similarity metric and single linkage as the clustering method. Java Treeview56 was used to visualize the results of the clustering. To obtain a list of enriched gene functions, we used the Classification SuperViewer Tool on the BAR website (http://bar.utoronto.ca/ntools/cgi-bin/ntools_classification_superviewer.cgi) with the MapMan classification source option. Analyses of enriched functional categories with nitrate upregulated and downregulated genes were performed using the MapMan classification source option on the Classification SuperViewer Tool with manual annotation based on literature. The fold enrichment is calculated as follows: (number in class /number of total )/ (number in class / number of total ). The P value is calculated in Excel using a hypergeometric distribution test. The data in Extended Data Fig. 4c and d were sorted by fold enrichment with a P < 0.05 cut-off. For the biological duplicate RNA-seq experiments for identifying NLP7 target genes in the mesophyll protoplast transient expression system, 500 μg HBT-NLP7-HA, HBT-NLP7(S205A)-HA or control plasmid DNA was transfected into 106 protoplasts and incubated for 4.5 h. Total RNA (0.5 μg) was used to construct the libraries with six different barcodes (biological duplicate samples) as described above. The sequencing result was performed and analysed as described above. Differentially expressed genes were determined with DESeq2 on NLP7 versus Ctl (Control) and NLP7(S205A) versus Ctl. Results were imported into Microsoft Excel for filtering (log ≥ 1 cut-off) and generating heatmaps. Transgenic seedlings expressing NLP6–MYC or NLP7–MYC were germinated and grown in basal medium containing 0.5 mM ammonium succinate as a sole nitrogen source (0.01% MES-KOH, pH 5.7) for 4 days at 23 °C under continuous light (60 μmol m−2 s−1). After replacement with fresh medium supplemented with 10 mM KCl or KNO , the seedlings were collected after incubation for 5, 10 or 30 min. To examine the effects of Ca2+ channel blockers and Ca2+ sensor inhibitors, the 4-day-old seedlings were placed in fresh basal medium supplemented with 2 mM LaCl , 2 mM GdCl , 250 μM W5 or 250 μM W7 for 20 min and induced by 10 mM KCl or KNO . The seedlings were weighed, frozen in liquid nitrogen and ground using a Multibeads Shocker (Yasui Kikai). The ground samples were suspended in 20 volume of 1× Laemmli sample buffer supplemented with twice the concentration of EDTA-free protease inhibitor cocktail (Roche) and heated at 95 °C for 30 s. Samples were then spun down and the supernatant was subjected to SDS–PAGE and immunoblotting with anti-MYC (Millipore, 05-419; 1:1,000) and anti-histone H3 (Abcam, ab1791; 1:5,000) antibodies. For calf intestinal alkaline phosphatase (CIP) treatment, proteins in 1.2-fold CIP buffer (60 mM Tris-HCl pH 8.0, 120 mM NaCl, 12 mM MgCl , 1.2 mM DTT, 2.4-fold concentration of EDTA-free Protease Inhibitor Cocktail) were mixed with CIP solution (New England Biolabs, M0290, 10 U μl−1) at a ratio of 5 (CIP buffer):1 (CIP solution) and incubated at 37 °C for 30 min. Heat-inactivated CIP was mixed as a control treatment. The reactions were stopped by adding an equal volume of 2× Laemmli sample buffer and heating at 95 °C for 30 s. To demonstrate that nitrate-induced NLP7 phosphorylation was abolished in icpk by protein mobility shift in SDS–PAGE, 4 × 104 protoplasts isolated from wild-type or icpk seedlings were transfected with 20 μg NLP7–HA or NLP7(S205A)–HA. To block CPK10(M141G) activity in icpk, 10 μM 3MBiP was added in the incubation buffer (WI) after transfection. After expressing protein for 4.5 h, protoplasts were induced by 10 mM KCl or KNO for 15 min. Protoplasts were spun down and re-suspended in 40 μl 1× Laemmli sample buffer. Samples (10 μl) were separated in a 6% SDS–PAGE resolving gel without a stacking gel layer. After transferring proteins to the PVDF membrane, the NLP7 (wild-type and S205A) proteins were detected with anti-HA-peroxidase (Roche, 11667475001; 1:2,000). RuBisCo was detected by an anti-rubisco antibody (Sigma, GW23153; 1:5,000) as a loading control. Transformation of T87 cell suspension culture derived from a seedling of A. thaliana L. (Heynh.) ecotype Columbia57 was conducted with the 35SΩ-NLP7-MYC construct in the pCB302 binary plasmid carrying the hygromycin B selection marker gene. Transformants mediated by Agrobacterium (GV3101) were selected on agar plates (JPL medium, 3 g l−1 gellun gum, 500 mg l−1 carbenicillin and 20 mg l−1 hygromycin), and the transformants were maintained in liquid JPL medium as described previously57. T87 cells expressing NLP7–MYC were incubated in nitrogen-free JPL liquid medium for 2 days, and then 10 mM KNO was added into the medium. After 30 min treatment, the T87 cells (approximately 4 g frozen weight) were frozen in liquid nitrogen and homogenized with Multi-beads Shocker (Yasui Kikai) in 10 ml of the buffer that contained 25 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.1% NP-40, 10% glycerol, 1× Complete Protease Inhibitor Cocktail and 1× PhosSTOP (Roche). Cell lysates obtained were incubated with anti-MYC antibodies crosslinked to Dynabeads (Invitrogen). Trapped proteins were eluted by 1× Laemmli sample buffer and separated by SDS–PAGE. Gel pieces containing NLP7–MYC were recovered and subjected to in-gel double digestion with trypsin (10 ng μl−1) and chymotrypsin (10 ng μl−1) (Promega). NanoLC–ESI-MS/MS analysis was performed as described previously58, 59 with minor modifications. To analyse NLP7 nuclear retention triggered by nitrate in protoplasts, nitrate-free mesophyll protoplasts (4 × 104 protoplasts in 200 μl) were co-transfected with 20 μg NLP7–GFP or NLP7(S205A)–GFP and 10 μg HBT-HY5-mCherry plasmid DNA and incubated for 6 h. Mesophyll protoplasts were spun down for 1 min at 100g. WI buffer with 10 mM KCl or KNO was added into mesophyll protoplasts for 30 min. The treated protoplasts were loaded onto slides and imaged with the 20× objective lens on a Leica DM5000B microscope operated with the Leica AF software. The images were collected and processed using Adobe Photoshop software. To analyse NLP7–GFP nuclear retention triggered by nitrate in transgenic lines, NLP7–GFP/nlp7-1 and NLP7(S205A)–GFP/nlp7-1 seedlings were germinated and grown on the basal medium supplemented with 2.5 mM ammonium succinate and with 1% phytoagar under constant light (150 μmol m−2 s−1) at 23 °C for 5 days. Plants were placed on the slide as described above and stimulated by 10 mM KNO . Confocal images were acquired as described for GCaMP6-based Ca2+ imaging in transgenic seedlings. To analyse CPK10, CPK30 and CPK32 nuclear localization in response to nitrate, nitrate-free mesophyll protoplasts (4 × 104 protoplasts in 200 μl) were co-transfected with 20 μg CPK10–GFP, CPK30–GFP or CPK32–GFP and 10 μg HBT-HY5-mCherry plasmid DNA and incubated for 12 h. Protoplasts were then treated with 10 mM KNO for 5 min. Confocal imaging was acquired using the Leica Application Suite X software on a Leica TCS SP8 (Leica) confocal microscope with the 40× objective lens. To obtain fluorescence images, the excitation was set to 489 nm (GFP) and 587 nm (mCherry), and images at emissions 508 nm (GFP) and 610 nm (mCherry) were collected. The scanning resolution was set to 1,024 × 1,024 pixels. The images were collected and processed using Adobe Photoshop software. To analyse NLP7–GFP nuclear retention in wild-type and icpk seedlings, nitrate-free mesophyll protoplasts (4 × 104 protoplasts in 200 μl) were co-transfected with 20 μg NLP7–GFP and 4 μg HBT-Td-Tomato plasmid DNA and incubated for 12–16 h. The transfected protoplasts were treated with inhibitor 10 μM 3MBiP 30 min before nitrate induction. Protoplasts were treated with 10 mM KNO for 15 min in the presence of 10 μM 3MBiP of WI buffer. The images were acquired as described above for the NLP7 nuclear retention in protoplasts. Nitrate-free mesophyll protoplasts (4 × 104 protoplasts in 200 μl) were co-transfected with 18 μg UBQ10-CPK10(KM)-YN, UBQ10-NLP7-YC, and 4 μg HBT-HY5-mCherry plasmid DNA, and incubated for 12–18 h. Protoplasts were then treated with 10 mM KNO for 2 h. Confocal images were acquired as described above for CPK localization in response to nitrate. The chosen sample sizes for all experiments were empirically determined by measuring the mean and s.d. for the sample population in pilot experiments, and then calculated (the 1-sample Z-test method, two-sided test) with the aim to obtain the expected mean of less than 25% significant difference with the alpha value ≤ 0.05 and the power of the test ≥ 0.80. For multiple comparisons, data were first subjected to one-way or two-way ANOVA, followed by Tukey’s multiple comparisons test to determine statistical significance. To compare two groups, a Student’s t-test was used instead. To compare wild-type and icpk lateral root development, data were categorized into two groups, and then subjected to a chi-square test, as indicated in the figure legends. Experiments were not randomized and investigators were not blinded to allocation during experiments and outcome assessment. RNA-seq data are available at the Gene Expression Omnibus (GEO) under accession number GSE73437. The Source Data for blots, gels and histograms are provided in the Supplementary Information. All other data are available from the corresponding author upon reasonable request.
