How does precursor RNA structure influence RNA processing and gene expression?

The sequence of an RNA influences its biological activity. Sequence information still predominates as the most studied aspect of a nucleic acid. However, due to its single-stranded and flexible nature, RNA naturally forms structures as soon as it is synthesized by an RNA polymerase, reviewed in [1,2]. There is clear evidence for robust and reproducible RNA structures in RNA transcripts, both in human cells and several model organisms [3–8]. RNA structures can range from stable, consistent folds to flexible structural ensembles, reviewed in [9]. Transcriptome-wide structure analysis has revealed patterns of functional structure in processed RNAs, including flexible structural transitions around start and stop codons [3,5,6,8]. However, little is known about RNA structure in nascent human RNAs before they undergo processing to become mature transcripts. There is evidence that RNA structure is altered at different stages of an RNA molecule’s life cycle. Liu et al. found major differences between nuclear and cytoplasmic RNA structures in Arabidopsis, which suggested that RNA structures changed significantly from the nascent RNA to the mature transcript [10]. Structures in 3′UTRs can vary during different stages of development in zebrafish [11]. In yeast, there are differences in structure between the same RNAs in vivo versus RNAs extracted from the cell [4]. Direct study of nascent RNA structure is important to understand the relationship between precursor and mature RNA structures and how precursor RNA structure can impact RNA processing and gene expression. Although still limited, several studies have identified transcriptome-wide patterns of RNA structures in nascent RNAs [10,12].

Experimentally based secondary structural models of RNAs can be created with enzyme and chemical probing data, reviewed in [13]. In particular, chemical probing combined with next-generation sequencing has become very popular due to its ability to generate data on long RNAs and multiple RNAs at the same time [14,15]. Chemical probes include those that react with the ribose backbone (selective 2′-hydroxyl acylation analyzed by primer extension (SHAPE) reagents) and those that react with nucleobases, such as dimethyl sulfate and carbodiimides [16,17]. There are a wide variety of chemical probing applications (Table 1). While chemical probing primarily reveals secondary structure, there are other structure modeling techniques capable of measuring tertiary structures including cryo-electron microscopy (cryo-EM), nuclear magnetic resonance (NMR) and small angle X-ray scattering (SAXS). Recent advances in cryo-EM technology have resulted in structures of large RNA and protein (RNP) complexes like the spliceosome and ribosome [18,19], suggesting that deriving tertiary structural models for stable structures in large pre-processed RNAs may be possible. Secondary and tertiary RNA structural models can be modelled computationally and incorporate experimental chemical probing data [20]. These calculated structures can be based on combinations of thermodynamic parameters, machine learning and experiments, including highly cited methods like RNAstructure, mFOLD and others [21–27]. However, though less time consuming than experimentally generated structures, the accuracy of de novo models of RNA structure, both secondary and tertiary, is questionable [28–30].

A major hurdle in acquiring experimental secondary structure data from in vivo systems is that most methods depend on a significant number of molecules to quantify reactivity (Table 1, Cons). Thus, current in vivo methods are limited to high abundance transcripts. Low abundance transcripts that do not meet copy number thresholds require in vitro or more elaborate techniques to deduce structure [31,32]. The low abundance of precursor RNAs is one reason that their structures are understudied. Some techniques are beginning to address the problem of low abundance RNAs, including enrichment for low abundance targets during chemical probing (Table 1, RNA pull-down and Hybridization capture) [31,33,34]. Another hurdle in studying the structure of precursor RNAs is the flexible nature of RNA molecules. Although some RNAs have stable structures, such as transfer RNAs and RNAs found in the spliceosome and ribosome, most RNAs have many possible structures that are energetically similar. This ensemble effect of RNAs must be considered when determining what structures are biologically relevant. The long length of most introns compounds this problem for precursor RNAs. Computational modeling to analyze ensembles is being applied to assist in this problem [35–37]. Additionally, many RNAs are likely to have long-distance and tertiary interactions that may be functional but remain difficult to map [38,39]. Little is known about the importance of long-distance interactions in pre-processed RNAs. Experimental mapping techniques are available to document long-distance interactions (Table 1, Cross-linking). Finally, the nature of co-transcriptional processing adds a temporal element to precursor RNA structure modeling. Introns can be spliced out of order, making it difficult to predict which nucleotides are available for structural interactions during transcription of an RNA [40–42]. There are experimental and computational approaches that partially address the temporal nature of RNA folding by targeting temporally associated proteins (Table 1, Protein immunoprecipitation) [12,43]. However, the specialized approaches that address the problems of RNA structure modeling in dynamic, long and low abundance RNAs are not easily broadly applied.

