“Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Michael Elowitz ( melowitz@caltech.edu ).

Fertile chicken (Gallus gallus) eggs, purchased from commercial sources, were incubated at in a humidified 37 C incubator, and staged by the criteria of Hamburger and Hamilton (HH) (). Embryos were electroporated at stage 12-13, replaced in the incubator, and dissected 20h later.

All cell lines used in this paper contained stable integrations of transgenes, and were typically clonal populations. To create each stable cell line, the following steps were followed: 1) Cells were first transfected with 800-1000 ng of plasmid DNA using Lipofectamine 2000 or Lipofectamine LTX. 2) 24 h later, cells were transferred to selection media containing 600 ug/ml Geneticin, 500 ug/ml Hygromycin, 400 ug/ml Zeocin, or 10 ug/ml Blasticidin as appropriate. 3) After selection for 1-2 weeks, the resulting polyclonal populations stably expressing the transgene were allowed to recover for ∼1 week. 4) Single clones were isolated through the technique of limiting dilution. 5) Single clonal populations were screened for desired behavior, usually high expression (for constitutive genes) or low background expression of the transgene and large dynamic range (for inducible genes and reporter genes). Cell lines incorporating multiple transgenes were constructed by sequential rounds of this process. For piggybac constructs, the initial transfection comprised of the target plasmid along with the construct expressing the piggybac transposase, typically in a 1:1 or 2:1 molar ratio.

C2C12 cells (Mouse cells, RRID:CVCL_0188, ATCC Catalog No. CRL-1772) were grown in DMEM (Life Technologies), supplemented with 20% Tet System Approved FBS (ClonTech), 100 U/ml penicillin, 100 ug/ml streptomycin, 0.584 mg/ml L-glutamine (GIBCO). C2C12 media was used for CHO-K1 + C2C12 co-culture assays ( Figure S4 ). All cells were grown at 37°C in 5% COin a humidified atmosphere. Cells were passaged every 2-3 days, depending on confluency, using 0.05% or 0.25% Trypsin-EDTA (Life Technologies).

CHO-K1 (Hamster cells, RRID:CVCL_0214, ATCC Catalog No. CCL-61) or CHO- TREx (RRID:CVCL_D586, Invitrogen) cells and their derivatives were grown on tissue-culture grade plastic plates (Thermo Scientific) in Alpha MEM Earle’s Salts (Life Technologies), supplemented with 10% Tet System Approved FBS (ClonTech), 100 U/ml penicillin, 100 ug/ml streptomycin, 0.292 mg/ml L-glutamine (GIBCO).

All constructs used in this paper were assembled using standard restriction enzyme-based cloning and/or Gibson cloning (). pcDNA3-hNECD-Gal4 ( Figures 1 2 , and 5 ) has been described previously (). The H2B-3xCitrine fluorescent reporter ( Figures 1 2 , and 5 ) was constructed by cloning 3 repeats of mCitrine in frame with H2B, downstream of a UAS promoter. The mRNA destabilized version of this reporter was constructed by fusing the 3′UTR of mouse Hes1 downstream of the STOP codon. Ligand constructs were cloned into pcDNA5 or piggyBac plasmids (System Biosciences Inc.) by fusing the complete rat Dll1 (kind gift from G.Weinmaster) or human Dll4 cDNA in frame with T2A-H2B-mCherry, downstream of a previously described inducible pCMV-TO promoter (). We note that hDll1 shows the same pulsatile behavior described here for rDll1. Chimeric ligands ( Figure 5 ) were constructed by exchanging the intracellular domains of rDll1 (aa 561 – 714) and hDll4 (aa 551 – 685). The hN1ΔECD gene ( Figure 3 ) was cloned from hN1 (kind gift from J. Aster) by removing residues 22-1716 and fused in frame with myc-T2A-H2B-mCherry, downstream of the CMV-TO promoter in a piggyBac construct. Constructs used for in ovo electroporation ( Figure 4 ) were made by cloning rDll1 or hDll4 cDNA (minus stop) upstream of, and in frame with, T2A-EGFP in a pCI-CAGG plasmid.