News Article | July 11, 2017
MADISON, Wis.--(BUSINESS WIRE)--Promega Corporation has become the first major forensic manufacturer to achieve third-party certification of the published ISO 18385 standard to minimize the risk of human DNA contamination in products used to collect, store and analyze biological material for forensic purposes. This Forensic DNA Grade certification comes following a recent onsite audit by international certification agency Bureau Veritas. “This critical milestone is the best way for forensic laboratories to ensure that products claimed to be produced under published ISO 18385 guidelines are indeed meeting those expectations,” said Charles York, Vice President, Manufacturing Operations at Promega. “As with our ISO 9001 and 13485 certifications, this certification further exemplifies Promega’s commitment to our customers, to our business, and to all those who rely on and benefit from the use of our products.” Promega products manufactured in alignment with the ISO 18385 standard will include a “Forensic Grade” certification logo. The new international standard was published on February 1, 2016, to improve the quality of forensic DNA testing by minimizing the risk of human DNA contamination during the manufacturing process. Two Promega scientists were selected by the United States Technical Advisory Group to represent the U.S. on the development of ISO 18385. They worked with forensic leaders from the Arkansas State Crime Laboratory and the National Institute of Forensic Science, Australia/New Zealand. ISO 18385 is based upon the Australian Standard. ISO 18385 certification for Promega culminated significant work by numerous departments throughout the organization including R&D, Operations, Marketing and Quality Assurance/Regulatory Affairs to ensure that processes meet the new standard requirements. This includes revised quality control testing and certificates of analysis, as necessary, and also treating plastics for the forensic product line, such as primary packaging materials and consumables, with ethylene oxide. “Promega is continually striving to not only meet and exceed the most current quality standards for our customers, but to also be a leader in the development of new and evolving guidelines that will best serve the forensic industry,” said Ron Wheeler, Senior Director of Quality Assurance and Regulatory Affairs. For more information about ISO 18385 and Promega efforts related to the standard, visit: www.promega.com/ForensicGradeFAQ Promega Corporation has provided products for DNA-based human identification for more than 25 years. Promega is a leader in providing innovative solutions and technical support to the life sciences industry. The company’s 3,500 products enable scientists worldwide to advance their knowledge in genomics, proteomics, cellular analysis, molecular diagnostics and human identification. Founded in 1978, the company is headquartered in Madison, WI, USA, with branches in 16 countries and over 50 global distributors. For more information about Promega, visit: www.promega.com
News Article | July 26, 2017
MADISON, Wis.--(BUSINESS WIRE)--Promega Corporation, a leading biotechnology manufacturer, released its 2017 Corporate Responsibility Report today highlighting continued sustainable expansion, community engagement, and initiatives supporting employee well-being to cultivate conscious long-term growth for the organization. Highlights from the 2017 Report include: Energy benchmarking completed in the last year from the Lawrence Berkeley National Laboratory shows that Promega facilities are in line with best-in-class laboratories on energy efficiency. In 2017, the company’s global distribution hub, the Kepler Center, saw its first full year in operation, contributing to an overall building footprint increase of 84% globally since 2008. More than 90% of this growth was in high-energy intensive laboratory, manufacturing and logistics spaces. Even with these increases, Promega has been able to hold carbon emissions constant relative to building footprint. Such evidence and outcomes validate a core Promega value of designing and building highly efficient facilities that are flexible, timeless and able to serve for many decades. As a global company, community engagement takes many forms. Each Promega location has the autonomy to focus on the unique needs of its community through an integrative and authentic approach. The goal is to provide meaningful global support at a local level by tapping into individual employee passions and providing the tools and resources to empower these employees to get involved in causes that are close to them. This is achieved through avenues such as paid time for volunteering, matching employee donations or collaborations with non-profits. The report explores the diverse organizations supported in the past year. To inspire and support employees, Promega creates unique and flexible workspaces, provides programs and space to encourage healthy lifestyles, promotes mindfulness and work-life balance, and invests in employee advancement. Predicated on decades of research and named the differentiating factor in a resilient culture with strong leadership, Promega has been proactively integrating emotional and social intelligence into the workplace and during the last year has introduced these principles to employees at various levels. For almost 40 years, Promega has integrated the values of corporate responsibility and sustainable business practices. The 2017 Corporate Responsibility Report, encompassing Promega philosophies and corporate mindset, product information, sustainability practices, work culture, and community outreach efforts, documents how the company continues to align business practices with positive social, environmental and business outcomes. Read the full report at www.promega.com/responsibility Promega Corporation is a leader in providing innovative solutions and technical support to the life sciences industry. The company’s 3,500 products enable scientists worldwide to advance their knowledge in genomics, proteomics, cellular analysis, drug discovery and human identification. Founded in 1978, the company is headquartered in Madison, WI, USA, with branches in 16 countries and over 50 global distributors. For more information about Promega, visit: www.promega.com
News Article | July 7, 2017
MADISON, Wis.--(BUSINESS WIRE)--Promega Corporation intends to seek US Food and Drug Administration approval and CE-IVD marking for a commercially available Microsatellite Instability (MSI) assay to assist oncologists and pathologists in determining colorectal cancer decisions. MSI status is a measure of mismatch repair deficiency commonly found in solid tumors. In addition to the colorectal cancer intended use, Promega intends to investigate additional possible applications for the MSI Diagnostic Test. The current Promega Research Use Only MSI Assay has been available and used in the market as part of Lab-Developed Tests since 2004. This patent protected technology is considered the gold standard molecular assay for detecting DNA mismatch repair deficiency. An FDA cleared Promega In Vitro Diagnostic version of this kit will extend the benefits of MSI detection to the specific needs of clinical labs. The MSI assay is reimbursable, has a fast turn-around time and most importantly there is a large body of evidence supporting use of MSI in colorectal cancer decisions. “The impact of having an MSI result on patient outcomes is becoming clearer each day,” says Randall Dimond, PhD, Chief Technical Officer at Promega. “Having a reliable, inexpensive functional assay with quick turnaround like MSI accessible to every pathology lab offers physicians a vital tool with which to make strategic decisions.” In clinical research, MSI is a biomarker that is proving to be increasingly important in understanding the most effective treatment methods for various types of cancers. MSI detection has captured the interest of researchers looking at DNA damage in many areas due to its reliability, simplicity and low cost. Recently the FDA announced fast-track approval of the Merck drug Keytruda based on MSI status for all advanced solid tumors. Leveraging this news, Promega is expanding its network of clinical researchers to better understand these new applications for MSI status in solid tumor types beyond colorectal cancer. “Following recent specific discussions with a number of labs, it is clear that they view the Promega MSI assay as an important tool for determining DNA mismatch repair deficiency that is a more straightforward measurement relative to next-generation sequencing,” says Heather Baird Tomlinson, PhD, Business Unit Leader - Molecular Diagnostics at Promega. “Our research showed that large marker panels and tumor mutation burden as defined by next-generation sequencing are excellent tools for research, but the impact of the information they provide is unclear, the assays are too costly, and the turnaround time is too long to be used routinely in a diagnostic setting in the near term. We will do everything we can to accelerate the delivery of the MSI technology for these labs in compliance with appropriate regulatory agencies.” Promega will rely on its extensive experience and expertise in designing fragment analysis and multiplex Polymerase Chain Reaction assays for the forensic industry to develop additional assays to expand and improve upon its current MSI product offering over the next several years. One example is a more sensitive MSI kit that is under development as an IVD at Promega Shanghai for CFDA approval in China. To learn more about the current Research Use Only Promega MSI Analysis system, visit www.promega.com/MSI. Promega Corporation is a leader in providing innovative solutions and technical support to the life sciences industry. The company’s 3,500 products enable scientists worldwide to advance their knowledge in genomics, proteomics, cellular analysis, drug discovery and human identification. Founded in 1978, the company is headquartered in Madison, WI, USA with branches in 16 countries and over 50 global distributors. For more information about Promega, visit www.promega.com.
News Article | May 9, 2017
MADISON, Wis.--(BUSINESS WIRE)--Join scientists, law enforcement professionals and forensic experts from around the world to learn about the latest developments in forensic DNA research, techniques and processes at the 28th International Symposium on Human Identification (ISHI), October 2-5, 2017, in Seattle, Washington. This symposium for forensic experts and suppliers is offered through Promega Corporation, a leader in providing innovative solutions and technical support to the life sciences industry. Promega Corporation has provided products for DNA-based human identification for over 20 years. The company’s 3,500 products enable scientists worldwide to advance their knowledge in genomics, proteomics, cellular analysis, drug discovery and human identification. Founded in 1978, the company is headquartered in Madison, WI, USA, with branches in 16 countries and over 50 global distributors. For more information about Promega, visit www.promega.com.