There is no self-evident reason to assume that the structures that RNA molecules form are inherently functional. However, careful research has identified many functional RNA structures and their mechanisms. RNA structures can act to block recognition motifs. Short hairpin motifs that block recognition of the 5′ splice site are common and one of the earliest known structures to functionally impact splicing [44,45], reviewed in [46]. For example, the nascent MAPT (microtubule associated protein tau) RNA has a hairpin element at the 5′ splice site of exon 10 (Figure 1A). MAPT is important in neural biology and its precursor RNA is alternatively spliced to create at least 6 isoforms, reviewed in [47]. A splice junction in MAPT (exon 10–intron 10) is normally spliced at an equal ratio, resulting in mix of mature isoforms with either 3 or 4 microtubule binding domain repeats, which code for Tau proteins with different biological activity [48]. A hairpin at the MAPT exon10–intron10 junction directly overlaps with the 5′ splice site and can be disrupted by disease-associated variants [49,50]. The MAPT hairpin blocks normal spliceosomal recognition by the U1 snRNP at the 5′ splice site and causes exon skipping and formation of the shorter 3R MAPT mature transcript [51] (Figure 1A). RNA structure in pre-processed transcripts has been shown to block U1 interactions and alter splicing in other precursor RNAs in addition to MAPT, including SMN2, VWF, ATM and BCL2L1 [52–56] (Table 2).

RNA structure can also function to collapse the distance within long sequences to bring RNA elements into close proximity. Some nascent RNAs may use structure to condense long intronic regions to form structures that are suitable scaffolds for early spliceosome recognition and activation. The human AdML (adenovirus 2 major late transcript IVS1) precursor RNA has a global fold important for splicing [57]. Disrupting the structure of pre-processed AdML RNA prevents it from being recognized efficiently by the U1 snRNP in in vitro studies. FRET analysis confirms that the 5′ and 3′ splice sites are in close proximity in the normal structure, but not in poorly spliced mutants with altered structures [57]. In the AdML precursor RNA the global fold is not dependent on proteins. Similar studies of the human HBB (β globin) precursor RNA support the role of global RNA structure in recruiting U1 and promoting splicing [58] (Figure 1B). However, the global fold of the pre-processed HBB RNA is influenced by binding of SRSF1 protein as a structural stabilizing factor (Figure 1B). The ability of RNA structure to influence splicing by collapsing the distance between the branchpoint and the 3′ splice site has been documented in several introns in yeast [59]. RNA structure may also collapse the distance between the polyadenylation recognition motif and the cleavage site during 3′ processing and polyadenylation of nascent RNAs [60].

RNA structures can be recognized specifically, often in combination with sequence elements. Small nuclear ribonucleoprotein polypeptide A′ (SNRPA1) recognizes intronic sequence-independent stem structures combined with sequence-dependent loops to promote splicing of cassette exons in multiple genes, including plectin (PLEC) precursor RNA [61]. SNRPA1 splicing of PLEC contributes to a prometastatic cellular environment and is associated with progression and poor prognosis in breast cancer [61]. MBLN1 precursor RNA is autoregulated by MBLN1 protein binding at primarily unpaired YGCY motifs close to the 3′ splice site [62] (Figure 1C). Binding of MBLN1 to MBLN1 RNA restructures a distal branchpoint and results in exon 5 skipping and an isoform of MBLN1 protein with different subcellular localization [31] (Figure 1C). RNA structural elements can utilize multiple attributes of folding. In addition to containing structures that specifically bind MBLN1, the global fold of the MBNL1 exon 5 has also been shown to bring the 3′ and 5′ splice sites into close proximity [31,62]. MBLN1 protein also binds to loop elements with YGCY sequences in other nascent RNAs, including cardiac troponin RNA (Table 2) [63].

