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News Article | May 9, 2017
Site: globenewswire.com

SAN DIEGO, May 09, 2017 (GLOBE NEWSWIRE) -- Royale Energy, Inc. (OTCQB:ROYL) announced it has signed a drilling contract for three additional wells in Rio Vista Field in Sacramento Basin and expects to begin drilling operations by the end of the month. The CRC 8-3, the CRC 8-4 and the CRC 8-5 wells will be drilled under Royale’s joint venture agreement covering the Rio Vista Field. Using 3-D seismic, Royale and its partner have identified several prospective locations in the Martinez, McCormick and Capay formations in the field. In 2016, Royale successfully completed three wells, two of the wells in the Martinez formation in Rio Vista which are presently producing over 1,500 MCF per day. Rio Vista Field is the largest natural gas field in California. Since 1936, the field has produced over 3.5 TCF of natural gas. Don Hosmer, President of Royale Energy said, "We are excited to begin our 2017 drilling campaign in the Rio Vista field. With natural gas prices continuing to stabilize and strengthen, we believe our Rio Vista development program will provide an excellent return for our shareholders and drilling investors." About Royale Energy, Inc. Founded in 1986, Royale Energy, Inc. (OTCQB:ROYL) is an independent exploration and production company focused on the acquisition, development, and marketing of natural gas and oil. Royale Energy has its primary operations in the Sacramento and San Joaquin basins in California. In addition to historical information contained herein, this news release contains “forward-looking statements” within the meaning of the Private Securities Litigation Reform Act of 1995, subject to various risks and uncertainties that could cause the company’s actual results to differ materially from those in the “forward-looking” statements. While the company believes its forward looking statements are based upon reasonable assumptions, there are factors that are difficult to predict and that are influenced by economic and other conditions beyond the company’s control. Investors are directed to consider such risks and other uncertainties discussed in documents filed by the company with the Securities and Exchange Commission.


News Article | May 9, 2017
Site: globenewswire.com

SAN DIEGO, May 09, 2017 (GLOBE NEWSWIRE) -- Royale Energy, Inc. (OTCQB:ROYL) announced it has signed a drilling contract for three additional wells in Rio Vista Field in Sacramento Basin and expects to begin drilling operations by the end of the month. The CRC 8-3, the CRC 8-4 and the CRC 8-5 wells will be drilled under Royale’s joint venture agreement covering the Rio Vista Field. Using 3-D seismic, Royale and its partner have identified several prospective locations in the Martinez, McCormick and Capay formations in the field. In 2016, Royale successfully completed three wells, two of the wells in the Martinez formation in Rio Vista which are presently producing over 1,500 MCF per day. Rio Vista Field is the largest natural gas field in California. Since 1936, the field has produced over 3.5 TCF of natural gas. Don Hosmer, President of Royale Energy said, "We are excited to begin our 2017 drilling campaign in the Rio Vista field. With natural gas prices continuing to stabilize and strengthen, we believe our Rio Vista development program will provide an excellent return for our shareholders and drilling investors." About Royale Energy, Inc. Founded in 1986, Royale Energy, Inc. (OTCQB:ROYL) is an independent exploration and production company focused on the acquisition, development, and marketing of natural gas and oil. Royale Energy has its primary operations in the Sacramento and San Joaquin basins in California. In addition to historical information contained herein, this news release contains “forward-looking statements” within the meaning of the Private Securities Litigation Reform Act of 1995, subject to various risks and uncertainties that could cause the company’s actual results to differ materially from those in the “forward-looking” statements. While the company believes its forward looking statements are based upon reasonable assumptions, there are factors that are difficult to predict and that are influenced by economic and other conditions beyond the company’s control. Investors are directed to consider such risks and other uncertainties discussed in documents filed by the company with the Securities and Exchange Commission.