News Article | May 10, 2017
HEK293 cells stably transfected with the STF plasmid encoding the firefly luciferase reporter under the control of a minimal promoter, and a concatemer of 7 LEF/TCF binding sites32, were obtained from J. Nathans. Mouse L cells stably transfected with the STF plasmid and a constitutively expressed Renilla luciferase (control reporter) were obtained from C. Kuo. L cells transfected with a mouse WNT3A expression vector to produce conditioned media were obtained from the ATCC. A375, SH-SY5Y and A549 cells were stably transfected with the BAR plasmid encoding the firefly luciferase reporter under the control of a minimal prompter and a concatemer of 12 TCF/LEF binding sites and a constitutively expressed Renilla luciferase (control reporter) using a lentiviral-based approach33. All reporter cell lines were cultured in complete DMEM medium (Gibco) supplemented with 10% FBS, 1% penicillin, streptomycin, and l-glutamine (Gibco), at 37 °C and 5% CO and cultured in the presence of antibiotics for selection of the transfected reporter plasmid. C3H10T1/2 cells were obtained from the ATCC. Human primary MSCs were obtained from Cell Applications, Inc. Mouse primary MSCs were obtained from Invitrogen. Cell lines have not been tested for mycoplasma contamination. The coding sequence of B12 containing a C-terminal 6×His-tag was cloned into the pET28 vector (Novagen) for bacterial cytoplasmic protein expression. Protein expression was performed in transformed BL21 cells, expression was induced with 0.7 mM IPTG at an OD of 0.8 for 3–4 h. Cells were pelleted, lysed by sonication in lysis buffer (20 mM HEPES, pH 7.2, 300 mM NaCl, 20 mM imidazole), and soluble fraction was applied to Ni-NTA agarose (QIAGEN). After washing the resin with lysis buffer containing 500 mM NaCl, B12 was eluted with 300 mM imidazole, and subsequently purified on a Superdex 75 size-exclusion column (GE Healthcare) equilibrated in HBS (10 mM HEPES, pH 7.2, 150 nM NaCl). XWnt8 was purified from a stably transfected Drosophila S2 cell line co-expressing XWnt8 and mouse FZD8 CRD–Fc described previously4. Cells were cultured in complete Schneider’s medium (Thermo Fisher Scientific), containing 10% FBS and supplemented with 1% l-glutamine, penicillin and streptomycin (Gibco), and expanded in Insect-Xpress medium (Lonza). A complex of XWnt8 and FZD8 CRD–Fc was captured from the conditioned media on Protein A agarose beads (Sigma). After washing with 10 column volumes of HBS, XWnt8 was eluted with HBS containing 0.1% n-dodecyl-β-d-maltoside (DDM) and 500 mM NaCl, while the FZD8 CRD–Fc remained bound to the beads. All other proteins were expressed in High Five (Trichoplusia ni) cells (Invitrogen) using the baculovirus expression system. To produce the B12-based surrogate, the coding sequences of B12, a flexible linker peptide comprising of 0, 1, 2 or 3 GSGSG-linker repeats, followed by the C-terminal domain of human DKK1 (residues 177–266), and a C-terminal 6×His-tag, were cloned into the pAcGP67A vector (BD Biosciences). To clone the scFv-based surrogate ligand, the sequence of the Vantictumab was retrieved from the published patent, reformatted into a scFv, and cloned at the N terminus of the surrogate variant containing the GSGSG linker peptide. To produce recombinant FZD CRD for crystallization, surface plasmon resonance measurements, SEC-MALS experiments and functional assays, the CRDs of human FZD1 (residues 113–182), human FZD4 (residues 42–161), human FZD5 (residues 30–150), human FZD7 (residues 36–163), human FZD8 (residues 32–151) and human FZD10 (residues 30–150), containing a C-terminal 3C protease cleavage site (LEVLFQ/GP), a biotin acceptor peptide (BAP)-tag (GLNDIFEAQKIEWHE) and a 6×His-tag were cloned into the same vector. The human FZD8 CRD used for crystallization contained only a C-terminal 6×His-tag, in addition to a Asn49Gln mutation to mutate the N-linked glycosylation site. FZD1/FZD8 CRD for inhibition assay contained a C-terminal 3C protease cleavage site, Fc-tag (constant region of human IgG), and a 6×His-tag. Human DKK1 (residues 32–266) with a C-terminal BAP-tag and 6×His-tag, and the two furin-like repeats of human RSPO2 (residues 36–143) with a N-terminal Fc-tag and a C-terminal 6×His-tag, were cloned also into the pAcGP67A vector. All proteins were secreted from High Five insect cells grown in Insect-Xpress medium, and purified using Ni-NTA affinity purification, and size-exclusion chromatography equilibrated in HBS (10 mM HEPES, pH 7.3, 150 nM NaCl). Enzymatic biotinylation was performed in 50 mM bicine, pH 8.3, 10 mM ATP, 10 mM magnesium acetate, 0.5 mM d-biotin with recombinant glutathione S-transferase (GST)-tagged BirA ligase overnight at 4 °C, and proteins were subsequently re-purified on a Superdex 75 size-exclusion column to remove excess biotin. We attempted to mimic the native Wnt–FZD lipid–protein interaction with a de novo designed protein–protein binding interface. A 13-residue alanine helix was docked against the lipid-binding cleft using Foldit34. This structural element was grafted onto a diverse set of native helical proteins using the Rosetta Epigraft35 application to discover scaffolds with compatible, shape-complementary backbones. Prototype designs were selected by interface size and optimized using RosettaScripts36 to perform side-chain redesign. 50 selected designs were further manually designed to ensure charge complementarity and non-essential mutations were reverted to the wild-type amino acid identity to maximize stability. DNA was obtained from Gen9 and screened for binding via yeast surface display as previously described with 1 μM biotinylated FZD8 CRD pre-incubated with 025 µM SAPE (Life Technologies)37. A design based on the scaffold with PDB code 2QUP, a uncharacterized four-helix bundle protein from Bacillus halodurans, demonstrated binding activity under these conditions, whereas knockout mutants Ala52Arg and Ala53Asd made using the Kunkel method38 abrogated binding, verifying that the functional interface used the predicted residues. Wild-type scaffold 2QUP did not bind, confirming that activity was specifically due to design. To improve the affinity of the original design, a full-coverage site-saturation mutagenesis library was constructed for design based on the 2QUP scaffold via the Kunkel mutagenesis method38 using forward and reverse primers containing a ‘NNK’ degenerate codon and 21-bp flanking regions (IDT). A yeast library was transformed as previously described39 and sorted for three rounds, collecting the top 1% of binders using the BD Influx cell sorter. Naive and selected libraries were prepared and sequenced, and the data was processed as previously described37 using a Miseq (Illumina) according to manufacturer protocols. The most enriched 11 mutations were identified by comparison of the selected and unselected pools of binders and were combined in a degenerate library containing all enriched and wild-type amino acid identities at each of these positions. This combination library was assembled from the oligonucleotides (IDT) listed below for a final theoretical diversity of around 800 k distinct variants. This library was amplified, transformed, and selected to convergence over five rounds, yielding the optimized variant B12. The B12–FZD8 CRD(N49Q) complex was formed by mixing purified B12 and FZD8 CRD(N49Q) in stoichiometric quantities. The complex was then treated with 1:100 (w/w) carboxypeptidase A (Sigma) overnight at 4 °C, and purified on a Superdex 75 (GE Healthcare Life Sciences) size-exclusion column equilibrated in HBS. Purified complex was concentrated to around 15 mg ml−1 for crystallization trials. Crystals were grown by hanging-drop vapour diffusion at 295 K, by mixing equal volumes of the complex and reservoir solution containing 42–49% PEG 400, 0.1 M Tris, pH 7.8–8.2, 0.2 M NaCl, or 20% PEG 3000, 0.1 M sodium citrate, pH 5.5. While the PEG 400 condition is already a cryo-protectant, the crystals grown in the PEG 3000 condition were cryoprotected in reservoir solution supplemented with 20% glycerol before flash freezing in liquid nitrogen. Crystals grew in space groups P2 (PEG 400 condition) and P2 (PEG 3000 condition), respectively, with 2 and 4 complexes in the asymmetric units. Cell dimensions are listed in Supplementary Table 1. Data were collected at beamline 8.2.2 at the Advanced Light Source (ALS), Lawrence Berkeley National Laboratory. All data were indexed, integrated, and scaled with the XDS package40. The crystal structures in both space groups were solved by molecular replacement with the program PHASER41 using the structure of the FZD8 CRD (PDB code 1IJY) and the designed model of a minimal core of B12 as search models. Missing residues were manually build in COOT42 after initial rounds of refinement. Several