Despite technical difficulties in performing transcriptome-wide studies of precursor RNA structure, recent studies have broadly analyzed precursor RNA structure to identify global patterns of functional base pairing. Saldi et al. found that there are higher-order structural ‘steps’ that demarcate efficiently spliced and structured introns from less efficiently spliced exons in human cells [12]. The researchers used tNET-Structure-Seq to acquire structural data on nascent RNAs in human cell lines. tNET-Structure-Seq combines enzymatic and chemical structure probing with RNA Polymerase II immunoprecipitation to determine the accessibility of nascent RNA nucleotides. Introns spliced co-transcriptionally were associated with clearer structural ‘steps’ at the 5′ and 3′ splice sites when compared with splice junctions in introns that were spliced post-transcriptionally [12]. Structural ‘steps’ are characterized by a disparity in nucleotide accessibility around the exon–intron junction. Typical structural ‘steps’ have higher accessibility on the exonic side of the junction and lower accessibility on the intronic side of the junction (Figure 1D). The magnitude of a structural ‘step’ also leads to splicing preferences in cassette exons with bigger differences between the accessibility of the exon and intron leading to more efficient splicing. For example, mutually exclusive exons, a type of cassette exon, are primarily spliced post-transcriptionally and exon-inclusion or exclusion is influenced by RNA structure. Excluded exons had more robust ‘steps’ at the farthest 3′ splice site, whereas included exons were more likely to have weak ‘steps’ at the farthest 3′ splice site [12]. These finding are consistent with the general lack of base-pairing at the 3′ splice site and structural differences based on splicing efficacy found in Arabidopsis seedlings [10,64]. The pattern of unpaired, accessible exonic nucleotides and paired intronic nucleotides in efficiently spliced transcripts has also been found in mouse precursor RNAs [65] and other organisms [31,57,64–67], reviewed by [68,69].

All RNAs have structural patterns, many of which are functionally important, yet it can be difficult to identify which RNA structures are relevant without prior knowledge. Unlike many nascent RNAs, the survival of motor neuron 2 (SMN2) precursor RNA has been extensively mapped for structural elements [55,70–72], reviewed in [73]. SMN2 splicing studies have focused on exon 7, which can either be skipped or included. Two terminal stem-loops (TSL1 and TSL2) within exon 7 and several structures within intron 7 influence splicing (Figure 2A) [55,70,71,74]. These structures block U1 recognition of the exon 7 at the 5′ splice site and influence RNA-binding protein (RBP) interactions with enhancer and silencer functions. Inclusion of exon 7 allows SMN2 mature transcripts to produce active SMN protein and promote neural survival in the event of defects at the SMN1 locus [75,76]. Similar to exon 7 in SMN2, exon 3 of MCL1 can be skipped or included, resulting in the production of short (MCL1-S) and long (MCL1-L) RNA and protein isoforms with either pro- or anti-apoptotic functions [77]. In contrast with the extensive research into SMN2 RNA splicing, very little is known about how RNA structure influences RNA processing in most other nascent RNAs, including MCL1. We use SMN2 and MCL1 nascent RNAs as examples to discuss how to identify functional nascent RNA structures for validation.

One approach to identifying functional regions that may have important RNA structures is to take advantage of conservation data [78,79]. Highly conserved regions within a gene may have functional importance. Both SMN2 (Figure 2B,C) and MCL1 (Figure 2F,G) follow a typical conservation pattern with higher conservation values in the exonic regions and lower conservation in the intronic sequences. A region of SMN2 that stands out is the continuation of high conservation scores into the intronic region of intron 7 (Figure 2B,C, boxed). This region overlaps with TSL2 and internal stem loop 1 (ISL1), which sequester the 5′ splice site and promote exon 7 skipping [55]. Additional conserved regions overlap with the functional long-distance internal stem 2 (ISTL2) [71] and TSL4. In MCL1, high conservation values are also extended into intron 2, overlapping with the 3′ splice site and experimentally identified branchpoints [80] (Figure 2F,G, boxed). In addition to standard conservation data, covariation analysis can be used to identify functional structures by looking for regions where base pairing is conserved [81,82]. Covariation has been important for identifying functional structures in lncRNAs; however, it is not commonly detected in human protein coding RNAs [83,84]. Conservation data is not structure-specific, however, in SMN2 conservation data indicates important structural regions, suggesting that highly conserved regions in MCL1 may have structural significance.