On Wednesday, shares in Houston, Texas headquartered EP Energy Corp. ended the session 2.29% higher at $4.47 with a total volume of 776,557 shares traded. The stock is trading 2.33% below its 50-day moving average and 5.69% below its 200-day moving average. Moreover, shares of the Company, which engages in the exploration for and the acquisition, development, and production of oil, natural gas, and natural gas liquids in the US, have a Relative Strength Index (RSI) of 46.30. On May 09th, 2017, research firm Citigroup upgraded the Company's stock rating from 'Sell' to 'Neutral'. Sign up and read the free research report on EPE at: On Wednesday, shares in Irving, Texas headquartered Pioneer Natural Resources Co. recorded a trading volume of 1.54 million shares. The stock ended the day 1.06% higher at $171.27. Pioneer Natural Resources' stock has advanced 6.23% in the past one year. The Company's shares are trading below its 50-day and 200-day moving averages by 5.94% and 5.56%, respectively. Furthermore, shares of Pioneer Natural Resources, which operates as an independent oil and gas exploration and production company in the US, have an RSI of 41.89. On April 13th, 2017, research firm Stifel resumed its 'Buy' rating on the Company's stock, with a target price of $267 per share. The complimentary research report on PXD can be downloaded at: Calgary, Canada headquartered Gran Tierra Energy Inc.'s stock finished Wednesday's session 2.01% higher at $2.54 with a total volume of 1.15 million shares traded. Over the last one month and the previous one year, Gran Tierra Energy's shares have advanced 2.83% in the past three months. The Company's shares are trading below its 50-day and 200-day moving averages by 0.87% and 8.76%, respectively. Shares of Gran Tierra Energy, which engages in the acquisition, exploration, development, and production of oil and gas properties in Colombia, Peru, and Brazil, has an RSI of 50.47. Register for free on Stock-Callers.com and access the latest report on GTE at: Los Angeles headquartered California Resources Corp.'s stock advanced 5.43%, to close the day at $13.79. The stock recorded a trading volume of 1.97 million shares, which was above its three months average volume of 1.94 million shares. The Company's shares are trading 2.40% and 6.03% below its 50-day and 200-day moving averages, respectively. Shares of the Company, which operates as an oil and natural gas exploration and production company in the State of California, are trading at a PE ratio of 2.13. Additionally, the stock has an RSI of 55.58. 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News Article | May 8, 2017
Site: globenewswire.com

FREMONT, Calif., May 08, 2017 (GLOBE NEWSWIRE) -- Electronics For Imaging, Inc. (Nasdaq:EFII) today announced that it has acquired privately held CRC Information Systems (“CRC”), a Reynolds and Reynolds company. Reynolds and Reynolds is headquartered in Dayton, Ohio. CRC, which is based in Scottsdale, Ariz., is a provider of business management information systems (MIS) for commercial printers and packaging label and forms printers throughout the United States and Canada. While financial terms of the acquisition were not disclosed, it is not expected to be material to EFI's Q2 or full year 2017 results. “We are excited to welcome CRC customers and employees into the EFI family,” said Jeff White, general manager, SMB Segment, in the EFI Productivity Software business unit. “We intend to continue to meet the needs of CRC’s existing client base with the same enthusiasm they have come to expect over their years in business, while offering those customers access to the full EFI portfolio.” "We are very excited to join the EFI family and are confident in the additional value they can bring to our customers," said Erv Ratazak, manager, Product and Services, CRC Information Systems. About EFI EFI™ is a global technology company, based in Silicon Valley, and is leading the worldwide transformation from analog to digital imaging. We are passionate about fueling customer success with products that increase competitiveness and boost productivity. To do that, we develop breakthrough technologies for the manufacturing of signage, packaging, textiles, ceramic tiles, and personalized documents, with a wide range of printers, inks, digital front ends, and a comprehensive business and production workflow suite that transforms and streamlines the entire production process. (www.efi.com) Follow us on Twitter: https://twitter.com/EFIPrint Follow us on Instagram: https://www.instagram.com/efiprint Find us on Facebook: www.facebook.com/EFIPrint View us on YouTube: www.youtube.com/EFIDigitalPrintTech The Electronics For Imaging, Inc. logo is available at http://www.globenewswire.com/newsroom/prs/?pkgid=15847. Safe Harbor for Forward Looking Statements Certain statements in this press release are forward-looking statements within the meaning of Section 27A of the Securities Act of 1933, as amended and Section 21E of the Securities Exchange Act of 1934, as amended. Statements in this press release that could be deemed forward-looking statements include, but are not necessarily limited to, statements regarding the impact of the acquisition on EFI's results, expansion of our product portfolio, our future product offerings to CRC customers, integration of CRC, future customer achievements, continuation of support to the existing CRC client base, the timing of payments to the former CRC shareholders, and any statements or assumptions underlying any of the foregoing. Forward-looking statements are subject to certain risks and uncertainties that could cause our actual future results to differ materially, or cause a material adverse impact on our results. Potential risks and uncertainties include, but are not necessarily limited to, unforeseen expenses; the difficulty of aligning expense levels with revenue; management's ability to forecast revenues, expenses and earnings; any world-wide financial and economic difficulties and downturns; adverse tax-related matters such as tax audits, changes in our effective tax rate or new tax legislative proposals; the unpredictability of development schedules and commercialization of products by the leading printer manufacturers and declines or delays in demand for our related products; changes in the mix of products sold; the uncertainty of market acceptance of new product introductions; intense competition in each of our businesses, including competition from products developed by EFI’s customers; challenge of managing asset levels, including inventory and variations in inventory levels; the uncertainty of continued success in technological advances; the challenges of obtaining timely, efficient and quality product manufacturing and supply of components; litigation involving intellectual property rights or other related matters; our ability to successfully integrate acquired businesses; the uncertainty regarding the amount and timing of future share repurchases by EFI and the origin of funds used for such repurchases; the market prices of EFI’s common stock prior to, during and after the share repurchases; any disruptions in our operations, the difficulty to retain employees; the compliance with the new requirements regarding the "conflict minerals," if they are found to be used in our products, and any other risk factors that may be included from time to time in the Company's SEC reports. The statements in this press release are made as of the date of this press release. EFI undertakes no obligation to update information contained in this press release. For further information regarding risks and uncertainties associated with EFI's businesses, please refer to the sections entitled "Risk Factors," "Factors That Could Adversely Affect Performance," and other similar sections in our SEC filings and reports, including, but not limited to, EFI's annual report on Form 10-K and its quarterly reports on Form 10-Q, copies of which may be obtained by contacting EFI's Investor Relations Department by phone at 650-357-3828 or by email at investor.relations@efi.com or EFI's Investor Relations website at http://www.efi.com. NOTE TO EDITORS:  The EFI logo is a registered trademark of Electronics For Imaging, Inc. in the U.S. and/or certain other countries. EFI is a trademark of Electronics For Imaging, Inc. in the U.S. and/or certain other countries. All other terms and product names may be trademarks or registered trademarks of their respective owners, and are hereby acknowledged. Nothing herein should be construed as a warranty in addition to the express warranty statements provided with EFI products and services.