residues at the N terminus (residues 1 to 16/17/20/21), at the C terminus (residues after 117) and several residues within loop regions were unstructured and could not been modelled. Furthermore, we observed that in both crystal forms, B12 underwent domain swapping, and one B12 molecule lent helix 3 and 2 to another B12, resulting in a closely packed B12 homodimer. The density of the loops connecting helixes 1 and 2, and 3 and 4 were clearly visible, and folded into helical turns. Yet, SEC-MALS experiments confirmed that B12 existed as a monomer in solution, and complexed FZD8 CRD with a 1:1 stoichiometry. PHENIX Refine43 was used to perform group coordinate refinement (rigid body refinement), followed by individual coordinate refinement using gradient-driven minimization applying stereo-chemical restraints, NCS restraints, and optimization of X-ray/stereochemistry weight, and individual B-factor refinement. Initial rounds of refinement were aided by restraints from the high-resolution mouse FZD8 CRD structure as a reference model. Real space refinement was performed in COOT into a likelihood-weighted SigmaA-weighted 2mF − DF map calculated in PHENIX. The final model in the P2 space group was refined to 3.20 Å with R and R values of 0.2002 and 0.2476, respectively (Supplementary Table 1). The quality of the structure was validated with MolProbity44. 99.5% of residues are in the favoured region of the Ramachandran plot, and no residue in the disallowed region. The structure within the P2 space group was refined to 2.99 Å with R and R values of 0.2253 and 0.2499, respectively, with 99.2% of residues in the favoured region of the Ramachandran plot, and no residues in the disallowed region. See Supplementary Table 1 for data and refinement statistics. Structure figures were prepared with the program PYMOL. Binding measurements were performed by surface plasmon resonance on a BIAcore T100 (GE Healthcare) and all proteins were purified on SEC before experiments. Biotinylated FZD1 CRD, FZD5 CRD, FZD7 CRD and FZD8 CRD were coupled at a low density to streptavidin on a SA sensor chip (GE Healthcare). An unrelated biotinylated protein was captured at equivalent coupling density to the control flow cells. Increasing concentrations of B12 and scFv–DKK1c were flown over the chip in HBS-P (GE Healthcare) containing 10% glycerol and 0.05% BSA at 40 μl ml−1. The chip surface was regenerated after each injection with 2 M MgCl in HBS-P or 50% ethylene glycol in HBS-P (scFv–DKK1c measurements), or 4 M MgCl in HBS-P (B12 measurements) for 60 s. Curves were reference-subtracted and all data were analysed using the Biacore T100 evaluation software version 2.0 with a 1:1 Langmuir binding model to determine the K values. To characterize the FZD-specificity of B12, the yeast display vector encoding B12 was transformed into EBY100 yeast. To induce the display of B12 on the yeast surface, cells were growing in SGCAA medium45, 46 for 2 days at 20 °C. 1 × 106 yeast cells per condition were washed with PBE (PBS, 0.5% BSA, 2 mM EDTA), and stained separately with 0.06–1,000 nM biotinylated FZD1/4/5/7/8/10 CRDs for 2 h at 4 °C. After washing twice with ice-cold PBE, bound FZD CRDs were labelled with 10 nM strepdavidin-Alexa647 for 20 min. Cells were fixed with 4% paraformaldehyde, and bound FZD CRD was analysing on an Accuri C6 flow cytometer. FZD8 fused to an N-terminal HaloTag47 and LRP6 fused to an N-terminal SNAP-tag48 were cloned into the pSEMS-26m vector (Covalys Biosciences) by cassette cloning49, 50. The template pSEMS-26m vectors had been coded with DNA sequences of the SNAP-tag or the HaloTag, respectively, together with an Igκ leader sequence (from the pDisplay vector, Invitrogen) as described previously50. The genes of full-length mouse Fzd8 or human LRP6 without the N-terminal signal sequences were inserted into pSEMS-26m via the XhoI and AscI or AscI and NotI, restriction sites, respectively. A plasmid encoding a model transmembrane protein, maltose-binding protein fused to a transmembrane domain, fused to an N-terminal HaloTag was prepared as described recently13. HeLa cells were cultivated at 37 °C, 5% CO in MEM Earle’s (Biochrom AG, FG0325) supplemented with 10% fetal calf serum and 1% nonessential amino acids. Cells were plated in 60-mm cell culture dishes to a density of 50% confluence and transfected via calcium phosphate precipitation49. 8–10 h after transfection, cells were washed twice with PBS and the medium was exchanged, supplied with 2 μM porcupine inhibitor IWP-2 for inhibiting maturation of endogenous Wnt in HeLa cells51. 24 h after transfection, cells were plated on glass coverslips pre-coated with PLL-PEG-RGD52 for reducing nonspecific binding of dyes during fluorescence labelling. After culturing for 12 h, coverslips were mounted into microscopy chambers for live-cell imaging. SNAP-tag and HaloTag were labelled by incubating cells with 50 nM benzylguanine-DY649 (SNAP-Surface 649, New England Biolabs) and 80 nM of HaloTag tetramethylrhodamine ligand (HTL-TMR, Promega) for 20 min at 37 °C. Under these conditions, effective degrees of labelling estimated from single molecule assays with a HaloTag–SNAP-tag fusion protein were ~40% for the SNAP-tag and ~25% for the HaloTag13. After washing three times with PBS, the chamber was refilled with MEM containing 2 μM IWP-2 for single-molecule fluorescence imaging. Single-molecule fluorescence imaging was carried out by using an inverted microscope (Olympus IX71) equipped with a triple-line total internal reflection (TIR) illumination condenser (Olympus) and a back-illuminated EMCCD camera (iXon DU897D, 512 × 512 pixel from Andor Technology). A 561-nm diode solid state laser (CL-561-200, CrystaLaser) and a 642-nm laser diode (Luxx 642-140, Omicron) were coupled into the microscope for excitation. Laser lights were reflected by a quad-line dichroic beam splitter (Di R405/488/561/647, Semrock) and passed through a TIRF objective (UAPO 150×/1.45, Olympus). For simultaneous dual-colour detection, a DualView microimager (Optical Insight) equipped with a 640 DCXR dichroic beamsplitter (Chroma) in combination with bandpass filters FF01-585/40 and FF01 670/30 (Semrock), respectively, was mounted in front of the camera. The overlay of the two channels was calibrated by imaging fluorescent microbeads (TetraSpeck microspheres 0.1 μm, T7279, Invitrogen), which were used for calculating a transformation matrix. After channel alignment, the deviation between the channels was below 10 nm. For single-molecule imaging, typical excitation powers of 1 mW at 561 nm and 0.7 mW at 642 nm measured at the objective were used. Time series of 150–300 frames were recorded at 30 Hz (4.8–9.6 s). An oxygen scavenging system containing 0.5 mg ml−1 glucose oxidase, 40 mg ml−1 catalase, and 5% (w/v) glucose, together with 1 μM ascorbic acid and 1 μM methyl viologene, was added to minimize photobleaching53. Receptor dimerization was initiated by incubating with 100 nM Wnt proteins or surrogates. Images were acquired after 5 min incubation in the presence of the ligands. All live-cell imaging experiments were carried out at room temperature. A 2D Gaussian mask was used for localizing single emitters54, 55. For colocalization analysis to determine the heterodimerization fraction, particle coordinates from two channels were aligned by a projective transformation (cp2tform of type ‘projective’, MATLAB 2012a) according to the transformation matrix obtained from microbead calibration measurement. Particles colocalized within a distance of 150 nm were selected. Only co-localized particles, which could be tracked for at least 10 consecutive frames (that is, molecules co-locomoting for at least 0.32 s) were accepted as receptor heterodimers or hetero-oligomers, which has been previously found to be a robust criterion for protein dimerization13. The fraction of heterodimerization or hetero-oligomerization was determined as the number of co-locomotion trajectories with respect to the number of the receptor trajectories. Since the receptor expression level of FZD8 or LRP6 was variable in the transiently transfected cells, only cells with similar receptor expression levels were considered (less than three times the excess of one subunit over the other). The smaller number of trajectories of either FZD8 or LRP6 was regarded as the limiting factor and therefore taken as a reference for calculating the heterodimerized/hetero-oligomerized fraction. Oligomerization values were not corrected for the degree of labelling. Single-molecule trajectories were reconstructed using the multi-target tracing (MTT) algorithm56. The detected trajectories were evaluated with respect to their step length distribution to determine the diffusion coefficients. For a reliable quantification of local mobilities, we estimated