RNA-protein interactions drive fundamental biological processes and are targets for molecular engineering, yet quantitative and comprehensive understanding of the sequence determinants of affinity remains limited. Here we repurpose a high-throughput sequencing instrument to quantitatively measure binding and dissociation of MS2 coat protein to >107 RNA targets generated on a flow-cell surface by in situ transcription and inter-molecular tethering of RNA to DNA. We decompose the binding energy contributions from primary and secondary RNA structure, finding that differences in affinity are often driven by sequence-specific changes in association rates. By analyzing the biophysical constraints and modeling mutational paths describing the molecular evolution of MS2 from low- to high-affinity hairpins, we quantify widespread molecular epistasis, and a long-hypothesized structure-dependent preference for G:U base pairs over C:A intermediates in evolutionary trajectories. Our results suggest that quantitative analysis of RNA on a massively parallel array (RNAMaP) relationships across molecular variants.

Highlights d A thermodynamic model quantitatively predicts PUM1/2 binding to any RNA sequence d Factors beyond simple recognition of consecutive residues influence binding d Comparison to X-linking data reveals thermodynamic control of PUM2 binding in cells d Analysis of RNA structure effects suggests disruption of RNA structure in cells

RNA-binding proteins (RBPs) control the fate of nearly every transcript in a cell. However, no existing approach for studying these posttranscriptional gene regulators combines transcriptomewide throughput and biophysical precision. Here, we describe an assay that accomplishes this. Using commonly available hardware, we built a customizable, open-source platform that leverages the inherent throughput of Illumina technology for direct biophysical measurements. We used the platform to quantitatively measure the binding affinity of the prototypical RBP Vts1 for every transcript in the Saccharomyces cerevisiae genome. The scale and precision of these measurements revealed many previously unknown features of this well-studied RBP. Our transcribed genome array (TGA) assayed both rare and abundant transcripts with equivalent proficiency, revealing hundreds of low-abundance targets missed by previous approaches. These targets regulated diverse biological processes including nutrient sensing and the DNA damage response, and implicated Vts1 in de novo gene "birth." TGA provided single-nucleotide resolution for each binding site and delineated a highly specific sequence and structure motif for Vts1 binding. Changes in transcript levels in vts1Δ cells established the regulatory function of these binding sites. The impact of Vts1 on transcript abundance was largely independent of where it bound within an mRNA, challenging prevailing assumptions about how this RBP drives RNA degradation. TGA thus enables a quantitative description of the relationship between variant RNA structures, affinity, and in vivo phenotype on a transcriptome-wide scale. We anticipate that TGA will provide similarly comprehensive and quantitative insights into the function of virtually any RBP.RNA | next-generation sequencing | systems biochemistry | RNA binding proteins | Vts1 R NA-binding proteins (RBPs) constitute 5-10% of the eukaryotic proteome (1-3) and collectively govern the localization, translation, and decay of virtually every transcript (4-6). Despite the ubiquity of RBPs and their central importance in gene regulation, decoding the links between RNA primary sequence and its cadre of regulators remains a major unresolved challenge (7). Current approaches for characterizing RBP function generally involve trade-offs between throughput, comprehensiveness, and quantitative precision. Biophysical measurements can be made with targeted biochemical approaches such as electrophoretic mobility shift assays (EMSAs) or fluorescence polarization (FP) (8, 9), but these methods can only interrogate known RNA-protein interactions and are inherently lowthroughput. Selection-based approaches [e.g., in vitro selection, high-throughput sequencing of RNA, and sequence-specificity landscapes (SEQRS)/RNA bind-n-seq (RBNS)] achieve higher throughput, but these techniques remove binding sites from their natural sequence context and identify "winners" based on more than simple affinity (10). Transcriptome-wide methods, which often use cross-linking and immunoprecipi...

The quantification of protein-ligand interactions is essential for systems biology, drug discovery, and bioengineering. Ligand-induced changes in protein thermal stability provide a general, quantifiable signature of binding and may be monitored with dyes such as Sypro Orange (SO), which increase their fluorescence emission intensities upon interaction with the unfolded protein. This method is an experimentally straightforward, economical, and high-throughput approach for observing thermal melts using commonly available real-time polymerase chain reaction instrumentation. However, quantitative analysis requires careful consideration of the dye-mediated reporting mechanism and the underlying thermodynamic model. We determine affinity constants by analysis of ligand-mediated shifts in melting-temperature midpoint values. Ligand affinity is determined in a ligand titration series from shifts in free energies of stability at a common reference temperature. Thermodynamic parameters are obtained by fitting the inverse first derivative of the experimental signal reporting on thermal denaturation with equations that incorporate linear or nonlinear baseline models. We apply these methods to fit protein melts monitored with SO that exhibit prominent nonlinear post-transition baselines. SO can perturb the equilibria on which it is reporting. We analyze cases in which the ligand binds to both the native and denatured state or to the native state only and cases in which protein:ligand stoichiometry needs to treated explicitly.The interaction of proteins with ligands is a fundamental aspect of biomolecular function. The quantification of such interactions is essential for systems biology, drug discovery, and bioengineering. Traditional, generalizable methods for measuring protein-ligand affinity such as equilibrium dialysis or isothermal titration calorimetry require large amounts of material or radiolabeled ligands. Ligand-induced changes in protein stability also provide a general, quantifiable signature of protein-ligand interactions (1-6), and hence of biological function (7). The measurement of protein stability typically also has required relatively large amounts of protein and low-throughput instrumentation and, consequently, has not been widely used as a tool to assess function. However, recently a number of techniques that allow protein stabilities to be determined with small amounts of material in a high-throughput manner have been developed (8-12). One such method is based on extrinsic fluorescent dyes that monitor protein (un)folding (2, 3, 13). This technique uses the relatively inexpensive fluorescent dye Sypro Orange (SO) 1 (14) in combination with readily available real-time polymerase chain reaction (RT-PCR) instrumentation (3, 13, 15) and is being adopted as a straightforward, economical, high-throughput screening tool for ligand discovery in the pharmaceutical industry (4, 15), structural genomics efforts (16,17), and high-throughput protein engineering (18). Here we present insights into the prop...