News Article | May 10, 2017
Site: www.biosciencetechnology.com

All bacteria have developed clever mechanisms for survival and propagation within host cells. Salmonella are a typical example: usually they hide in membrane-bound particles with only very few bacteria escaping to the cell's interior. Those escapees are extremely dangerous as they proliferate and spread at enormous speed. To stop such an invasion, cells have developed very effective defense strategies. An interdisciplinary team around Prof. Ivan Dikic (Institute of Biochemistry II) and Prof. Mike Heilemann (Institute of Physical and Theoretical Chemistry), both from Goethe University Frankfurt, now studied such a cellular defense mechanism by visualizing protein patterns at the near-molecular level. Upon bacterial invasion, cells react fast: They flag escaped bacteria with a small protein called ubiquitin, which is known to regulate numerous cellular processes. The attached flags contain chains of differently linked ubiquitin molecules, resulting in a secret code, which has so far only partially been decoded. Similar to mobile transmission towers, these ubiquitin chains relay specific signals from the surface of the bacteria into the cell. Employing super-resolution microscopy, the Frankfurt team now succeeded with visualizing different ubiquitin chains on the bacterial surface and analyzing their molecular organization in detail. They discovered that one chain type, so called linear chains, plays an essential role during a bacterial invasion. Linear ubiquitin chains trigger degradation of bacteria and kick off an inflammatory signaling cascade which results in restricting bacterial proliferation. In addition, the researchers identified the enzyme Otulin as an important regulator capable of limiting this reaction - a very important notion considering the fact that excessive inflammation is one of the major causes of tissue damage following bacterial infection. Signaling the cells' need for pathogen defense is just one important role of ubiquitin. The small protein is also involved in development and progression of inflammatory and neurodegenerative diseases as well as of cancer. Until now, however, very little is known about how small errors in the ubiquitin system contribute to these serious human diseases, and how the system can be targeted pharmaceutically. These new findings pave the way for many follow-up projects which may ultimately lead to novel therapeutic approaches. Very recently, Ivan Dikic obtained one of the prestigious ERC Advanced Grants of 2.5 M € in which he will investigate the role of ubiquitin in modulating the host-pathogen interaction in more detail. The work of the Frankfurt team is an excellent example for interdisciplinary collaboration and was enabled by funding of several large research networks, e.g. the Cluster of Excellence Macromolecular Complexes, the CRC 1177 on selective autophagy and the LOEWE ubiquitin network. The results are now published in the latest online issue of Nature Microbiology, back-to-back with complementary insights generated by colleagues in Cambridge (UK).