Method Details

Co-culture assays and time-lapse microscopy 2, 5, S1, Used in Figures 1 S2 , and S5 Surface treatment In preparation for plating of cells, glass-bottom multi-well plates (MatTek, No. 1.5 glass, 10 mm radius) were coated with 5 ug/ml Hamster Fibronectin (Oxford Biomedical Research) diluted in 1x Phosphate-Buffered Saline (PBS) for 1h at room temperature. Cell culture After trypsinization, sender cells (pre-induced for > 48h with 4-epiTc, Sigma) or CHO-K1 cells were mixed in suspension with similarly trypsinized receiver cells at a ratio of 100:1 or 1:100, for excess sender or excess receiver assays, respectively. A total of 8x104 cells (60% confluence) were plated for each experiment, with continued 4-epiTc induction when appropriate. Imaging commenced 2-4h post-plating. Time-lapse microscopy Movies were acquired at 20X (0.75 NA) on an Olympus IX81 inverted epi-fluoresence microscope equipped with hardware autofocus (ZDC2) and an environmental chamber maintaining cells at 37C, 5% CO2. Automated acquisition software (METAMORPH, Molecular Devices) was used to acquire images every 30 min in multiple colors (YFP, RFP, CFP) or differential interference contrast (DIC), from multiple stage positions.

Plate-bound Dll1 assay Used in Figures S1 D and S1E Recombinant human Dll1ext-Fc fusion proteins (kind gift from I. Bernstein) were diluted to 1 ug/ml in PBS, and the solution was used to coat the tissue-culture surface. After 1h incubation at room temperature, the solution was removed, and cells were plated for the experiment.

Image segmentation, tracking, and single-cell fluorescence calculation 2, 5, S1, Used in Figures 1 S2 , and S5 Custom MATLAB code (2013a, MathWorks) was used to segment cell nuclei in images based on constitutive CFP/RFP fluorescence or background YFP fluorescence. The segmentation procedure uses edge detection, adaptive thresholds, and the Watershed algorithm to detect nuclear edges. Nuclear segments were then matched in pairs of images corresponding to consecutive time frames, and thus tracked through the duration of the movie. Single-cell tracks were subsequently curated manually. In particular, there were periods where any given cell could not be automatically segmented (typically due to high density) but could be visually followed. In such cases, the tracks corresponding to the cell prior to and after such time frames were manually linked if fewer than ∼5 frames were missing. Fluorescence data was extracted from nuclear segments by calculating the integrated fluorescence within the segment and subtracting a background fluorescence level estimated from the local neighborhood of the segment. This fluorescence was linearly interpolated across time frames where nuclei could not be segmented automatically. Division events were detected automatically, and fluorescence traces were corrected for cell division by adding back fluorescence lost to sister cells. The resulting ‘continuized’ traces were smoothed and the difference in fluorescence between consecutive time frames was calculated. A smoothed version of this difference was used as the rate of change or promoter activity of the fluorescence.

