Eukaryotic tRNA splicing – one goal, two strategies, many players

Janina L. Gerber; Sandra Köhler; Jirka Peschek

doi:10.1515/hsz-2021-0402

Artikel Open Access

Eukaryotic tRNA splicing – one goal, two strategies, many players

Janina L. Gerber , Sandra Köhler und Jirka Peschek

Veröffentlicht/Copyright: 30. Mai 2022

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Aus der Zeitschrift Biological Chemistry Band 403 Heft 8-9

Abstract

Transfer RNAs (tRNAs) are transcribed as precursor molecules that undergo several maturation steps before becoming functional for protein synthesis. One such processing mechanism is the enzyme-catalysed splicing of intron-containing pre-tRNAs. Eukaryotic tRNA splicing is an essential process since intron-containing tRNAs cannot fulfil their canonical function at the ribosome. Splicing of pre-tRNAs occurs in two steps: The introns are first excised by a tRNA-splicing endonuclease and the exons are subsequently sealed by an RNA ligase. An intriguing complexity has emerged from newly identified tRNA splicing factors and their interplay with other RNA processing pathways during the past few years. This review summarises our current understanding of eukaryotic tRNA splicing and the underlying enzyme machinery. We highlight recent structural advances and how they have shaped our mechanistic understanding of tRNA splicing in eukaryotic cells. A special focus lies on biochemically distinct strategies for exon-exon ligation in fungi versus metazoans.

Keywords: enzyme catalysis; ligase; nuclease; protein structure; RNA processing

Introduction

The cellular RNA pool is immensely diverse and complex, with various classes of RNAs functioning in a wide spectrum of biological processes. During their biosynthesis, RNA molecules undergo a vast number of co- and post-transcriptional processing and modification steps, which require the activity of dedicated enzymes. One unique example of RNA processing is the protein-catalysed splicing of transfer RNAs (tRNAs), an essential step during tRNA maturation (Phizicky et al. 1992). Intron-containing tRNA genes are found in all three kingdoms of life. However, the biochemical strategies for tRNA splicing differ between phylogenetic groups (Yoshihisa 2014). In this review, we focus on the eukaryotic tRNA splicing machinery including recent advances in identifying new “players” and understanding their structure-function relationship.

tRNAs are ubiquitous and highly abundant small non-coding RNA molecules that deliver amino acids to ribosomes and thereby aid in translating the genetic code. This primary function is conserved among the three domains of life. In eukaryotic cells, numerous additional functions of tRNAs in signalling, cell viability and metabolic regulation have been uncovered (reviewed in Kirchner and Ignatova 2015; Phizicky and Hopper 2010). Correct tRNA biosynthesis including transcription, modification and post-transcriptional processing by dedicated enzymes is essential for cell viability.

In eukaryotes, tRNAs are transcribed as precursor transcripts (pre-tRNAs). While pre-tRNA processing is initiated in the nucleus, the cellular path towards mature tRNAs takes place at different subcellular locations. The complex dynamics between nuclear and cytoplasmic tRNA processing as well as the precise order of all steps are not fully understood. After export into the cytosol, pre-tRNAs are subjected to multiple further post-transcriptional processing events before they can fulfil their function as mature molecules. A subset of pre-tRNAs contains an intervening intronic sequence (intron) that needs to be removed during maturation. These intron-containing pre-tRNAs undergo a fully enzyme-catalysed splicing reaction. This reaction occurs in two steps: The intron is first excised by a tRNA splicing endonuclease (SEN) and the resulting tRNA exon halves are ligated by tRNA ligase to form the fully matured, functional tRNA (Greer et al. 1983; Peebles et al. 1983). The subcellular localisation of these events seems to differ depending on the organism and is not conclusively solved. In contrast to the yeast SEN complex, which acts in the cytoplasm and is attached to the outer mitochondrial membrane (Yoshihisa et al. 2007, 2003; Zahedi et al. 2006), the human SEN (also named TSEN) complex shows nuclear localisation (Paushkin et al. 2004). Whether the sites of pre-tRNA cleavage also determine the localisation of the tRNA ligases, remains an open question (Lopes et al. 2015).