News Article | May 10, 2017
Site: www.prweb.com

Covenant Retirement Communities (CRC), a non-profit faith-based senior living organization headquartered in Skokie, announces the addition of Jody Holt as Chief Financial Officer (CFO) and Janine Wilson as Chief Operating Officer (COO). Jody Holt, a certified public accountant, brings more than 28 years of experience in both the public and private sectors and will work closely with CRC leadership on major strategic and operational issues while providing oversight on all financial and planning functions. Holt comes to CRC after serving as CFO for a leading global provider of high speed internet. In addition, she has served on the CRC board of directors. She holds a BS in accounting from The Pennsylvania State University. Janine Wilson brings more than 30 years of experience in operations, change management and resource development. As COO, she will oversee campus operations on CRC’s 12 senior living communities across the country, along with Lean Six Sigma, risk management and purchasing. She is a Master Black Belt in Six Sigma, a certified public accountant and certified project manager. Wilson has an MBA in strategic planning and a BS in business and engineering from the University of Toledo. “With the addition of Jody and Janine to the leadership team at CRC, we are poised to continue providing our 5,000 residents and 3,000 team members with solid operational excellence,” said Terri Cunliffe, CRC president. “Their breadth and depth of knowledge helps to round out our team as we provide day-to-day operations and strategically plan for the future with fiscal responsibility as a key focus.” ABOUT COVENANT RETIREMENT COMMUNITIES Chicago-based Covenant Retirement Communities (CRC) is the nation’s sixth largest, non-profit, senior services provider. It serves 5,000 residents at 15 retirement communities in 10 states and offers independent living, assisted living, skilled nursing, memory care, and rehabilitation. Through CRC’s LifeConnect® Wellness Partnership, residents can access resources and opportunities that are designed to enrich the mind, body, and spirit and complement each person’s unique journey. CRC is a ministry of the Evangelical Covenant Church and has been serving seniors since 1886. For more information, visit http://www.covenantretirement.org, Facebook, LinkedIn, and Twitter.