Many nascent RNA processing steps rely on RBP interactions. Mapping RBP-binding sites onto RNA can highlight important regulatory regions. Structures nearby or that overlap with RBP sites may influence protein interactions. Experimentally determined RBP binding sites are available in published enhanced cross-linking and immunoprecipitation (eCLIP) databases, such as the ENCORE dataset in ENCODE [85–87]. In the ENCORE dataset, MCL1 has many RBPs bound to intronic and exonic regions, including SRSF1, which is known to affect MCL1-S to MCL1-L isoform ratios [88] (Figure 2H, boxes). The ENCODE SRSF1 binding site overlaps with a hotspot of RBP binding where 15 additional RBPs have been identified at the same position. The structures of hotspot elements may influence competitive RBP binding. Studies of MCL1 suggest that hnRNPF/H regulates MCL1 splicing and binds within intron 2 [89] (Figure 2H, boxed). However, each individual study is limited to select tissues and time-points. SMN2 is a neural factor and is not expressed in the ENCODE project cell lines. Only one RBP, U2AF2 is significantly associated with SMN2 in this dataset. However, experimental studies on SMN2 have been performed in multiple laboratories suggesting that more than 40 RBPs bind pre-processed SMN2 RNA around exon 7 and influence its splicing, reviewed in [90]. For example, the RBP TIA1 promotes SMN2 exon 7 inclusion by binding with intron 7 and recruiting the U1 complex in proximity to the 5′ splice site [91]. TIA1 binding overlaps with the intronic TSL3 structure (Figure 2D, boxed). Antisense oligonucleotides targeting SMN2 near the TIA1 site are predicted to open TSL3, make the TIA1 binding site more accessible, and promote exon 7 inclusion [72]. Predicting RBP sites is another option for genes that are less studied than SMN2 but not expressed in commonly used cell lines [92]. As we learn more about the preference of different RBPs for sequence and structural features we will be able to better predict the impact of RNA structures within RBP-binding sites. Since protein binding sites can be influenced by RNA structure, they are important regions for structural studies.

Another way to identify functional regulatory elements that may affect RNA processing is to map disease-associated variants onto the RNA, such as variants found through GWAS and familial studies and in databases such as ClinVar and HGMD [93,94]. Additional variant-based approaches can use quantitative trait loci (QTLs), which are variants associated with a variety of phenotypes like expression (eQTLs), splicing (sQTLs) and alternative polyadenylation (apaQTLs) [95–97]. However, there are generally few disease-associated variants mapped to genes, particularly in non-coding regions such as introns. In MCL1 and SMN2 there are only a handful of disease-associated variants (Figure 3B,E). Rare variants and somatic mutations may also be informative for identifying functional regions of an RNA [98,99]. To directly connect variants with RNA structure elements there are computational models to estimate the impact of a variant on local and global RNA structure [100–103], reviewed in [104]. Variants that change RNA structure are relatively common and are called riboSNitches [3,100,105], reviewed in [106]. We find that across SMN2 there are 95 riboSNitches, while in MCL1 there are 40 riboSNitches [102] (Figure 3B,E). RiboSNitches that overlap with rare or phenotypic variants are candidates that indicate functional structures that may be involved in phenotypic differences. In SMN2 a disease-associated variant found in ClinVar is also predicted to be a riboSNitch (Figure 3B, arrow). This riboSNitch falls within IS1, which is known to have structure that regulates SMN2 splicing [70]. Another riboSNitch in SMN2 overlaps with a somatic variant from COSMIC in the regulatory SMN2 intron 7 hairpin element 2 (Figure 3B, arrow) [74]. Similar somatic variants are predicted to be riboSNitches in MCL1, including a riboSNitch in close proximity to a ClinVar variant (Figure 3E, arrows).