Mature tRNA adopts a secondary structure composed of four helical stems resembling a cloverleaf structure with a central four-way junction (Figure 1A). At the tertiary structure level, tRNAs fold into an “L” shape, where the acceptor domain and the anticodon domain are perpendicular to each other (Figure 1B). At the acceptor domain end, amino acids are attached via the last nucleotide of the 3′ CCA tail in the acceptor stem. The opposite end of the molecule harbours the anticodon, which pairs with its complementary codon on the messenger RNA during translation (Kim et al. 1974). Introns found in eukaryotic tRNA genes are mostly inserted at one nucleotide 3′ to the anticodon (Hopper 2013). Because intron insertion at this canonical position disrupts the anticodon stem-loop structure, the removal of the introns is an essential process. The phylogenetic origin, as well as the function of tRNA introns are still being explored. After excision, the intron is either degraded by exoribonucleases (Wu and Hopper 2014) or circularised by a tRNA ligase. The resulting tRNA intronic circular RNA (tricRNA) is a recently identified RNA species of unknown cellular function (Lu et al. 2015; Schmidt et al. 2019).

Figure 1:

The eukaryotic tRNA splicing process.

(A) Schematic overview of the two separate, enzyme-catalysed cleavage and ligation reactions. The 5′ exon is depicted in purple, the 3′ exon in yellow and the intron in grey. The intron is circularised and forms tricRNA (grey dashed pathway). (B) Crystal structure of the yeast tRNA^Phe (PDB: 6TNA). The anticodon is highlighted in red.

The combined activities of an endonuclease and an RNA ligase catalyse the splicing of tRNAs. Depending on the organism, various additional enzymes and structural proteins are involved (which we discuss in later sections of this review). Together, these proteins form the tRNA splicing machinery, which performs the intricate process of splicing intron-containing pre-tRNAs during their maturation (Lopes et al. 2015; Schwer et al. 2004). Knowledge about the function and structure of the core enzymes and additional tRNA splicing factors has expanded vastly during the last years.

Intron removal by splicing endonucleases

The endonucleolytic removal of tRNA introns is the essential first step during tRNA splicing. It depends on two precise cuts by the SEN complex at the exon-intron boundaries (Ho et al. 1990; Trotta et al. 1997). Despite differences in length, location and abundance of the introns, tRNA splicing in archaea is an enzyme-catalysed mechanism that is similar to the eukaryotic pathways (Yoshihisa 2014). Most of our structural understanding of intron removal stems from the archaeal SEN complex (Kleman-Leyer et al. 1997; Li et al. 1998). The cleavage sites in archaeal pre-tRNAs are defined by a bulge-helix-bulge (BHB) motif (Kjems and Garrett 1988; Marck 2003; Tang 2002). Archaeal SEN complexes form varied quaternary structures such as a homodimer (Fujishima et al. 2011; Kleman-Leyer et al. 1997), homotetramer (Li et al. 1998; Lykke-Andersen 1997) or dimer of homodimers (Mitchell et al. 2009). In eukaryotes, SEN is a hetero-tetrameric complex comprised of two catalytic subunits, SEN2 and SEN34, as well as two structural proteins, SEN54 and SEN15 (Figure 2B) (Paushkin et al. 2004; Trotta et al. 1997). The eukaryotic SEN complex was first identified and characterised by Abelson and co-workers in the yeast Saccharomyces cerevisiae (Trotta et al. 1997). They showed that two dimers, SEN2/SEN54 and SEN34/SEN15, assemble into the tetrameric holo-complex. While there is no solved structure of the fully assembled tetramer, a recent study revealed the structure of a truncated SEN34/SEN15 dimer (Sekulovski et al. 2021).

Figure 2:

State of knowledge on the fungal tRNA splicing machinery.

(A) Enzyme-mediated steps of the intron excision and exon ligation processes. The structure of tRNA with intron was modelled with RNA-Composer out of the crystal structure of tRNA^Phe (PDB: 6TNA). The 5′ exon is depicted in purple, the 3′ exon in yellow and the intron in grey. The circled part of the tRNA structure is enlarged and simplified represented. The colour of the arrows corresponds to the enzyme structure colour. (B) Model of splicing endonuclease complex composition. The tetrameric enzyme consists of two dimeric subcomplexes (dark and light orange) each consisting of a catalytic (Pacman-like circle) and a structural subunit (rectangle). A pre-tRNA molecule is schematically depicted so that the two splice sites at the 3′ and the 5′ exons can be cleaved by SEN2 and SEN34, respectively (indicated by arrows). (C) Domain organisation of Trl1 enzyme. The crystal structure of the ligase domain (LIG) is shown in blue (PDB: 6N67), the polynucleotide kinase domain (KIN) in pink and the cyclic phosphodiesterase (CPD) in green (PDB: 6U05). The linker region between the CtTrl1 and CaKIN-CPD structures is depicted as dashed line. Enzyme and tRNA structures are depicted in relative size.

