A robust and versatile method for production and purification of large-scale RNA samples for structural biology.

Recent findings in genome-wide transcriptomics revealed that RNAs are involved in almost every biological process, across all domains of life. The characterization of native RNAs of unknown function and structure is particularly challenging due to their typical low abundance in the cell and the inherent sensitivity toward ubiquitous RNA degrading enzymes. Therefore, robust in vitro synthesis and extensive work-up methods are often needed to obtain samples amenable for biochemical, biophysical, and structural studies. Here, we present a protocol that combines the most recent advances in T7 in vitro transcription methodology with reverse phase ion pairing and ion exchange HPLC purification of RNAs for the production of yield-optimized large-scale samples. The method is easy to follow, robust and suitable for users with little or no experience within the field of biochemistry or chromatography. The complete execution of this method, for example, for production of isotopically labeled NMR samples, can be performed in less than a week.

Efficient Detection of Structure and Dynamics in Unlabeled RNAs: The SELOPE Approach.

The knowledge of structure and dynamics is crucial to explain the function of RNAs. While nuclear magnetic resonance (NMR) is well suited to probe these for complex biomolecules, it requires expensive, isotopically labeled samples, and long measurement times. Here we present SELOPE, a new robust, proton-only NMR method that allows us to obtain site-specific overview of structure and dynamics in an entire RNA molecule using an unlabeled sample. SELOPE simplifies assignment and allows for cost-effective screening of the response of nucleic acids to physiological changes (e.g. ion concentration) or screening of drugs in a high throughput fashion. This single technique allows us to probe an unprecedented range of exchange time scales (the whole µs to ms motion range) with increased sensitivity, surpassing all current experiments to detect chemical exchange. For the first time we could describe an RNA excited state using an unlabeled RNA.

A guide to large-scale RNA sample preparation.

RNA is becoming more important as an increasing number of functions, both regulatory and enzymatic, are being discovered on a daily basis. As the RNA boom has just begun, most techniques are still in development and changes occur frequently. To understand RNA functions, revealing the structure of RNA is of utmost importance, which requires sample preparation. We review the latest methods to produce and purify a variation of RNA molecules for different purposes with the main focus on structural biology and biophysics. We present a guide aimed at identifying the most suitable method for your RNA and your biological question and highlighting the advantages of different methods. Graphical abstract In this review we present different methods for large-scale production and purification of RNAs for structural and biophysical studies.

Production of Structured RNA Fragments by In Vitro Transcription and HPLC Purification.

The understanding of the functional importance of RNA has increased enormously in the last decades. This has required research on the RNA molecules themselves, with the concomitant need for obtaining purified RNA samples, such as for structural studies by NMR or other methods. The main method to create labeled and unlabeled RNA, T7 in vitro transcription, suffers from sequence-dependent yield and often low homogeneity for short constructs (<100 nt) and requires laborious purification. Additionally, the design of structured RNA fragments mimicking the structure of a larger biological RNA is often not straightforward. Secondary structure simulations can be used to make reliable predictions about the folding of a particular RNA fragment. In this article, we describe how to design an RNA construct of interest from a larger sequence, and we combine several previously published improvements of the in vitro transcription method, such as the use of 2'-methoxy modifications and dimethyl sulfoxide or the use of tandem repeats, to increase yield and purity of in vitro-transcribed RNA. Together with a high-performance liquid chromatography (HPLC) purification procedure using both reversed-phase ion-pairing and ion-exchange HPLC, we provide a robust protocol to obtain highly pure RNA of short to intermediate length in large quantities. The protocol optimizes yield, especially for RNA starting with nucleotides other than G. At the same time, it is simplified, and the required time is reduced. The protocols described here constitute a versatile pipeline for the production of purified RNA samples and are suitable for users with little experience in liquid chromatography. © 2021 The Authors. Basic Protocol 1: RNA construct design Basic Protocol 2: DNA template production and in vitro transcription Alternate Protocol: Tandem transcription and RNase H cleavage Basic Protocol 3: Reversed-phase ion-pairing HPLC purification Basic Protocol 4: Ion-exchange HPLC purification.

Mutate-and-chemical-shift-fingerprint (MCSF) to characterize excited states in RNA using NMR spectroscopy.

It is important to understand the dynamics and higher energy structures of RNA, called excited states, to achieve better understanding of RNA function. R1ρ relaxation dispersion NMR spectroscopy (RD) determines chemical shift differences between the most stable, ground state and the short-lived, low-populated excited states. We describe a procedure for deducing the excited state structure from these chemical shift differences using the mutate-and-chemical-shift-fingerprint (MCSF) method, which requires ~2-6 weeks and moderate understanding of NMR and RNA structure. We recently applied the MCSF methodology to elucidate the excited state of microRNA 34a targeting the SIRT1 mRNA and use this example to demonstrate the analysis. The protocol comprises the following steps: (i) determination of the secondary structure of the excited state from RD chemical shift data, (ii) design of trapped excited state RNA, (iii) validation of the excited state structure by NMR, and (iv) MCSF analysis comparing the chemical shifts of the trapped excited state with the RD-derived chemical shift differences. MCSF enables observation of the short-lived RNA structures, which can be functionally and structurally characterized by entrapment.

Choice of labeling and cell line influences interactions between the Fab fragment AbD15179 and its target antigen CD44v6.

Medical imaging by use of immunotargeting generally relies on a labeled molecule binding to a specific target on the cell surface. It is important to utilize both cell-based and time-resolved binding assays in order to understand the properties of such molecular interactions in a relevant setting. In this report we describe the detailed characterization of the interaction properties for AbD15179, a promising CD44v6-targeting antibody fragment for radio-immunotargeting. Influence of labeling and cell-line model on the protein interaction kinetics was assessed using three different labeling approaches ((111)In, (125)I and FITC) on three different squamous carcinoma cell lines. Interactions were measured using time-resolved assays on living cells, and further analyzed with Interaction Map®. Results demonstrated a general biphasic appearance of a high- and a low-affinity binding event in all cases. The relative contribution from these two interactions differed between conjugates. For (125)I-Fab, the population of low-affinity binders could be significantly increased by extending the chloramine T exposure during labeling, whereas the (111)In-labeling predominantly resulted in a high-affinity interaction. Interactions were also shown to be cell line dependent, with e.g. SCC-25 cells generally mediating a faster dissociation of conjugates compared to the other cell lines. In conclusion, we report both cell line dependent and labeling associated variations in interaction kinetics for AbD15179 binding to CD44v6. This has implications for cell-based kinetic assays and applications based on labeled conjugates in general, as well as in a clinical setting, where each individual tumor may create different kinetic profiles for the same conjugate.