Finally, rather than starting from functional elements and analyzing whether RNA structure may impact that function, we can also start from experimental or predicted structural models. There are many webservers and software packages that can predict minimum free energy models or base-pairing probabilities with reasonably high expected accuracy [21–26]. Within these structural models, highly probable hairpin/stem-loop motifs are common regulatory elements, reviewed in [107]. In SMN2, selection of strong hairpin elements from an experimentally based structural model for further study would have identified internal stem 1 (IS1), TSL2 and TSL3, all of which have been shown to structurally influence SMN2 splicing (Figure 3C) [55,70]. These elements are predicted despite the limited region selected for folding. In addition to these known structures in SMN2 there are two highly probably hairpin motifs within SMN2 intron 6 that could influence processing of SMN2 precursor RNA (Figure 3C). Structural models of SMN2 have been used by others to predict novel functional elements in SMN2 [73]. MCL1 precursor RNA also has highly probable hairpin and nested hairpin motifs in an experimentally based structural model (Figure 3F). In the nested hairpin, base-pairing is predicted between the branchpoint region and exonic sequence suggesting that this structure could affect 3′ splice site recognition and contribute to regulatory processing of MCL1 RNA.

Currently it is difficult to identify functional structures within a precursor RNA. SMN2 RNA is a well-studied model that shows the ability of conservation data, RBP binding site analysis, variant mapping and structural models to identify RNA structures that many influence SMN2 splicing. We also highlight regions within the understudied MCL1 RNA that may be of interest structurally to understand MCL1 splicing. As only a handful of human genes have clear precursor RNA structural annotation, the lack of known functional structures even in a highly regulated gene like MCL1 is unsurprising. The field will continue to acquire better secondary and tertiary structure models for SMN2 and MCL1 as new technologies for RNA structure modeling emerge, such as chemical probing techniques (Table 1) and cryo-EM. Although individual gene annotation for functional RNA structures is slow, an important goal is to identify enough functional structures in nascent RNAs to accurately predict functional structures directly from sequence.

Proteins are principal effectors of cellular function. Proteins that interact with RNA are common and many studies have explored the sequence and structural preferences of RBPs [108–111]. The interaction between RBPs and RNA can be understood as a modular interaction between RNA-binding domains (RBDs) in the protein and target RNA elements. There are 16 well known RBDs, and likely additional non-canonical domains, reviewed in [112]. RBDs can be repeated or varied within a protein and the spacing between RBDs can influence how the protein interacts with its target RNA. Interactions between RBDs and RNA elements are governed by basic physical principles: hydrogen bonding, electrostatics, and base-stacking, reviewed in [113]. All three of these mechanisms are used by most RBDs for structure and sequence specific interactions. Most RBPs have degenerate sequence preferences that favor structurally accessible RNA sequences [108]. In general, sequence-specific interactions between RBPs and RNA occur through hydrogen bonding. Hydrogen bonds cannot readily form when the target nucleotides are base paired. However, even though most RBPs target unpaired nucleotides for sequence specificity, structural context does influence RBP interaction. This allows RBPs to differentiate between RNA binding elements even when sequence motifs are degenerate or similar to other RBP binding sites [108].

One of the most common types of RBD is the RNA recognition motif (RRM). The U1A/SNF/U2A″ family is an example of RRM-containing proteins that can recognize structured RNA [114]. U1A is part of the spliceosome U1 snRNP. It binds specifically to the U1 stem-loop II (SLII) through structure specific base-stacking (Figure 4A, maroon), electrostatic interactions (Figure 4B, pink) and sequence-specific hydrogen bonding interactions (Figure 4A, orange) [115,116]. U1A binding is very specific and discriminates between stem-loops in U1 and U2 RNAs [117]. Despite this specificity, U1A is also capable of binding PIE RNA elements in its own 3′UTR and several other RNAs, including SMN2, to regulate polyadenylation [118,119] (Table 2). PIE elements are similar to the U1 SLII at the structure and sequence levels. Within a PIE element, duplicated stem-loops dimerize U1A (Figure 4C) and directly interact with polyA polymerase [120–122]. Although U2B contains a leucine-rich RBD rather than an RRM like U1A, it has a similar multifunctional ability. U2B interacts with the U2 RNA stem-loop IV with both structure and sequence specificity. However, in cancer cells, U2B has an extra-spliceosomal function wherein U2B binds intronic stem-loop structures and promotes splicing of cassette exons [61].