diffusion constants from the displacements with three frames (96 ms). Step-length histograms were obtained from all single molecule trajectories and fitted by a two-component model of Brownian diffusion, thus taking into account the intrinsic heterogeneity of protein diffusion in the plasma membrane57, 58. A bimodal probability density function p(r) was used for a nonlinear least square fit of the step-length histogram: where is the percentage of the fraction, contains the diffusion coefficient of each fraction (nδt = 96 ms). Average diffusion coefficients were determined by weighting the diffusion coefficients with the corresponding fractions. Single-molecule intensity distribution of individual diffraction-limited spots was extracted from the first 50 images of the recorded time lapse image sequence, in which photobleaching of dyes was kept below 10%13. Oligomerization of receptors was evaluated by fitting the obtained single molecule intensity with a multi-component Gaussian distribution function59. To ensure a reliable analysis, monomeric receptors were first distinguished based on the observation that monomers diffused much faster than oligomers. Therefore, the characteristic intensity distribution of monomeric receptor subunits was obtained by tracking of the fast mobile fraction. Fractions of the monomer, dimer, trimer and higher oligomers were then de-convoluted from the single molecule intensity distribution, presuming that intensities of clusters were multiples of the monomer intensity distribution. Immortal cells were seeded in triplicate for each condition in 96-well plates, and stimulated with surrogates, XWnt8, WNT3A conditioned media, control proteins, or other treatments for 20–24 h. After washing cells with PBS, cells in each well were lysed in 30 μl passive lysis buffer (Promega). 10 μl per well of lysate was assayed using the Dual Luciferase Assay kit (Promega) and normalized to the Renilla luciferase signal driven constitutively by the human elongation factor-1 alpha promoter to account for cell variability. A375 BAR, SH-SY5Y BAR, L STF and HEK293 STF cells were plated at a density of 10,000–20,000 cells per well, and treatment was started after 24 h in fresh medium. A549 BAR cells were plated at a density of 5,000 cells per well in the presence of 2 μM IWP-2 (Calbiochem) to suppress endogenous Wnt secretion, and treatment was started after 48 h in fresh medium containing fresh IWP-2. To induce β-catenin accumulation, SH-SY5Y BAR cells were treated for 2 h with scFv–DKK1c, WNT3A conditioned media (positive control), B12 (negative control protein) and mock conditioned media (from untransfected L cells, negative control) at 37 °C, 5% CO . After, cells were washed twice with PBS. For β-catenin stabilization assay, cells were scraped into hypotonic lysis buffer (10 mM Tris-HCl pH 7.4, 0.2 mM MgCl , supplemented with protease inhibitors), incubated on ice for 10 min, and homogenized using a hypodermic needle. Sucrose and EDTA were added to final concentration of 0.25 M and 1 mM, respectively. For LRP6 phosphorylation assay, cells were lysed in RIPA buffer (50 mM Tris pH 8.0, 150 mM NaCl, 0.5% sodium deoxylate, 1% Triton X-100), supplemented with protease inhibitor and phosphatase inhibitor for 1 h at 4 °C. Lysates were centrifuged at 12,000g for 1 h at 4 °C. Supernatants were then diluted into SDS sample buffer. For immunoblotting, samples were resolved on a 12% Mini-PROTEAN(R)TGX precast protein gel (Bio-Rad) and transferred to a PVDF membrane. The membranes were cut horizontally approx. at the 64 kDa mark of the SeeBlue plus 2 molecular mass marker (Invitrogen). Top half of the blot was incubated with anti-β-catenin primary antibody ((D10A8)XP, rabbit, Cell Signaling 8480), LRP6 antibody ((C47E12), rabbit, Cell Signaling 3395), and P-LRP6 (S1490) antibody (rabbit, Cell Signaling 2568), and the bottom part with the anti-α-tubulin primary antibody (mouse, DM1A, Sigma) in PBS containing 0.1% Tween-20 and 5% BSA overnight at 4 °C. Blots were then washed, incubated with the corresponding secondary antibodies in the same buffer, before washing and developing using the ECL prime western blotting detection reagent (GE Healthcare). To induce β-catenin accumulation, K562 and cells were stimulated for 0, 15, 30, 45, 60, 90 and 120 min with 10 nM scFv–DKK1c, recombinant Wnt3a (R&D Systems), B12 (negative control protein) or plain complete growth medium at 37 °C, 5% CO . After, cells were washed twice with PBS, fixed with 4% PFA for 10 min at room temperature, and permeabilized in 100% methanol for at least 30 min at −80 °C. The cells were than stained with Alexa-647 conjugated anti-β-catenin antibody (L54E2) (Cell Signaling Technology, 1:100–1: 50 dilution). Fluorescence was analysed on an Accuri C6 flow cytometer. Total RNA was isolated using either TRIZOL (Invitrogen) or RNeasy plus micro kit (QIAGEN) according to manufacturer’s protocols. A total of 2 μg RNA were used to generate cDNA using the RevertAid RT kit (Life Technologies) using oligo(dT)18 mRNA primers (Life Technologies) according to manufacturer’s protocol. 12 ng of cDNA per reaction were used. qPCR was performed using SYBR Green-based detection (Applied Biosystems) according to the manufacturer’s protocol on a StepOnePlus real-time PCR system (ThermoFisher Scientific). All primers were published, or validated by us. Transcript copy numbers were normalized to GAPDH for each sample, and fold induction compared to control was calculated. The following gene-specific validated primers were used: human FZD1: F: 5′-ATCCTGTGTGCTCCTCTTTTGG-3′, R: 5′-GATTGCTTTTCTCCTCTTCTTCAC-3′; human FZD2: F: 5′-CTGGGCGAGCGTGATTGT-3′, R: 5′-GTGGTGACAGTGAAGAAGGTGGAAG-3′; human FZD3: F: 5′-TCTGTATTTTGGGTTGGAAGCA-3′, R: 5′-CGGCTCTCATTCACTATCTCTTT-3′; human FZD4: F: 5′-TGGGCACTTTTTCGGTATTC-3′, R: 5′-TGCCCACCAACAAAGACATA-3′; human FZD5: F: 5′-CCATGATTCTTTAAGGTGAGCTG-3′, R: 5′-ACTTATTCAAGACACAACGATGG-3′; human FZD6: F: 5′-CGATAGCACAGCCTGCAATA-3′, R: 5′-ACGGTGCAAGCCTTATTTTG-3′; human FZD7: F: 5-TACCATAGTGAACGAAGAGGA-3′, R: 5′-TGTCAAAGGTGGGATAAAGG-3′; human FZD8: F: 5′-ACCCAGCCCCTTTTCCTCCATT-3′, R: 5′-GTCCACCCTCCTCAGCCAAC-3′; human FZD9: F: 5′-GCTGTGACTGGAATAAACCCC, R: 5′-GCTCTGCTTACAAGAAAGACTCC-3′; human FZD10: F: 5′-CTCTTCTCTGTGCTGTACACC, R: 5′-GTCTTGGAGGTCCAAATCCA-3′; mouse Fzd1: F: 5′-GCGACGTACTGAGCGGAGTG, R: 5′-TGATGGTGCGGATGCGGAAG-3′60; mouse Fzd2: F: 5′-CTCAAGGTGCCGTCCTATCTCAG, R: GCAGCACAACACCGACCATG-3′60; mouse Fzd3: F: 5′-GGTGTCCCGTGGCCTGAAG-3′, R: 5′-ACGTGCAGAAAGGAATAGCCAAG-3′60; mouse Fzd4: F: 5′-GACAACTTTCACGCCGCTCATC-3′, R: 5′-CAGGCAAACCCAAATTCTCTCAG-3′60; mouse Fzd5: F: 5′-AAGCTGCCTTCGGATGACTA-3′, R: 5′-TGCACAAGTTGCTGAACTCC-3′60; mouse Fzd6: F: 5′-TGTTGGTATCTCTGCGGTCTTCTG-3′, R: 5′-CTCGGCGGCTCTCACTGATG-3′60; mouse Fzd7: F: 5′-ATATCGCCTACAACCAGACCATCC-3′, R: 5′-AAGGAACGGCACGGAGGAATG-3′60; mouse Fzd8: F: 5′-GTTCAGTCATCAAGCAGCAAGGAG-3′, R: 5′-AAGGCAGGCGACAACGACG-3′60; mouse Fzd9: F: 5′-ATGAAGACGGGAGGCACCAATAC-3′, R: 5′-TAGCAGACAATGACGCAGGTGG-3′60; mouse Fzd10: F: 5′-ATCGGCACTTCCTTCATCCTGTC-3′, R: 5′-TCTTCCAGTAGTCCATGTTGAG-3′60; human AXIN2: F: 5′-CTCCCCACCTTGAATGAAGA-3′, R: 5′-TGGCTGGTGCAAAGACATAG-3′; human GAPDH: F: 5′-TGAAGGTCGGAGTCAACGGA-3′, R: 5′-CCATTGATGACAAGCTTCCCG-3′; mouse Gapdh: F: 5′-CCCCAATGTGTCCGTCGTG-3′, R: 5′-GCCTGCTTCACCACCTTCT-3′. Differentiation of C3H10T1/2, and human and mouse primary MSCs were performed essentially as described previously61. In brief, approximately 10,000 cells cm−2 were plated in normal culture medium (αMEM + FBS + penicillin/streptomycin), and allowed to adhere overnight. The following day, the medium was replaced with osteogenic medium (αMEM, 10% FBS, 1% penicillin/streptomycin, 50 μg ml−1 ascorbic acid, 10 mM β-glycerol phosphate (βGP), and replaced every other day. To determine alkaline phosphatase enzymatic activity, cells were fixed for 10 min with 10% formalin in PH7 PBS, before incubation in NBT-BCIP solution (1-Step(tm) NBT/BCIP Substrate Solution (Thermo Fisher Scientific, 34042) for 30 min. qPCR reactions were done with the SYBR method using the following primers: human ACTB F: 5′-GTTGTCGACGACGAGCG-3′, R: 5′-GCACAGAGCCTCGCCTT-3′; human ALPL: F: 5′-GATGTGGAGTATGAGAGTGACG-3′, R: 5′-GGTCAAGGGTCAGGAGTTC-3′; mouse Alpl: F: 5′-AAGGCTTCTTCTTGCTGGTG-3′, R: 5′-GCCTTACCCTCATGATGTCC-3′; mouse Actb: F: 5′-GGAATGGGTCAGAAGGACTC-3′, R: 5′-CATGTCGTCCCAGTTGGTAA-3′; mouse Col2a1 F: 5′-GTGGACGCTCAGGAGAAACA-3′, R: 5′-TGACATGTCGATGCCAGGAC-3′. P26N, normal adult human colon organoids, were established from a tumour-free colon segment of a patient diagnosed with CRC as described18, 62, 63. CFTR-derived colorectal organoids were obtained from a patient at Wilhelmina Children’s Hospital WKZ-UMCU. Informed consent for the generation and use of these organoids for experimentation was approved by the ethical committee at University Medical Center Utrecht (UMCU) (TcBio 14-008). Human stomach organoids, derived from normal corpus and pylorus, were from patients that underwent partial or total gastrectomy at the University Medical Centre Utrecht (UMCU) and were established as described19, 64, 65. Pancreas organoids were obtained from the healthy part of the pancreas of patients undergoing surgical resection of a tumour at the University Medical Centre Utrecht Hospital (UMC) and were established as described66, 67. The liver organoids were derived from freshly isolated normal liver tissue from a patient with metastatic CRC who presented at the UMC hospital (ethical approval code TCBio 14-007) and were established as described20, 68. For the performance of 3D cultures, Matrigel (BD Biosciences) was used and overlaid with a liquid medium consisting of DMEM/F12 advanced medium (Invitrogen), supplemented with additional factors as outlined below. 