Intron removal generates two tRNA halves: The 5′ half possesses a terminal 2′,3′-cyclic phosphate (2′,3′-cP), and the 3′ half possesses a 5′-hydroxyl (Figure 2A). A catalytic triad similar to other archaeal or eukaryotic endonucleases acts in the cleavage of the pre-tRNA. The catalytic subunit SEN2 is responsible for cleavage at the 5′ splice site and SEN34 cuts at the 3′ splice site of the pre-tRNA (Ho et al. 1990; Trotta et al. 1997). However, the precise mode of splice site recognition by eukaryotic SEN has remained largely unknown. The cleavage site in eukaryotes is determined by the distance between the cleavage site and a conserved anticodon-intron (A-I) pair in the anticodon stem-loop (Baldi et al. 1992; Di Nicola Negri et al. 1997; Greer et al. 1987; Reyes and Abelson 1988). The correct distance to the 5′ splice site is established by a suggested “molecular ruler” mechanism in which the distance to the splice site is “measured” by SEN54 and its interaction with the D-arm of the tRNA (Hayne et al. 2020; Reyes and Abelson 1988; Sekulovski et al. 2021; Trotta et al. 2006; Xue 2006; Yoshihisa 2014).

Using a co-immunoprecipitation approach combined with mass spectrometry, Paushkin et al. identified the polyribonucleotide 5′-hydroxyl-kinase 1 (CLP1) as additional pre-tRNA cleavage factor associated with the human TSEN complex (Paushkin et al. 2004). CLP1 possesses a kinase activity and is responsible for the phosphorylation of the 5′ end of the 3′ exon (Weitzer and Martinez 2007). CLP1 is conserved among eukaryotes but its function within the SEN complex remains unknown.

Exon-exon joining by tRNA ligases

Succeeding endonucleolytic digest by the SEN complex, the pre-tRNA exon halves, containing a 2′,3′-cP at the 5′ exon end and a 5′-OH group at the 3′ exon end, are joined by a dedicated tRNA ligase (Englert et al. 2011; Peebles et al. 1983; Popow et al. 2011; Tanaka et al. 2011). In general, RNA ligases are found in all domains of life. They catalyse the ligation of RNA molecules via phosphodiester bonds during different RNA processing events, such as repair, editing and splicing. The term “tRNA ligase” refers to a functional group of tRNA splicing enzymes belonging to distinct RNA ligase classes (Trl1: EC 6.5.1.3, RTCB: EC 6.5.1.8). Despite the conceptual similarity, the biochemical mechanisms of tRNA exon joining differ vastly depending on the organism. There are two main branches in eukaryotes that can be divided into a fungal (also employed in plants) as well as a metazoan/human strategy. During the final step of tRNA splicing, the tRNA exon halves are ligated by the tRNA ligase Trl1 in fungi (Englert 2005; Greer et al. 1983; Phizicky et al. 1986) or the multi-protein RTCB ligase complex in metazoans (Popow et al. 2011).

The fungal strategy

In fungi, the pre-tRNA exon halves cannot be ligated directly after endonucleolytic digest by the SEN complex (Greer et al. 1983). Two different enzymatic steps proceed to modify the chemical groups at both RNA termini in preparation for their final ligation (Figure 2A). This generation of the two biochemically distinct ends, also referred to as “healing”, occurs by RNA end-modifying enzymes (Schwer et al. 2004).