Double-stranded RNA-binding domains (dsRBDs) generally have strong structural preferences for their target RNAs. The cleavage factors Drosha and Dicer both contain dsRBDs and interact with precursor microRNAs through structure-dependent mechanisms [123,124]. MicroRNAs are processed from precursor RNAs (pri-miRNAs) that form extended stem-loops and are cleaved into pre-microRNAs and finally mature miRNAs. Pri-miRNAs interact with the microprocessor complex containing Drosha based on their helical RNA structure [125,126]. Drosha-dsRNA interactions are mediated by electrostatic interactions between the dsRBD (Figure 4D, gray) and two minor grooves, with minimal major groove interactions [127,128]. The extended stem-loop structure of pri-miRNAs is recognized at the transition between unpaired and paired segments at the bottom and top of the stem region. Flexibility in the pri-miRNA stem alters Drosha cleavage, ultimately resulting in the production of isoforms of mature miRNAs that can target different transcripts for translational repression [123]. The RBP Drosha and its partner DGCR8 recognize these structural junctions and structural changes in pri-miRNA, partially mediated by DDX3X, that result in different Drosha processing [123]. In addition to structure-specific recognition, Drosha also recognizes specific nucleotide sequences that are important for processing of pri-miRNAs. After Drosha processing, Dicer cleaves pre-miRNAs into duplex miRNAs. Dicer also recognizes structural elements of the pre-miRNA to discriminate between pre-miRNAs and other classes of RNA to cleave true pre-miRNAs into mature miRNAs [124].

While many RBPs have been extensively studied and their specificity determined, there are more than 2000 predicted RBPs in the human genome, and many ‘moon-lighting’ proteins with unrecognized RNA binding potential, reviewed in [112,113]. High-throughput studies to broadly characterize RNA-binding protein characteristics demonstrate the importance of both sequence and structural recognition [108,109]. Although the majority of RBPs prefer unpaired sequence motifs, they display varying sensitivity to structure. For example, RBPs including RBM22, RBM6, PRR3 and BOLL display preferences for motif-based partial pairing or a motif surrounded by paired nucleotides [108]. The little-known RBP ZNF326 even prefers sequence-based recognition within a completely paired structural motif [108]. By varying their ability to recognize structured sequences, RBPs effectively create a binding preference by incorporating the surrounding structural context of short sequence motifs [108]. Additional studies on the binding preferences of RBPs can help determine how RBPs recognize RNA and regulate gene expression.

While the sequence of an RNA is the primary input for many computational structure models, biological regulation affects the structure of RNA in the cell. One mechanism of regulation is the speed of transcription. Slow transcription by RNA pol II can influence the folding of known RNA structures, such as the hairpin structure at the 3′ end of histone transcripts [129]. Overall, slow transcription speeds increase base-pairing in nascent RNAs and lead to more efficient splicing [12]. The speed of RNA polymerase II transcription is a function of modifications to its carboxy-terminal domain; these modifications can be influenced by many biological processes, reviewed in [130,131]. Cell signaling by the myc pathway influences RNA pol II elongation speed, suggesting that RNA structure and processing can be globally altered under certain conditions, reviewed in [130,132]. In addition to global changes, individual gene loci may be prone to fast or slow transcription speeds based on their DNA and chromatin composition. For example, DNA G-quadruplexes influence transcription speed [133]. Additionally, several DNA- and RNA-binding proteins influence RNA pol II modification and elongation speed [134]. Each of these factors can be regulated by other cellular pathways, potentially making transcription speed a dynamic factor regulated globally and fine-tuned at individual loci. More research is required to explain how perturbation of transcription speed influences the ensemble of structures formed by a nascent transcript and how a structural ensemble may influence processing of the precursor transcript and its downstream output.

Nucleotide modifications are another cellular mechanism that can impact RNA structure. There are about 180 known types of RNA nucleotide modifications [135]. Two common modifications are methyladenosine (m6A) and pseudouridine (pseudoU), reviewed by [136,137]. Methylation of adenosine at the 6th position makes the residue more likely to be unpaired, resulting in changes to RNA structure than can affect RBP interactions [138,139]. Modification of uridine to pseudoU stiffens the RNA backbone, usually resulting in more stable RNA structure, but the impact on structure is dependent on the context of the pseudoU modification [140]. Lack of pseudoU modification is associated with altered structural dynamics in the ribosome, specifically in the way the sections of the ribosome rotate with respect to one another [141]. The ability of pseudoU to change the structure of the ribosome and modulate its dynamics probably contributes to the translational defect in cells with ribosomes that lack pseudoU [141]. Although modifications are the rule for noncoding RNAs such as ribosomal RNA and tRNAs, RNA modifications are also present in pre-processed mRNAs, reviewed in [137]. For example, both m6A and pseudoU are common in precursor RNAs and affect splicing [142–144]. Updates to computational modeling algorithms are beginning to consider the effect of m6A on the biophysical characteristics of structure folding [24]. The growing volume of functional RNA modifications suggests that these modifications constitute a dynamic cellular mechanism that regulates RNA structure.