2% RSPO3-CM (produced via the r-PEX protein expression platform at U-Protein Express BV), WNT3A conditioned medium (50%, produced using stably transfected L cells in the presence of DMEM/F12 advanced medium supplemented with 10% FBS), and Wnt and Wnt/RSPO2 surrogates at different concentrations were added as indicated. Single-cell suspensions of normal human organoids were cultured in duplicate or triplicate in round-bottom 96-well plates to perform a cell viability test using Cell Titer-Glo 3D (Promega). In brief, organoids were trypsinized to single-cell suspension and plated in 100 μl medium in the presence of the different reagents. 3 μM IWP-2 was added to inhibit endogenous Wnt lipidation and secretion. After 12 days, 100 μl of Cell Titer-Glo 3D was added, plates were shaken for 5 min, incubated for an additional 25 min and centrifuged before luminescence measurement. All animal experiments were conducted in accordance with procedures approved by the IACUC at Stanford University. Experiments were not randomized, the investigators were not blinded, and all samples/data were included in the analysis. Group sample sizes were chosen based on (1) previous experiments, (2) performance of statistics analysis, and (3) logistical reasons with respect to full study size, to accommodate all groups. Adenoviruses (E1 and E3 deleted, replication deficient) were constructed to express scFv–DKK1c or scFv–DKK1c–RSPO2 with an N-terminal signal peptide and C-terminal 6×His-tag (Ad-scFv–DKK1c or Ad-scFv–DKK1c–RSPO2), respectively. Adenoviruses expressing mouse IgG2α Fc (Ad-Fc), human RSPO2–Fc fusion protein (Ad-RSPO2–Fc) and mouse WNT3A (Ad-Wnt3a) were constructed and described in the companion paper by Yan et al.26 The adenoviruses were cloned, purified by CsCl gradient, and titred as previously described69. Adult C57Bl/6J mice were purchased from Taconic Biosciences. Adult C57Bl/6J mice between 8–10 weeks old were injected intravenously with a single dose of adenovirus at between 1.2 × 107 p.f.u. to 6 × 108 p.f.u. per mouse in 0.1 ml PBS. Serum expression of Ad-scFv–DKK1c or Ad-scFv–DKK1c–RSPO2 were confirmed by immunoblotting using mouse anti-6×His (Abcam ab18184, 1:2,000) or rabbit anti-6×His (Abcam ab9108, 1:1,000), respectively. All experiments used n = 4 mice per group and repeated at least twice. qRT–PCR on liver samples were performed as following. Total cDNA was prepared from each liver sample using Direct-Zol RNA miniprep kit (Zymo Research) and iScript Reverse Transcription Supermix for RT-qPCR (BIO-RAD). Gene expression was analysed by -ΔΔC or fold change (2−ΔΔCt). Unpaired Student’s t-test (two tailed) was used to analyse statistical significance. Primers for mouse Axin2 and Cyp2f2 were previously published70. Additional primers used were listed as below: For the parabiosis experiment, age- and gender-matched C57Bl/6J mice were housed together for at least 2 weeks before surgery. At 2 days before surgery, the ‘donor’ mice were injected intravenously with a single dose of adenovirus at between 1.2 × 107 pfu to 6 × 108 pfu per mouse in 0.1 ml PBS and were separated from the ‘recipient’ mice until surgery. The parabiosis surgery was performed as described previously71. The establishment of shared circulation was confirmed at day 5 after surgery by presence of adenovirus-expressed proteins in the serum of both donors and recipients. Mouse livers were collected and fixed in 4% paraformaldehyde. 5 μm paraffin-embedded sections were stained with the following antibodies after citrate antigen retrieval and blocking with 10% normal goat serum: mouse anti-glutamine synthetase antibody (Millipore MAB302, 1:200), mouse anti-PCNA (BioLegend 307902, 1:200), and rabbit anti-HNF4α (Cell Signaling 3113S, 1:500). The immunostained tissue sections were analysed and images were captured on a Zeiss Axio-Imager Z1 with ApoTome attachment. Atomic structure factors and coordinates have been deposited to the Protein Data Bank (PDB) under accession numbers 5UN5 and 5UN6. All other data are available from the corresponding author upon reasonable request.
News Article | May 10, 2017
No statistical methods were used to predetermine sample size. The experiments were not randomized, and investigators were not blinded to allocation during experiments and outcome assessment. Recombinant adenoviruses were constructed with the following inserts. Full-length mouse Dkk1 (ref. 6), and mouse Rspo1-Fc16 with full-length Rspo1 fused to a mouse antibody IgG2α Fc fragment at the C terminus have been described. Human RSPO2 and mouse Rnf43 ECD and Znrf3 ECD similarly contained full-length open reading frames with a C-terminal mouse IgG2α Fc fragment. Mouse Fzd8 CRD (residues 25–173) was cloned with an N-terminal haemagglutinin (HA) epitope tag and C-terminal IgG2α Fc fragment. In addition, a recombinant adenovirus was engineered to express human LGR5 ECD with both C-terminal FLAG and histidine tags. The construction of the adenoviruses encoding the scFv–DKK1c Wnt surrogate agonist and scFv–DKK1c–RSPO2 single-chain polypeptide fusion, each with a C-terminal His tag is described in a companion paper by Janda et al.25 On day 2 after intravenous injection, scFv–DKK1c was found to be expressed in vivo at ~10–20 μg ml−1 (280–560 nM) in mouse sera and the serum potently induced TOPflash activity in vitro. Full-length Wnt3a cDNA (a gift from R. Nusse) was cloned without any epitope tags and detected by western blotting with anti-WNT3A (Cell Signaling 2391) against a recombinant WNT3A protein. No detectable WNT3A protein was found in mouse sera after intravenous injection. All adenoviral constructs contained an N-terminal signal peptide sequence to allow for their secretion. These adenoviruses were cloned by homologous recombination into E1− E3− adenovirus strain 5, purified by double CsCl gradient, and titred as previously described33. Recombinant proteins were expressed in serum-free CD293 medium (Invitrogen) of HEK293 cells infected by adenovirus. Recombinant LGR5-ECD protein was purified by nickel-NTA affinity chromatography (Qiagen) from Ad-LGR5-ECD-infected CD293 medium. Likewise, recombinant RNF43 and ZNRF3 ECD-Fc fusion proteins were purified by protein A affinity chromatography (KPL) from Ad-Rnf43-ECD-infected or Ad-Znrf3-ECD-infected CD293 medium, respectively. Protein purity was verified by Coomassie-stained SDS–PAGE. Adult Lgr5-eGFP-IRES-creER mice7 (Jax) or Axin2-LacZ mice (Jax) between 8 and 12 weeks old were injected intravenously with adenoviruses (doses of 5 × 108 to 1 × 109 pfu per mouse). Lgr5-eGFP-IRES-creER mice were crossed with Rosa26-tdTomato mice to generate Lgr5-eGFP-IRES-creER; Rosa26-tdTomato compound heterozygous mice. Similarly, Villin-creER or Actin-creER mice were crossed to Rosa26-Rainbow mice to generate Villin-creER; Rosa26-Rainbow or Actin-creER; Rosa26-Rainbow compound heterozygous mice. Mice were dosed with adenoviruses as above, and serum expression of all ECDs was confirmed by immunoblotting and histological assessment of intestinal crypt hyperplasia for those treated with Ad-Rspo1 and Ad-RSPO2. Adult mice between 8 and 12 weeks of age were administered tamoxifen (Sigma) dosed at 4 mg per 40 g body weight to genetically label for lineage tracing experiments using the various Rosa26 reporter strains. All in vivo experiments used n = 3–5 mice per group and were repeated at least twice except for the RNA-seq studies. Both male and female mice were used. All animal experiments were conducted in accordance with procedures approved by the IACUC at Stanford University. FACS experiments were performed using fresh small intestine epithelial preparations. A standardized 3 cm segment of proximal jejunum was used for quantitative FACS analysis of ISC populations. Intestinal epithelial cells were extracted from en bloc resected small intestine with 10 mM EDTA and manual shaking, followed by enzymatic dissociation with collagenase/dispase (Roche) to generate a single-cell suspension. Singlet discrimination was sequentially performed using plots for forward scatter (FSC-A versus FSC-H) and side scatter (SSC-W versus SSC-H). Dead cells were excluded by scatter characteristics and viability stains. All FACS experiments were performed on an Aria II sorter (BD) or LSRII analyser (BD) at the Stanford University Shared FACS Facility and FACS data were analysed using FlowJo software (TreeStar). Intestinal tissue was collected and fixed in 4% paraformaldehyde. 