Analysis of single-cell traces 2, 5, S1, Used in Figures 1 S2 , and S5 Alignment For each receiver cell trace, including those of cells in control conditions (showing background fluorescence levels) an average rate of fluorescence increase (‘average slope’) was calculated by dividing the change in total fluorescence of the reporter by the duration of the trace. Traces showing activation were automatically selected for further analysis based on their average slopes surpassing a threshold value, chosen to be higher than average slopes observed in receiver cells under control conditions. Activating traces were aligned at the point of activation, defined as the time point when their promoter activity crosses an absolute threshold level, chosen based on typical promoter activities corresponding to background activity. Note that activations occurring during the first 15h of the movie were typically not considered, to eliminate transient effects produced by cell transfer to imaging conditions. The same thresholds were always used when direct comparisons were made between ligands or conditions, and we verified (by varying threshold levels) that qualitative results did not depend strongly on the choice of threshold. For C2C12 dynamics ( Figure S4 ) promoter activity could not be reliably used to align traces due to the low levels of reporter activity and resulting noise in the promoter activity data. These traces were instead aligned based on when the total fluorescence levels increased a threshold level. Double-pulse alignment th percentile) in the 0-7.5h window during which the first pulse is expected to reach maximum levels. Starting at 7.5h, i.e., after the peak of the first pulse, traces were re-aligned at the point when the subsequent promoter activity values cross Peak1, and re-normalized to the 90th percentile of values in the period from 7.5h (relative to the first activation point) to the end of the trace. In order to align traces showing two pulses in response to Dll1 ( Figure S1 D) at the second pulse, the following procedure was used: the first activation was determined using the usual procedure (see above). Traces were then normalized by the peak activity (‘Peak1’, 95percentile) in the 0-7.5h window during which the first pulse is expected to reach maximum levels. Starting at 7.5h, i.e., after the peak of the first pulse, traces were re-aligned at the point when the subsequent promoter activity values cross Peak1, and re-normalized to the 90percentile of values in the period from 7.5h (relative to the first activation point) to the end of the trace. Normalization th percentile value during the analysis time window, except in th percentile value occurring within 15h after activation. When applied, the object of normalizing the response trace by its amplitude is to demonstrate its stereotyped features, such are relative rise time and duration. Un-normalized averaging would distort the shape of the response because higher-amplitude signals are also prolonged, since the timescales of the reporter are fixed by the half-lives of its components (Gal4 protein, H2B-3xCitrine mRNA) and do not scale with amplitude. Traces were typically normalized to the 90percentile value during the analysis time window, except in Figure S2 H, where traces were normalized to the 90percentile value occurring within 15h after activation. Amplitudes th percentile of (absolute, non-normalized) promoter activity values between 0 and 7.5h (after alignment) in the traces. This time window is chosen to simultaneously estimate the promoter activity at the peak of pulses and at steady-state levels of sustained signaling. In th percentile of promoter activity values during the 25h after activation (the period over which activities are averaged). While normalized traces were used to make comparisons of the stereotyped shapes of responses (see above), absolute values of promoter activity, calculated from non-normalized promoter activity, are reported in all amplitude comparisons. Except in Figure 2 C, this amplitude represents the 95percentile of (absolute, non-normalized) promoter activity values between 0 and 7.5h (after alignment) in the traces. This time window is chosen to simultaneously estimate the promoter activity at the peak of pulses and at steady-state levels of sustained signaling. In Figure 2 C, the amplitude represents the 95percentile of promoter activity values during the 25h after activation (the period over which activities are averaged). Trace filtering th percentile) in the 0-7.5h period (after alignment). This criterion was designed to automatically detect single pulses in the data. In In Figure 1 D, traces were included in the Dll1 alignment if the median promoter activity between 20-25h fell below 50% of the peak activity (95percentile) in the 0-7.5h period (after alignment). This criterion was designed to automatically detect single pulses in the data. In Figure 2 B traces were only included in the Dll1 alignment if the normalized value at 20h fell below 0.7. This filter eliminates traces consisting of multiple pulses, especially in the high Dll1 cases. A similar filter applied to Dll4 traces reveals a small fraction of cells activated transiently, but displaying qualitatively different behavior, such as a systematic increase in duration and amplitude with increasing Dll4 levels in senders. For C2C12 experiments in Figures 3 G and 3H, activating cells were identified based on an increase in total fluorescence levels above a threshold.

Estimating Gal4 and mRNA half-lives, Related to Figure S1H Gal4 after inhibition of the pathway using DAPT, at time 0h. d G a l 4 d t = − γ G a l 4 G a l 4

Reporter mRNA m is produced through non-cooperative binding of Gal4 to the promoter, with dissociation constant K and maximum rate β m . m is degraded with rate constant γ m . d m d t = β m G a l 4 K + G a l 4 − γ m m

The parameters γ G a l 4 , K and γ m were calculated by fitting the Citrine mRNA m to the experimentally measured decay in Citrine fluorescence rate using the lsqnonlin function in MATLAB. The fit was constrained using bounds for γ G a l 4 and γ m of log(2)/5h – log(2)/3h, based on Sprinzak et al. (2010) Sprinzak D.

Lakhanpal A.

Lebon L.

Santat L.A.

Fontes M.E.

Anderson G.A.

Garcia-Ojalvo J.

Elowitz M.B. Cis-interactions between Notch and Delta generate mutually exclusive signalling states. Bintu et al. (2016) Bintu L.

Yong J.

Antebi Y.E.

McCue K.

Kazuki Y.

Uno N.

Oshimura M.