News Article | May 10, 2017
Site: www.nature.com

To generate CRISPR–Cas9 plasmids targeting the last exon of LGR5 (exon 18) or KRT20 (exon 8), 20-bp target sequences were cloned into a pX330-U6-Chimeric_BB-CBh-hSpCas9 plasmid (Addgene 42230) to obtain single vectors bicistronically expressing sgRNA and human codon-optimized Cas9 nuclease as previously described36. The sgRNA sequences targeting LGR5 or KRT20 are available in Supplementary Table 1. To construct donor vectors for LGR5–GFP- and KRT20–GFP-knock-in, 5′ and 3′ homology arms (1 kbp each) were amplified by PCR and cloned into an Ires-GFP-loxp-pEF1α-RFP-T2A-puro-loxp plasmid (HR180PA-1, SBI) using the In-Fusion HD Cloning kit (Clontech). For CreER or iCaspase9-T2A-tdTomato knock-in, we replaced GFP of the LGR5–GFP or KRT20–GFP construct with CreER or iCaspase9-T2A-tdTomato, respectivley. The final plasmid sequences were verified by DNA sequencing. To obtain a rainbow reporter, the rainbow cassette was excised from a CMV-Brainbow-2.1R plasmid (Addgene 18723) and cloned into a PiggyBac vector (PB510B-1, SBI). For bioluminescent imaging, we cloned optimized firefly luciferase luc2 into a GFP-expressing PiggyBac vector (PB513B-1, SBI). Knock-in efficiency and diver mutation profiles for each CCO line are available in Supplementary Table 2. All organoids were established as previously reported16 from patients who had given informed consent under the ethical committee of Keio University School of Medicine. The organoids were embedded in Matrigel and cultured with previously described basal culture medium37, specifically Advanced Dulbecco’s modified Eagle’s medium/F12 supplemented with penicillin/streptomycin, 10 mM HEPES, 2 mM GlutaMAX, 1× B27 (Life Technologies), 10 nM gastrin I (Sigma) and 1 mM N-acetylcysteine (Sigma). The following niche factors were added to the basal culture medium depending on the niche requirements of CRC organoid lines: 50 ng ml−1 mouse recombinant EGF, 100 ng ml−1 mouse recombinant noggin (PeproTech) and 500 nM A83-01 (Tocris). We electroporated the vectors under previously reported conditions37. Three days after electroporation, the organoids were selected with puromycin (2 μg ml−1) treatment for two days. For in vitro ablation experiments, we treated the organoids with 1 nM dimerizer (AP20187, Clontech). Drug-resistant organoid clones were manually selected and expanded individually. Genomic DNA was isolated using the QIAamp DNA blood mini kit (Qiagen). Legitimate knock-in was determined by PCR. Southern blotting was performed based on the standard procedure using 1 μg of genomic DNA. The sequences of PCR primers and Southern blot probes are shown in Supplementary Table 1. The puromycin-RFP selection cassette flanked by loxP sequences was excised by transient infection of Cre-expressing adenovirus (TaKaRa) at multiplicity of infection of 5–10. After the infection, we manually selected and cloned RFP− organoids. Deletion of the puromycin cassette was validated by PCR diagnostics. Percentage of successful knock-in for each line is shown in Supplementary Table 2. Once a knock-in reporter CCO was cloned, we used the same clone for further experiments. Organoids were dissociated into single cells with TrypLE Express (Life Technology), and large clusters were removed with a CellTrics 20-μm cell strainer (Partec). The cells were washed with cold PBS and stained with 7-amino-actinomycin D (7-AAD) staining solution (BD Biosciences) to exclude dead cells. Single cells were gated based on the SSC-H versus SSC-W profile. The cells were subsequently analysed using a flow cytometer with a 70-μm nozzle (FACS JAZZ, BD Biosciences). Then, 1,000 sorted cells were embedded in 25 μl of Matrigel and cultured in a 48-well plate for 7–10 days. We added 10 μM Y27632 for the first two days of culture, and the organoid colony formation was assessed using a BZX-700 fluorescence microscope (Keyence). RNA was extracted from 1 × 106 sorted cells using the RNeasy Plus mini kit (Qiagen). The RNA quality was determined by the RNA integrity number (RIN) value with the RNA6000 assay (Agilent). Only specimens with RIN > 7.0 were used in this study. Gene expression was determined by microarray (GeneChip PrimeView Human Gene Expression Array, Affymetrix) according to the manufacturer’s instructions. The data were normalized using the robust multi-array analysis implemented in the R package affy. The probes were summarized into genes by selecting probes with the highest median absolute deviation value per gene. GSEA was performed using gsea (in the R package phenoTest) with 1,000 permutations. Two independent intestinal stem cell signature gene sets from refs 17, 38 were used. All animal procedures were approved by the Keio University School of Medicine Animal Care Committee. NOD/Shi-scid,IL-2Rγnull (NOG) mice39 (7–12 weeks of age, male) were obtained from the Central Institute for Experimental Animals (CIEA, Japan). Organoids with the indicated genetic reporter and with or without GFP-luc2-reporter, equivalent to 1 × 105 cells, were xenotransplanted subcutaneously or into the renal subcapsules as previously described40. We monitored the tumour size with a calliper or through bioluminescence imaging. Tumour volumes were measured according to the formula (length × width2) / 2. Once any individual tumour reached 2 cm in size, the mouse was euthanized. For bioluminescence imaging, we intraperitoneally administered 3 