8-μm OCT frozen sections or 5-μm paraffin-embedded sections were TUNEL-stained using the DeadEnd Fluorometric TUNEL system per manufacturer’s instructions (Promega) or immunostained using the following primary antibodies: anti-Ki67 (ThermoFisher RM-9106), anti-MUC2 (Santa Cruz sc-15334), anti-lysozyme (Dako A0099), anti-chromogranin A (Santa Cruz sc-1488), anti-FABP1 (Novus NBP1-87695), anti-CD44 (BD Pharmingen 550538), anti-cyclin D1 (Abcam ab134175) and anti-CD166 (R&D AF1172). All primary antibodies were used at 1:100 to 1:200 dilutions. Cy3- and Cy5-conjugated secondary antibodies (Santa Cruz and Jackson ImmunoResearch) were used at 1:500 to 1:1,000 dilutions. Alexa Fluor 594-conjugated phalloidin (Invitrogen) was used at 1:500. CD166 immunostained tissue sections34 were analysed and confocal images acquired as 0.5-μm planes using an IX81 Inverted Microscope equipped with Fluoview FV1000-Spinning Disc Confocal scan head and FV10 ASW 1.7 software (Olympus). All other images were captured on a Zeiss Axio-Imager Z1 with ApoTome or Leica SP5 confocal microscope. In situ hybridization for Olfm4 mRNA was performed using the RNAscope kit (Advanced Cell Diagnostics) according to the manufacturer’s instructions. In brief, 5 μm formalin-fixed, paraffin-embedded tissue sections or 8 μm OCT frozen sections were pre-treated with heat and protease before hybridization with a target probe to Olfm4 mRNA. A horseradish peroxidase (HRP)-based signal amplification system was then hybridized to the target probes followed by colorimetric development with DAB. Negative control probes for the bacterial gene DapB were also included for each slide. Adult Lgr5-eGFP-IRES-creER mice (Jax) between 10 and 12 weeks old were treated with intravenous adenovirus. After 48 h, these mice were treated by oral gavage for 4 days with twice daily dosing interval with either 50 mg kg−1 of PORCN inhibitor C59 (Cellagen Technology) or vehicle consisting of 0.5% methylcellulose plus 0.1% Tween80, as previously described35. Mice were euthanized 20 h after the last dose of C59 and the intestine was harvested for FACS and histological analysis. Small intestine tissue samples were fixed with 2.5% glutaraldehyde and post-fixed in 1% osmium tetroxide in 100 mM phosphate buffer. Tissue was dehydrated, embedded in epoxy resin, and visualized by a JEOL transmission electron microscope at 120 kV (model JEM-1210). L cells stably transfected with TOPflash dual reporter plasmid system (a gift from J. Chen) were used in TOPflash dual luciferase assays (Promega Dual Luciferase kit) with WNT3A conditioned medium from a stably transfected WNT3A-expressing cell line (a gift from R. Nusse) from which activation of the TOPflash reporter has been confirmed; mycoplasma contamination was not tested. Recombinant WNT3A (R&D) was alternatively used. Recombinant mouse RSPO1–RSPO4 proteins (R&D) were used at 5 pM concentration each in these assays. Recombinant LGR5, RNF43 and ZNRF3 ECD proteins were expressed and purified as above and their purity and protein concentrations were determined by Coomassie-stained SDS–PAGE and Bradford assays. Assays were visualized with a Tecan M1000 luminometer. Recombinant scFv–DKK1c was expressed and purified as described in the companion paper25. The kinetics and affinity of interactions between RSPO1–RSPO4 and Flag- and histidine-tagged LGR5 ECD, Fc-tagged RNF43 ECD or Fc-tagged ZNRF3 ECD were determined by surface plasmon resonance. Data were collected on the BIAcore T100 instrument (GE Healthcare). Approximately 1,000 resonance units (RU) of recombinant mouse RSPO1, RSPO2, RSPO3 or RSPO4 (R&D) were immobilized on a CM5 sensor chip (GE Healthcare) using standard amine coupling. Increasing concentrations of LGR5 ECD, RNF43 ECD or ZNRF3 ECD were passed over the chip in HBS supplemented with 0.005% surfactant P20 (HBS+P). Binding phases for the LGR5-ECD were performed at 50 μl min−1 for 240 s and dissociation phases were performed at 50 μl min−1 for 1,850 s. The chip was regenerated after each injection with 240-s washes with 0.5 M magnesium chloride. Binding and dissociation phases for RNF43 ECD and ZNRF3 ECD were each performed at 50 μl min−1 for 120 s. The chip was regenerated after each injection with 120-s washes with 1 M magnesium chloride. All curves were reference-subtracted from a flow cell containing 1,000 RU of a negative control protein (hen egg white lysozyme or BSA). Curves were fitted using the BIAcore T100 evaluation software to a 1:1 model to determine the association rate (k ), dissociation rate (k ) and dissociation constant (K ). The kinetics and affinity of anti-RSPO antibody interactions with RSPO1–RSPO4 were determined as described for RNF43 and ZNRF3, except that the regeneration buffer was 25% ethylene glycol and 2.25 M magnesium chloride. The kinetics and affinity of Fc-tagged RNF43 and ZNRF3 ECDs are enhanced by avidity effects due to Fc-dimerization. The furin 1 and 2 repeats of human RSPO2 were cloned into the pCT302 vector as a C-terminal fusion to a c-Myc epitope and the cell-wall protein AGA2. RSPO2 was displayed on the EBY100 strain of Saccharomyces cerevisiae as previously described36. Competent yeast cells were electroporated with the RSPO2 expression plasmid and recovered in SDCAA selection media. The cultures were harvested in log phase, and yeast cells were then pelleted and resuspended in SGCAA induction media. Surface expression of RSPO2 was detected by staining yeast with a 488-labelled antibody to the c-Myc epitope (Cell Signaling 279), and then analysed by flow cytometry. Binding of LGR5, RNF43 and ZNRF3 ECDs was tested by incubating yeast with 200 nM recombinant Flag-tagged LGR5 ECD or with Fc-tagged RNF43 ECD or ZNRF3 ECD in PBS and 0.1% BSA for 2 h, washing twice with PBS and 0.1% BSA and then incubating for 30 min with an Alexa Fluor 647-labelled antibody to the Flag epitope (Cell Signaling 3916S) (for LGR5 binding) or a PE-labelled anti-IgG antibody (eBioscience 12-4998-82). Cells were washed twice with PBS and 0.1% BSA and then analysed by flow cytometry. Sequential staining of yeast was performed by incubating samples with 200 nM LGR5-ECD, 200 nM RNF43 ECD, or 200 nM ZNRF3 ECD alone, washing and then incubating with a mixture of (200 nM LGR5-ECD and 200 nM RNF43-ECD) or (200 nM LGR5-ECD and 200 nM ZNRF3 ECD). Cells double-stained with both LGR5-ECD and either RNF43 or ZNRF3 ECD were then washed and incubated with a mixture of PE-anti-IgG and 647-anti-Flag before a final wash and analysis by flow cytometry. Cells were isolated by flow cytometry into RNEasy lysis buffer (Qiagen) from n = 2–3 mice per condition, 1.5 days after injection of the appropriate adenoviruses. A 1.8× volume of AMPure beads (Beckman Coulter) was added to the thawed cell lysates. After a 30-min incubation at room temperature, the samples were washed twice with 70% ethanol and eluted in 22 μl water. The samples were then digested with 0.6 mAU Proteinase K (Qiagen) in the presence of 1× NEB buffer 1 (NEB) at 50 °C for 20 min, followed by a heat-inactivation step at 65 °C for 10 min. A DNase digestion was performed using the RNase-Free DNase Set (Qiagen) at 37 °C for 30 min. The samples were cleaned with a 1.8× volume of AMPure XP beads (Beckman Coulter). 1 ng of purified total RNA, as determined by Agilent Bioanalyzer (Agilent Technologies), was processed with the mRNA direct micro kit (Life Technologies) to select for poly A RNA. Each entire sample was input into the Ambion WT Expression Kit (Life Technologies) to perform double-stranded cDNA synthesis followed by in vitro transcription to generate amplified cRNA. The cRNA was purified following the manufacturer’s instructions and the concentration was determined with a NanoDrop instrument (ThermoFisher). 1 μg of cRNA was fragmented in 1× fragmentation buffer (mRNA-Seq Sample Prep Kit, Illumina) at 94 °C for 5 min, then placed on ice and the reaction was stopped by the addition of 20 mM EDTA. The fragments were precipitated with 70 mM sodium acetate (Life Technologies), 40 μg glycogen (Life Technologies) and 70% ethanol at −80 °C for 1 h followed by centrifugation and washing with 70% ethanol. 