Elowitz M.B. Dynamics of epigenetic regulation at the single-cell level. For this model, we assume that the free Gal4 protein produced due to cleavage of N1ECD-Gal4 degrades with first-order kinetics with rate γafter inhibition of the pathway using DAPT, at time 0h.Reporter mRNAis produced through non-cooperative binding of Gal4 to the promoter, with dissociation constantand maximum rateis degraded with rate constantThe parametersandwere calculated by fitting the Citrine mRNAto the experimentally measured decay in Citrine fluorescence rate using the lsqnonlin function in MATLAB. The fit was constrained using bounds forandof log(2)/5h – log(2)/3h, based onand. Bootstrapped 95% confidence intervals were computed from 100 iterations of fitting 30 points, chosen randomly with replacement, out of a total 50 measured time points.

Mathematical model for estimating duration of Notch activation, Related to Figure S1J Gal4 for a duration τ act , and degrades with first-order kinetics with rate γ Gal4 . d G a l 4 d t = { β G a l 4 − γ G a l 4 G a l 4 , t ≤ τ a c t − γ G a l 4 G a l 4 , t > τ a c t }

Reporter mRNA m is produced through non-cooperative binding of Gal4 to the promoter, with dissociation constant K and maximum rate β m . m is degraded with rate constant γ m . d m d t = β m G a l 4 K + G a l 4 − γ m m

For the results of Gal4 = 1, β m = 1, and K = 6.6 (also fitted in For this model, we assume that Gal4 is produced at a rate βfor a duration τ, and degrades with first-order kinetics with rate γReporter mRNAis produced through non-cooperative binding of Gal4 to the promoter, with dissociation constantand maximum rateis degraded with rate constantFor the results of Figure S1 , β= 1, β= 1, and K = 6.6 (also fitted in Figure S1 E), and estimated mean values from Figure S1 E were used for the Gal4 and mRNA degradation rates.

Simulations of Dll1 pulse trains and analysis, Related to Figures S2 A–S2F This model constructs pulse-trains composed of Dll1-like pulses occurring at various frequencies and regularities based on each of three underlying pulse models, and analyzes the features of the resulting simulated signaling traces. Pulse train construction ( Figure S2 B) For each simulation we construct 200 pulse trains. Each pulse train is constructed from a series of pulses with the average Dll1 promoter activity pulse shape ( Figure S1 I), scaled by an amplitude randomly sampled from the empirically measured distribution of Dll1 pulse amplitudes (from the Figure 1 D dataset). The first pulse occurs at 0h, representing activation at time 0 in the aligned Dll4 traces. Subsequently, new pulses are introduced after successive time intervals τ chosen based on one of the underlying pulse models (see below), and the composite signal is constructed until it extends at least 10h beyond the 25h time period averaged in Figure 1 D. Feature analysis ( Figure S2 D) 1) Amplitude: The amplitude of each constructed trace is its median value over 25h.

2) Intra-trace variability: After calculation of the amplitude, each trace is normalized to its 90th percentile value. For each point t in this trace, the local temporal variability is estimated by the standard deviation of values in a 10h window starting at t. The overall intra-trace variability calculated for each trace is the median of the local variability value at each point, calculated by moving a 10h time window through the trace. For each trace, two features are analyzed: For each simulation (200 constructed traces), the medians of the calculated amplitudes and intra-trace variability are tabulated, and the SEM calculated. Pulse models ( Figure S2 C) 1) period , that can range from 1h to 8h. Since the Dll1 pulse decay becomes apparent after 7.5 h ( period greater than 8h are not considered in the simulation. Periodic model: In this model, the interval τ between adjacent pulses is fixed at a value T, that can range from 1h to 8h. Since the Dll1 pulse decay becomes apparent after 7.5 h ( Figure 1 D), intervals greater than 8h will result in pulse trains in which the individual pulses can be clearly discerned in each trace, and the average behavior will show oscillations. Since neither individual Dll4 traces, nor the average shape display overt oscillatory features, values for Tgreater than 8h are not considered in the simulation.

2) Poisson model: In this model, the interval between successive pulses i and i+1, τ i , represents the inverse of a pulse rate, r i , drawn from a Poisson distribution with parameter, λ, ranging from 1/h-1/15h.