RNA-binding helicases can change RNA structures in the cell. At least eight helicases, including DEAH-box helicases DHX19, DHX38, DHX8 and DHX15, are essential for splicing in human cells, reviewed in [145,146]. These helicases primarily assist with the release of splicing factors and precursor RNA at different stages of the splicing cycle. In addition, several helicases recognize stalled or improper splicing and are associated with degradation of these precursor RNAs [145]. In precursor microRNAs (pri-miRNAs), the DEAD-box helicase DDX3X impacts the structural flexibility of the pri-miRNA and influences alternative processing by Drosha [123]. DDX3X binds double-stranded RNA as a dimer, with one DDX3X primarily interacting with one RNA strand [147]. DDX3X regulation of pri-miRNAs results in differences in mature miRNA isoform composition across tissues and between normal and cancerous samples [123]. There are at least 64 human RNA helicases, primarily identified by homology to DEAD and DEAH helicases [148]. These helicases and other RBPs can alter RNA structure by mechanisms other than conventional helicase unwinding, reviewed in [112]. The interaction between helicases and RNAs is often dependent on the modification state of the helicase, allowing for dynamic regulation of RNA:protein interaction [149]. Additional research is needed to understand the function and regulation of helicases.

There is evidence that RNA structure is altered at different stages of an RNA molecule’s life cycle. Liu et al. found major differences between nuclear and cytoplasmic RNA structures in Arabidopsis, which suggested that RNA structure changed significantly from the precursor RNA to the mature transcript [10]. Structures in 3′UTRs can vary during different stages of development in zebrafish [11]. Differences in structure between RNAs in vivo versus purified from cells have been documented in yeast [4]. These studies demonstrate that RNA structure is not static. It is not clear in humans how structure is altered during processing of particular transcripts and whether refolding is a general characteristic that alters structure in predictable ways. Because RNA structure guides interactions with regulatory RBPs and nucleic acids, refolding of transcripts during processing has downstream implications for the fate of the transcript and gene expression.

In this review we have discussed multiple examples of how RNA structures in human precursor RNAs impact RNA processing and alluded to the effect of altered processing on subsequent gene expression. One impact of altered splicing is production of an alternative protein isoform (Figure 5A–C). In MAPT, when exon 10 is skipped the mature transcript produces a Tau protein with three rather than four microtubule binding domains (Figure 5A) [48]. These Tau protein isoforms have different biological activities and altering their ratio is correlated with development of frontotemporal dementia and other neurodegenerative diseases [48,49]. In SMN2, exon 7 is normally skipped, resulting in an unstable SMN protein isoform (Figure 5B) [150]. Spinal muscle atrophy occurs when neither SMN1 nor SMN2 can produce stable SMN protein. Exon 5 skipping in MBNL1 RNA results in loss of part of the bipartite nuclear localization element in MBNL1 protein and cell-wide localization rather than nuclear localization [151–153] (Figure 5C). The sequestration of MBNL1 in toxic repeats is an important factor in myotonic dystrophy [154]. RNA degradation can also be influenced by RNA structures that affect processing. For example, U1A precursor RNAs contains a PIE element that is bound by two U1A proteins [118]. Binding of U1A to its own transcript results in inhibition of polyadenylation and a decrease in U1A RNA [118,155] (Figure 5D). PIE elements in SMN2 and other transcripts also can be bound by U1A and inhibit polyadenylation, ultimately resulting in lower levels of RNA [118,119]. Altered RNA processing can also influence nonsense-mediated decay [156], RNA localization [157] and protein expression [158]. Nascent RNA structure has ripple effects on all aspects of the RNA life cycle and can contribute to human diseases.

