Engineered aptamers for new compounds are typically produced by using in vitro selection methods. However, aptamers that are developed in vitro might not function as expected when introduced into complex cellular environments. One approach that addresses this concern is the design of initial RNA pools for selection that contain structural scaffolds from naturally occurring riboswitch aptamers. Here, we provide guidance on design and experimental principles for developing riboswitch-inspired aptamers for new ligands. The in vitro selection protocol (based on Capture-SELEX) is generalizable to diverse RNA scaffold types and amenable to multiplexing of ligand candidates. We discuss strategies to avoid propagation of selfish sequences that can easily dominate the selection. We also detail the identification of aptamer candidates using next-generation sequencing and bioinformatics, and subsequent biochemical validation of aptamer candidates. Finally, we describe functional testing of aptamer candidates in bacterial cell culture.

Key features

• Develop riboswitch-inspired aptamers for new ligands using in vitro selection.

• Ligand candidates can be multiplexed to conserve time and resources.

• Test aptamer candidates in bacterial cells by grafting the aptamer back onto its expression platform.

Graphical overview

The importance of studying the mechanistic aspects of long non-coding RNAs is being increasingly emphasized as more and more regulatory RNAs are being discovered. Non-coding RNA sequences directly associate with generic RNA-binding proteins as well as specific proteins, which cooperate in the downstream functions of the RNA and can also be dysregulated in various physiologic states and/or diseases. While current methods exist for identifying RNA–protein interactions, these methods require high quantities of input cells or use pooled capture reagents that may increase non-specific binding. We have developed a method to efficiently capture specific RNAs using less than one million input cells. One single oligonucleotide is used to pull down the target RNA of choice and oligonucleotide selection is driven by sequence accessibility. We perform thermal elution to specifically elute the target RNA and its associated proteins, which are identified by mass spectrometry. Ultimately, two target and control oligonucleotides are used to create an enrichment map of interacting proteins of interest.

Graphical abstract

Schematic representation of the SOCRATES workflow. SOCRATES utilizes a single 20-mer oligonucleotide for RNA pull down followed by a temperature elution series and liquid chromatography–mass spectrometry (LC-MS)/MS to identify specific RNA–protein interactions.

Palmitoylation refers to the modification of the cysteine thiols in proteins by fatty acids, most commonly palmitic acid, through ‘thioester bond’ formation. In vivo, palmitoylation of proteins is catalyzed by palmitoyl acyltransferases (PATs or DHHC-PATs). Palmitoylation has recently emerged as a crucial post-translational modification in malarial parasites. The expression and activity of palmitoyl transferases vary across different developmental stages of the malarial parasite’s life cycle. The abundance of palmitoylated proteins at a given stage is a measure of overall PAT activity. The PAT activity can also change in response to external signals or inhibitors. Here, we describe a protocol to ‘image’ palmitoyl-transferase activity during the asexual stages using Click Chemistry and fluorescence microscopy. This method is based on metabolic labeling of a clickable analog of palmitic acid by parasitic cells, followed by CuAAC (Copper-catalyzed Alkyne-Azide Cycloaddition reaction) Click Chemistry to render palmitoylated proteins fluorescent. Fluorescence allows the quantitation of intracellular palmitoylation in parasite cells across various development stages. Using this method, we observed that intracellular palmitoylation increases as the parasite transitions from ring to schizont stages and appears to be most abundant during the schizont stages in Plasmodium falciparum.

The 5′ cap is a ubiquitous feature of eukaryotic mRNAs. It is added in the nucleus onto newly synthesized pre-mRNA, and in the cytoplasm onto mRNAs after decapping or endonuclease cleavage. Cytoplasmic recapping can occur after loss of the cap at the native 5′ end, or downstream within the body of the mRNA. The identification and location of recapping events is key to understanding the functional consequences of this process. Here we present an approach that addresses this problem, using the Lexogen TeloPrime^® cDNA synthesis kit to tag recapped 5′ ends. TeloPrime uses a proprietary DNA ligase to add a double stranded DNA oligonucleotide onto the 3′ end of cDNA while it is base paired with mRNA. Specificity for capped ends is obtained by the oligonucleotide having an unpaired C residue that base pairs weakly with ^m7G on the mRNA 5′ end. This is followed by PCR amplification of double-stranded cDNA using primers to the appended oligonucleotide and the mRNA of interest. The resulting products are gel purified and sequenced directly (if a single band) or cloned and sequenced. The sequence at the junction between the ligated oligonucleotide and the target mRNA provides the location of the cap on the corresponding transcript. This assay is applicable to all capped transcripts. It can be used with Sanger sequencing for small numbers of transcripts or adapted for use with Illumina library sequencing.

Studying monocytic cells in isolated systems in vitro contributes significantly to the understanding of innate immune physiology. Functional assays produce read outs which can be used to measure responses to selected stimuli, such as pathogen exposure, antigen loading, and cytokine stimulation. Integration of these results with high quality in vivo models allows for the development of therapeutics which target these cell populations. Current methodologies to quantify phagocytic function of monocytic cells in vitro either measure phagocytic activity of individual cells (average number of beads or particles/cell), or a population outcome (% cells that contain phagocytosed material). Here we address technical challenges and shortcomings of these methods and present a protocol for collecting and analyzing data derived from a functional assay which measures phagocytic activity of macrophage and macrophage-like cells. We apply this method to two different experimental conditions, and compare to existing work flows. We also provide an online tool for users to upload and analyze data using this method.