3) Mixed model: In these models, the interval τ between adjacent pulses is drawn from a normal distribution with mean T period (range 1h - 15h) and standard deviation σ (2.5h or 5h). This model therefore combines the regular pulsing inherent to the periodic model with the trace-to-trace variability of the Poisson model (thus preventing ‘constructive interference’ of pulse peaks, which would lead to apparent oscillations in the average signal shape). Three models are considered for the underlying pulsing process: period , λ, or σ, as appropriate) in each of the models, 36 simulations were run and the average of the median amplitudes and median intra-trace variabilities (see above) were calculated. These values are plotted in For every parameter value (T, λ, or σ, as appropriate) in each of the models, 36 simulations were run and the average of the median amplitudes and median intra-trace variabilities (see above) were calculated. These values are plotted in Figure S2 E. Bootstrapped analysis of variability in measured Dll4 signaling trace ( Figure S2 F) Finally, for direct comparison to simulation data, the Dll4 dataset of traces (200 traces in total) was subsampled 30 times (50 traces per sample) to generate a bootstrapped distribution of measured median intra-trace variability, and a corresponding median value was calculated. This bootstrapped median is compared to simulation data in Figure S2 F.

Sender cell categorization in excess receiver assays Used in Figures 2 and S2 Dll1- and Dll4-T2A-H2B-mCherry sender cells were induced with different 4epi-Tc concentrations, to access their full dynamic range of ligand expression. Following co-culture with receiver cells and timelapse analysis, individual sender cell nuclei were automatically segmented, and mCherry levels were calculated. At the same time, each receiver cell response was automatically associated with the closest sender cell. All data, across 4epi-Tc induction levels, were then pooled, and sender cells re-categorized into ‘low’, ‘medium’, or ‘high’ expression along with their associated receiver cell responses. This process of pooling and recategorization was necessary because of the broad, overlapping distributions in mCherry expression produced by 4epi-Tc treatment.

Detection of surface ligand Used in Figure S5 ext-Fc chimeric protein (R&D Systems) was used for surface-detection of ligands at a concentration of 10 ug/ml, based on a previously described protocol ( LeBon et al., 2014 LeBon L.

Lee T.V.

Sprinzak D.

Jafar-Nejad H.

Elowitz M.B. Fringe proteins modulate Notch-ligand cis and trans interactions to specify signaling states. ext-Fc protein in binding solution (blocking solution containing 100 ug/ml CaCl 2 , R&D Systems) for 45 min at RT. Following this, cells were washed 3x with binding solution, then incubated with anti-mouse secondary antibody conjugated to AlexaFluor-488 (1:1000 dilution, Life Technologies) for 30 min. Cells were then trypsinized and analyzed using flow cytometry. Recombinant mouse Notch1-Fc chimeric protein (R&D Systems) was used for surface-detection of ligands at a concentration of 10 ug/ml, based on a previously described protocol (). Sender cells were first cultured and induced with 4epiTc for 48h, then transferred from media to blocking solution (2% FBS in Phosphate Buffered Saline, PBS) for 30 min at room temperature (RT). Cells were then incubated with recombinant mouse Notch1-Fc protein in binding solution (blocking solution containing 100 ug/ml CaCl, R&D Systems) for 45 min at RT. Following this, cells were washed 3x with binding solution, then incubated with anti-mouse secondary antibody conjugated to AlexaFluor-488 (1:1000 dilution, Life Technologies) for 30 min. Cells were then trypsinized and analyzed using flow cytometry.

C2C12 N1ΔECD activation assays Used in Figures 3 and S3 The procedure for activating the Notch pathway in C2C12-hN1ΔECD cells was as follows: Cells were cultured in 10 μM DAPT (Sigma-Aldrich) until the experiment. In order to wash out DAPT, cells were washed quickly twice and a third time for 5 min with media at room temperature. Finally, cells were incubated in medium containing the appropriate activating DAPT concentration (0, 0.3, or 0.5 μM) at 37 C for the required activation duration (5 min, 15 min, 30 min, or until RNA extraction, i.e., sustained). In order to generate a pulse of activation, medium was then replaced with fresh 10 μM DAPT medium.

RNaseq Used in Figures 3 and S3 RNA was prepared using the RNeasy kit (QIAGEN) and submitted to the Caltech sequencing core facility, where cDNA libraries for RNaseq were prepared according to standard Illumina protocols. 100 base single-end read (100SR) sequencing was performed on a HiSeq2500 machine at the same facility. Reads were assembled, aligned, and mapped to the mouse genome (mm9 assembly) on a local instance of the Galaxy server, using Tophat. Cufflinks was used to calculate FPKM values. Castel et al., 2013 Castel D.