3 μg of random hexamer (Life Technologies) was added to the fragmented, purified cRNA and incubated at 70 °C for 10 min to anneal the primer. The first strand reaction was performed with 200 units of SuperScript II (Life Technologies) with 0.625 mM dNTPs (NEB) and 8U SUPERase RNase Inhibitor (Life Technologies) at 25 °C for 10 min, then 42 °C for 50 min, then 75 °C for 15 min and cooled to 4 °C. In second-strand synthesis, 1× second strand buffer (Illumina) and 0.3 mM dNTPs (Illumina) were added and the samples were incubated at 4 °C for 5 min before adding 50 U of DNA Polymerase (NEB) and 5 U of Rnase H (NEB). The samples were mixed well and incubated at 16 °C for 2.5 h, followed by purification with the MinElute Kit (Qiagen). To perform library prep, the samples were end repaired using a Quick Blunting Kit (NEB) and incubated at 20 °C for 1 h, then 75 °C for 30 min to inactivate the enzyme. To produce overhangs aimed to improve subsequent ligation efficiency, a single A base was added to the 3′ ends of each fragment with 2 mM dATP and 5 units of Klenow fragment 3′-5′ exo- DNA Polymerase (NEB) at 37 °C for 45 min, followed by 75 °C for 30 min to inactivate the enzyme. Using a quick ligase kit (NEB), 0.5 μM of adaptors containing single T base overhangs were ligated to the cDNA fragments at 12 °C for 75 min, then 80 °C for 20 min and cooled to 4 °C. These adaptors contain barcodes to facilitate sample multiplexing during sequencing. The adaptor sequence is preceded by four random nucleotides to add diversity to the pooled library. The samples were pooled by combining 5 μl of each library. After AMPure XP cleanup, one-half of the pooled library was run on the Pippin Size Selection Instrument (Sage Sciences) to select for 200 bp fragments. Library amplification was performed on one-half of the Pippin eluate in 1× Phusion GC buffer with 0.2 mM dNTPs, 0.1 μM forward primer (IDT), 0.1 μM reverse primer, 1 U Phusion Hot Start II Polymerase (Thermo Fisher Scientific). The reaction was run with the following program: 98 °C for 30 s, then 15 cycles of 98 °C for 10 s, 65 °C for 30 s, 72 °C for 30 s, then 72 °C for 4 min and cooled to 4 °C. The amplified library was cleaned using a 1× volume of AMPure XP beads and QC was run with the Agilent Bioanalyzer DNA 1000 kit, followed by concentration determination by qPCR using the KAPA Library Quantification Kit (KAPA Biosystems). To perform sequencing, the library was diluted to 4 nM and denatured with 0.1 N NaOH. Following denaturation, the library was further diluted to 4 pM and run on the Illumina HiSeq 2500 in paired-end, 100 × 100 bp format. Sequenced reads were aligned to the mouse reference genome mm9 (UCSC) using TopHat37 with the transcript annotation supplied. The mapped reads was assigned to gene using the tool htseq-count of the Python package HTseq38, with the default union-counting mode. The output of htseq-count was used as input for DESeq2 (ref. 39) to perform differential expression analysis, with a false discovery rate (FDR) of 10% as the cutoff. In addition, a filtering criterion of mean fragments per kilobase of transcript per million mapped reads (FPKM) of 1 in at least one condition was used to define expressed transcripts in each differential expression analysis. Cufflinks40 was used to calculate gene count and perform FPKM normalization. Gene Ontology term analysis was performed using DAVID functional annotation tool41. A FDR of 10% was applied to evaluate the significance. Lgr5-eGFP-IRES-creER mice were treated with adenovirus in vivo, and then 26 h after treatment the proximal jejunum was harvested to generate a single-cell suspension and FACS isolated using the endogenous GFP signal, as above. The sorted cellular suspensions were loaded on a GemCode Single Cell Instrument (10x Genomics) to generate single-cell gel beads in emulsion (GEMs). Approximately 1,200–2,800 cells were loaded per channel. Two technical replicates were generated per sorted cell suspension. Single-cell RNA-seq libraries were prepared using GemCode Single Cell 3′ Gel Bead and Library Kit (now sold as P/N 120230, 120231, 120232, 10x Genomics) as described previously29. Sequencing libraries were loaded at 2.1 pM on an Illumina Next-Seq500 with 2 × 75 paired-end kits using the following read length: 98 bp read1, 14 bp I7 index, 8 bp I5 index and 5 bp read2. Note that these libraries were generated before the official launch of GemCode Single Cell 3′ Gel Bead and Library Kit. Thus, 5 bp UMI was used (the official GemCode Single Cell 3′ Gel Bead contains 10 bp UMI). The Cell Ranger Single Cell Software Suite was used to perform sample de-multiplexing, barcode processing, and single-cell 3′ gene counting (http://software.10xgenomics.com/single-cell/overview/welcome). 5 bp UMI tags were extracted from read2. We analysed a total of 13,247 single cells, consisting of 11,268 FACS-sorted Lgr5–eGFP+and 1,979 Ad-Fc-treated Lgr5–eGFP− cells. Two technical replicates (the number of cells recovered per channel ranges from around 400 to 1,400 cells) were generated from each treatment condition. The mean raw reads per cell varied from ~45 k to 86 k. Each sample was downsampled to 28,439 confidently mapped reads per cell. Then the gene-cell barcode matrix from each sample was concatenated. The gene-cell barcode matrix was filtered based on number of genes detected per cell (any cells with less than 400 or more than 4,400 genes per cell were filtered) and percentage of mitochondrial UMI counts (any cells with more than 10% of mitochondrial UMI counts were filtered). Altogether, 13,176 cells, and 15,865 genes were kept for analysis by the Seurat R package30. Among these 13,176 cells, 74 did not show any epithelial cell markers so they were removed leaving a final total of 13,102 cells, consisting of 1,925 Ad-Fc-treated Lgr5–eGFP− cells and 11,177 Lgr5–eGFP+ cells across six conditions. 2,289 variable genes were selected based on their expression and dispersion (expression cutoff = 0.0125, and dispersion cutoff = 0.5). The first 11 principal components were used for the t-SNE projection and clustering analysis (resolution = 0.3, k.seed = 100). We applied sSeq from ref. 42 to identify genes that are enriched in a specific cluster (the specific cluster is assigned as group a, and the rest of clusters is assigned as group b). There are a few differences between our implementation and ref. 42. First, we used the ratio of total UMI counts and median of total UMI counts across all cells as the size factors. Second, the quantile rule of thumb was used to estimate the shrinkage target. Third, for genes with large counts, an asymptotic approximation from the edgeR package43 was used instead of the negative binomial exact test to speed up the computation. For the heatmap in Extended Data Fig. 9h, the gene list was furthered filtered requiring minimum UMI counts of 5 in each group, with a positive log fold change of mean expression between the two groups, and an adjusted P < 0.01. The top 10 genes specific to each cluster were picked, and their mean expression was centre scaled before used for the heatmap. Classification of cells was inferred from the annotation of cluster-specific genes. The stem cell clusters (clusters 0 and 1) were marked by enrichment of Lgr5, Olfm4 and Ascl2. Non-cycling and cycling stem cells were distinguished by the enrichment of cell cycle markers such as Mki67 and Tuba1b. Transit amplifying cells (cluster 2) were classified based on the enrichment of cell cycle markers and lack of Lgr5+ stem-cell marker expression. Enterocytes (clusters 3 and 4) were annotated based on the enrichment of markers such as Alpi and Reg1 and prior studies31. Goblet cells (cluster 5) were annotated based on the enrichment of markers such as Muc2 and Guca2a. Paneth cells (cluster 6) were annotated based on the enrichment of Defa genes. Tuft cells (cluster 7) were annotated based on the enrichment of markers such as Dclk1. EE cells (cluster 8) were annotated based on the enrichment of markers such as Chga and Chgb. To compare the global expression difference between samples and the Fc control, we first normalized gene expression by the sum of their UMI counts across all cells in the sample (adding 1 to the numerator and denominator to avoid dividing by 0 for genes that were not detected at all). Then we compared the normalized gene expression between the samples and the Fc control. To generate the heatmap, we furthered filtered the gene list: (1) Only genes with UMI counts >2 in each sample and a log fold change of >1 were considered. (2) The top 15 up- or downregulated genes were picked per sample–Fc comparison, and the union of all genes was used for the heatmap. Data generated during this study are available in the Gene Expression Omnibus (GEO) repository under accession numbers GSE92377 and GSE92865. All other data are available from the corresponding author upon reasonable request.