Because RNA structure influences its functional interactions with other molecules, structure is a target for intervention, including at the nascent RNA stage. Antisense oligonucleotides (ASOs) can be designed to alter RNA structure, reviewed in [159,160]. In a structured RNA, bases normally interact in cis to form standard hairpins or stem-loop structures. An ASO can compete for base pairing in trans. The hybridization between the ASO and its target RNA opens up nucleobases for interaction with other nucleotides or proteins and could have global effects on structure. In the 7SK snRNP structural rearrangement is important for release of kinases involved in phosphorylation of the Poll II carboxy-terminal domain, leading to transcriptional control. ASOs that target sequences within 7SK dynamic hairpins block the structural transition of 7SK from one state to another and alter the ability of 7SK to regulate transcription [161]. The SARS-CoV2 corona virus has a highly structured single-stranded RNA genome [162]. One strategy currently under development for treatment of SARS infection is an ASO designed to disrupt a 3′ stem-loop involved in viral replication, reviewed in [163]. A similar mechanism of competing hybridization has been used to develop toehold switches, which switch structures in the presence of a particular RNA sequence to allow translation [164]. Toehold sensors have been developed for many applications including as a method to detect viral infections like SARS-CoV2 and Zika [165,166] and identify genomic variation [167]. There is software available to design toehold structures (NUPACK) [168]. Other hybridization methods that impact RNA structure have been developed to target transcriptional regulation [169].

Although hybridization offers a straightforward mechanism of structure change, ASOs are difficult to deliver to human tissue, whereas small molecules are generally more tractable for medical treatment. Small molecules can target and stabilize or destabilize specific RNA structures, reviewed in [170–172]. The capacity of small molecules to act on RNA structures was evident early on from bacterial riboswitches, which are designed to recognize a variety of different small molecules (e.g., metabolites) and change conformation to effect transcriptional or translational regulation, reviewed in [173]. Although most proof-of-principle molecules target noncoding or viral RNAs [174,175], small molecules that target RNA structures can be used to control nascent RNA processing. The FDA approved small molecule risdiplam has been developed to target SMN2 splicing at exon 7, reviewed in [176]. Although the exact mechanism of action is still under investigation, these molecules may function to stabilize the interaction between the U1 spliceosome and the 5′ splice site [177,178]. Modulating RNA structure to diagnosis or to treat disease is a rapidly growing field. Targeting function RNA structures in precursor RNAs is an important direction for therapeutic development.

RNA molecules are naturally structured. Due to the low abundance, long-length, and flexible nature of nascent RNAs, precursor RNA structure is understudied. New structural methods are continuing to advance our technological capabilities and document structures within precursor RNAs. In particular, chemical probing and cryo-EM methods have expanded our understanding of secondary and tertiary structures. However, even when structural models are available, it is difficult to identify functional structures and understand their mechanisms. Structures within precursor RNAs determine how nascent transcripts interact with protein and nucleic acid co-factors. By influencing these interactions, RNA structure influences RNA processing pathways. Most studies have focused on the impact of structure on splicing and polyadenylation, but future research may tell us more about how RNA structure affects other processing pathways like RNA editing. Due to their impact on processing, RNA structures impact gene expression and play a role in disease. We are beginning to develop antisense oligonucleotides and small molecule methods to alter RNA structure in vivo; these methods can be broadly applied to target functional RNA structures, including those that regulate RNA processing.

The authors declare that there are no competing interests associated with the manuscript.

This work was supported by the National Institutes of Health [grant number R35GM142851].

Austin Herbert: Conceptualization, Formal analysis, Investigation, Visualization, Writing—review & editing. Abigail Hatfield: Conceptualization, Writing—review & editing. Lela Lackey: Conceptualization, Visualization, Writing—original draft, Writing—review & editing.

The Genotype-Tissue Expression (GTEx) Project was supported by the Common Fund of the Office of the Director of the National Institutes of Health, and by NCI, NHGRI, NHLBI, NIDA, NIMH, and NINDS. The data used for the analyses described in this manuscript were obtained from: GTEx Analysis Release V8 (dbGaP Accession phs000424.v8.p2) from the GTEx Portal on 6/01/2022.

ASO

antisense oligonucleotide

dsRBD

double-stranded RNA-binding domain

eCLIP

enhanced cross-linking and immunoprecipitation

RBD

RNA-binding domain

RNP

Ribonucleoprotein

RRM

RNA recognition motif