Mourikis P.

Bartels S.J.J.

Brinkman A.B.

Tajbakhsh S.

Stunnenberg H.G. Dynamic binding of RBPJ is determined by Notch signaling status. In the analysis, we focused first on genes that showed > 5 fold-changes in their FKPM values (highlighted in Table S1 ). We further narrowed our subsequent analyses to the transcription factors Hes1, Hey1, and HeyL, because their promoters were shown to directly bind NICD by ChIP-Seq, they show early and strong (> 10-fold) responses to NICD, and they are key factors mediating Notch responsive behaviors in many contexts. These are also the only Hes and Hey family genes that activate in response to Notch in C2C12 cells (). The RNaseq experiment did show upregulation of other genes, but we did not focus on them either because they were not transcription factors (such as Jag1 or Nrarp), or were not direct NICD targets based on the ChIP-Seq data.

RT-qPCR Used in Figures 3 and S3 RNA was prepared using the RNeasy kit (QIAGEN). cDNA was prepared from 500ng RNA using the iScript cDNA synthesis kit (Bio-Rad). 0.5 μL cDNA was used per 10 μL RT-qPCR reaction mix containing 1X iqSYBR Green Supermix (Bio-Rad) and 450 nM total forward and reverse primers. Reactions were performed on a BioRad CFX Real-Time PCR Detection System using a 2-step amplification protocol, with the following thermocycling parameters: 95 C, 3 min followed by 40 cycles of 95 C, 10 s (melting) and 55 C, 30 s (annealing + extension). All reactions were performed in duplicate.

Western blot analysis of NICD Used in Figure S3 For this analysis, 0.5x106 - 1x106 cells were trypsinized after treatment, spun down in excess PBS, and lysed using Lithium Dodecyl Sulfate (LDS) buffer also containing reducing agents (DTT + 2-Mercaptoethanol) and Protease Inhibitors (Roche). Standard procedure was used for LDS-PAGE gel electrophoresis and transfer to nitrocellulose (iBlot, Thermo Fisher Scientific). Cleaved NICD (1:1000, Cell Signaling Technology, Catalog # D3B8) and GAPDH (1:5000, Abcam, Catalog #6C5) were detected using monoclonal antibodies. The blots were subsequently stained using HRP-conjugated secondary antibodies and detected using the Enhanced ChemiLuminescence system (Pierce).

CHO-C2C12 co-culture assay Used in Figure S4 In preparation for the co-culture, C2C12-hN1 cells (4-6x104 cells in 12 well multi-well plate wells) were transfected with 60 pmol siRNA directed against mouse Notch2 (5′-UGAACUUGCAGGAUGGGUGAAGGUC-3′), using Lipofectamine RNAiMAX (Life Technologies). 24h later, 3x104 CHO-K1 based Dll1- and Dll4- sender cells (pre-induced for > 48h) were plated within the two chambers of ibidi culture inserts (Ibidi USA) on hamster fibronectin-treated (5 μg/ml in PBS, incubated for 3-5h at RT) surfaces of 24-well glass bottom plate wells. Once cells had attached to the surface (< 6h), inserts were removed and previously prepared C2C12-hN1 cells were plated, in 5 μM DAPT media, at high density so as to cover the gaps on the surface. After 12h, DAPT was washed out and cells were allowed to signal for 6h, after which the cultures were fixed in 4% formaldehyde at room temperature for 10 mins.

in ovo Electroporation Used in Figures 4 and S4 Batches of eggs were selected at random for electroporation with either Dll1 or Dll4, and the final data represents experiments conducted on at least two separate batches. The neural tubes of HH stage 12-13 embryos were injected with plasmid DNA (5 mg/ml) and electroporated by applying a series of current pulses (25V, 5x, 30 ms pulses separated by 100 ms) at the level of the pre-somitic mesoderm. 20h post-electroporation, embryos were screened for GFP fluorescence. Healthy embryos showing strong fluorescence in the neural crest were dissected (to remove extra-embryonic tissue) in Ringer’s solution and transferred to freshly prepared 4% paraformaldehyde, on ice. Embryos were fixed overnight at 4 C.