mg of D-luciferin (SPI, Tokyo) to tumour-bearing mice 10–20 min before imaging and anaesthetized the animals with isoflurane. The bioluminescence signal was measured with an IVIS imaging system (Xenogen), and the specific signal was calculated as the ratio of photon counts from the region of interest to counts from a background region. The grafts were fixed for subsequent histological analyses. An investigator blinded to the experimental conditions measured the tumour sizes. For the lineage-tracing experiments, each mouse received a single intraperitoneal injection of 0.25 mg (clonal dose) or 1 mg of tamoxifen (Sigma-Aldrich) diluted in corn oil. For the ablation studies, 40 μg of dimerizer was administered for five days daily for short term ablation and on alternate days for long term ablation. To label the proliferating cells, we intraperitoneally administered BrdU (40 mg kg−1, BD Biosciences) and EdU (10 mg kg−1, Life Technologies) at the indicated times. For chemotherapeutic studies, CTX (40 mg kg−1, Merck Serono) or oxaliplatin (15 mg kg−1, AdooQ Bioscience) was administered intraperitoneally at the indicated times. We isolated tumours from xenografted mice and immediately fixed them with 4% paraformaldehyde. Eight-micrometre OCT frozen tissue sections or 5-μm paraffin-embedded tissue sections were processed using a standard histological protocol. For rainbow fluorescent imaging, the frozen sections were visualized using an SP8 confocal microscope (Leica) with the following settings: mCFP was excited at 405 nm and collected using a 480–485-nm filter, nuclear GFP was excited at 488 nm and collected using a 494–507-nm filter, EYFP was excited at 514 nm and collected using a 560–566-nm filter, and RFP was excited at 552 nm and collected using a 601–665-nm filter. Nuclei were counterstained with the near-infrared nuclear dye DRAQ5 (BioStatus). For DLS 3D imaging, the whole tumours were cut into 1–2 mm3 pieces, fixed and embedded in agarose gel. 3D images were acquired using the Leica SP8 DLS system. The proportion of surviving clones was determined by counting the number of RFP+ cells at day 3 and day 31 after tamoxifen administration. Clone identification and raw volume measurement were carried out automatically using the ImageJ 3D-image processing package ‘3D object counter’41, 42. False identification rate of this automatic measurement was determined manually by random sampling. Raw volume was adjusted by randomly subtracting a proportion of clones according to the false-rate. The threshold volume for total colonies on day 3 was set as <2 × 105 μm3 and for large colonies on day 31 as >2 × 105 μm3 (equivalent to 20 cells). Colony-formation efficiency was defined as the ratio of the number of large colonies on day 31 to the number of clones on day 3. For immunostaining, the following primary antibodies were used: mouse anti-cytokeratin-20 (M7019, clone K 20.8, Dako, 1:50), goat anti-GFP (ab6673, Abcam, 1:200), rabbit anti-Ki67 (ab16667, Abcam, 1:100), mouse anti-α smooth muscle actin ab-1 (MS-113-P, Thermo Scientific, 1:800), mouse anti-BrdU (347580, BD, 1:100), anti-cleaved caspase-3 (9661, Cell Signaling, 1:100) and anti-tdTomato (600-401-379, ROCKLAND, 1:500). Alexa Fluor 488-, 568- or 647-conjugated secondary antibodies (Life Technologies, donkey anti-mouse, rabbit, rat or goat antibodies) were used at 1:200 dilution. For EdU staining, we used the Click-IT Plus EdU Imaging kit (Life Technologies) according to the manufacturer’s instructions. Nuclei were counterstained with Hoechst 33342 or DAPI. Images were captured with a Leica SP8 confocal microscope or a BZX-700 fluorescence microscope (Keyence). To count the number of BrdU/EdU-stained cells, we used Imaris (Bitplane). In situ hybridization was performed using an RNAscope 2.5HD kit (Advanced Cell Diagnostics) according to the manufacturer’s instructions. For each experiment, we used PPIB and DapB genes as positive and negative controls, respectively. Tumour tissues were homogenized using TissueLyser LT (Qiagen) and RNA was extracted with the RNAeasy mini kit (Qiagen) according to the manufacturer’s instructions. cDNA was synthesized using the Omniscript RT kit (Qiagen). Quantitative real-time PCR was performed on LightCycler 96 (Roche Diagnostics) using FastStart Essential DNA Probes Master (Roche Diagnostics) and the cDNAs as templates. Relative LGR5 expression to ACTB was calculated based on the comparative C method. Primers and probes for LGR5 and ACTB are available in Supplementary Table 1. The sample size was determined by previous experience and preliminary experiments. The vehicle/dimerizer/chemotherapy-treated group was randomly assigned on the basis of tumour size at the time of injection. Appropriate statistical analyses were performed dependent on the comparisons referenced in the figure legends. The n values represent biological replicates. All graphs show mean and error bars represent the standard error of the mean (s.e.m.). Genetic mutation data of organoids are summarized in Supplementary Table 2 and described in ref. 16. The microarray dataset generated in this study is available in the Gene Expression Omnibus (accession number: GSE83513). All other data are available from the corresponding author upon reasonable request.


News Article | May 10, 2017
Site: www.nature.com

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.

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