Hybridization Chain Reaction Fluorescence In Situ Hybridization Used in Figures 4 and S4 Choi et al., 2016 Choi H.M.T.

Calvert C.R.

Husain N.

Huss D.

Barsi J.C.

Deverman B.E.

Hunter R.C.

Kato M.

Lee S.M.

Abelin A.C.T.

et al. Mapping a multiplexed zoo of mRNA expression. The hybridization chain reaction fluorescence in situ hybridization (HCR-FISH) protocol was based on a previously described protocol (). Briefly, in situ HCR-FISH detection involves the following steps: 1. Dehydration and rehydration of embryos in MeOH, 2. Overnight hybridization with probes at 45 C, 3. Removal of unbound excess probes through washes at 45 C, 4. Overnight amplification at room temperature, and 5. Removal of excess amplifier. Each gene of interest was detected using 6 probes. At most three genes were detected simultaneously, typically EGFP, MyoD1, and Hes1, Hey1, or HeyL. After HCR processing, portions of the embryos anterior to the forelimbs were removed. Embryos were then mounted on glass-bottom multiwell plates in 1% agarose, with the dorsal surface in contact with the glass.

Confocal laser-scanning microscopy of embryos Used in Figures 4 and S4 Samples were imaged on a Zeiss LSM700 or using a 20x (0.8 NA) dry objective. For embryos, Z stacks were acquired using Zen software (ZEISS) and 3D-reconstructed in Imaris 8.0 (Bitplane). Optical slices in Imaris were used to remove obscuring auto-fluorescence from residual extra-embryonic tissue in the reconstructed images, without affecting signal in the areas of interest. For cell-culture Z stacks, the sum was projected in 2D using ImageJ.

Quantitation of effect on MyoD1 and Notch targets Blind scoring of embryos for changes in MyoD1 (Used in Table 1 Rios et al., 2011 Rios A.C.

Serralbo O.

Salgado D.

Marcelle C. Neural crest regulates myogenesis through the transient activation of NOTCH. 3D images of transverse optical sections of the interlimb region of the trunk (containing 3-5 pairs of somites per image), were sorted randomly, and then scored blindly for differences in somite MyoD1 levels between the electroporated and control sides of the embryo. The scoring procedure was as follows: any features that might reveal the specific experimental perturbation (Dll1 or Dll4 ectopic expression), such as image filenames, differences in pseudo-color attributes, or information from secondary channels, were removed before the files were re-ordered using a pseudorandom sequence. Subsequently, images were scored blindly, comparing MyoD1 signal in somites on the electroporated side with signal in the corresponding somites on the control side, as long as the two somites were level with each other. This requirement minimizes imaging artifacts. Finally, sample images were re-matched with the perturbation type and scores were tallied. The number of embryos scored per condition (11 Dll1 expressing embryos, 10 Dll4 expressing embryos, 61 somites for each perturbation) is standard for this type of quantification (). Quantification of fold-changes in MyoD1, Hes1, and Hey1 gene-expression (Used in Figure S5 C) The DML regions of the somites on the electroporated and control sides were manually identified in Z-projections of 3D-reconstructed confocal images (see above), and the maximal HCR-FISH staining intensities (90th percentile values within identically-sized areas on both sides) were calculated. The reported fold-changes represent the ratio of these values for electroporated versus control DMLs.

Immunofluorescence detection of transendocytosed Notch in co-cultures Used in Figures 5 and S5 Sender cells and receiver cells were co-cultured on glass-bottom dishes, in the excess sender configuration, as described above. After 24h of co-culture, cells were fixed in 4% formaldehyde (diluted in PBS). All subsequent steps were carried out in blocking solution (2% Bovine Serum Albumin diluted in PBS). Following 1h of incubation at room temperature, samples were incubated overnight at 4 C with 1:250 mouse anti-hNotch1 (Biolegend Catalog No. 352014, RRID AB_10899408 ). Samples were then washed and incubated in an anti-mouse secondary antibody conjugated to Alexa Fluor 488 (Life Technologies). After room temperature washes, samples were permeabilized in 0.3% Triton X-100 (Sigma-Aldrich) for 1h. Samples were then again incubated in 1:250 anti-hNotch1 overnight at 4C, following which they were incubated in Alexa Fluor 647 conjugated anti-mouse antibody (Life Technologies).