The essential fungal enzyme needed for modifying the tRNA ends is the same as for ligation. Trl1 (previously named Rlg1) is a tripartite enzyme (Figure 2C), consisting of (I) an N-terminal adenylyltransferase domain (ligase; LIG), which belongs to the nucleotidyltransferase superfamily alongside DNA ligases and RNA capping enzymes, (II) a C-terminal cyclic phosphodiesterase domain (CPD) and (III) a central polynucleotide kinase (KIN) domain (Phizicky et al. 1986; Xu et al. 1990). The CPD activity opens the 2′,3′-cP to form a 3′-OH/2′-phosphate terminus, and the KIN activity phosphorylates the 5′-OH in a GTP-dependent manner. All three enzymatic activities are essential for cell viability (Phizicky et al. 1992). The crystal structure of full-length, tripartite Trl1 still remains unresolved. We speculate that the flexible linker regions between the individual enzymes hamper crystallisation of the full-length protein (Figure 2C). However, recent crystallographic efforts yielded several atomic-resolution structures of the individual subdomains derived from various fungal species (Banerjee et al. 2019a, 2019b; Peschek and Walter 2019; Remus et al. 2017). They resulted in a couple of well-resolved snapshots of the three catalytic domains providing insights into protein dynamics, co-factor dependency and substrate binding.

The Candida albicans Trl1-KIN structure explains the preference for GTP as phosphate donor during 5′-OH phosphorylation. The structure of the kinase domain with GDP and Mg²⁺ in the active site reveals a P-loop phosphotransferase fold, together with a unique G-loop element that explains its guanine nucleotide specificity (Remus et al. 2017). Further, structures of the kinase-dead (D445N variant) Trl1-KIN domain apoenzyme and in complexes with GTP·Mg²⁺, IDP·PO₄, dGDP·PO₄, and GDP·Mg²⁺ reveal conformational changes in the G-loop (Banerjee et al. 2019b) upon different states of the bound nucleotide. While there is currently no crystal structure of the separate Trl1-CPD, the bipartite C. albicans Trl1-KIN-CPD was solved allowing for a detailed understanding of the C-terminal Trl1 domain (Banerjee et al. 2019b). In this structure, the two catalytic domains within the same polypeptide chain do not show any relevant interaction but are rather connected by an unstructured linker (Figure 2C). CPD harbours a conserved central β-sheet with two His-X-Thr motifs (with X indicating any canonical amino acid). Dual presence of this motif is characteristic for the 2H phosphoesterase superfamily, which is named after their two conserved histidine residues (Mazumder et al. 2002). A phosphate ion in the active site identifies some of the residues involved in substrate binding that are part of the conserved active site motif within fungal 2H enzymes (Banerjee et al. 2019b).

In the tripartite fungal tRNA ligase Trl1, the LIG domain catalyses the last step of joining the tRNA halves. The LIG activity seals the healed ends through phosphodiester bond formation. This final nucleotidyl transfer reaction is ATP-dependent and progresses via three steps (Figure 3A). First, a covalent Trl1-LIG-(lysyl-ζ)-AMP intermediate is formed between the active site lysine and ATP. Subsequently, the bound AMP is transferred to the 5′-PO₄ end and forms a 5′-5′ RNA-adenylate. Finally, Trl1-LIG catalyses the attack by the 3′-OH on the RNA-adenylate to form a phosphodiester bond, thereby releasing AMP (Greer et al. 1983; Phizicky et al. 1986). Despite being identified by Abelson and co-workers in the 1980s, there have been no structures of Trl1 or any of its separate domains until recently. Crystal structures of the N-terminal ligase have been solved from the thermophilic fungus Chaetomium thermophilum (Banerjee et al. 2019a; Peschek and Walter 2019). They provide new structural insights into the active site residues and nucleotide binding of Trl1-LIG. Crystal structures of Trl1-LIG exist in three distinct states: (I) bound to a α,β-non-hydrolysable ATP analogue (Peschek and Walter 2019) (Figure 2C), (II) a covalent LIG-(lysyl-Nζ)–AMP·Mn²⁺ intermediate and (III) a LIG·ATP·(Mn²⁺)₂ complex, the latter two as K148M ligase-dead variant (Banerjee et al. 2019a). These structures reveal a two-domain architecture comprised of an N-terminal canonical adenylyltransferase domain and a C-terminal domain with a unique all-helical fold. This building principle of a conserved adenylyltransferase domain in conjunction with a unique C-terminal fold is also present in other RNA and DNA ligases (Shuman and Lima 2004). Previous structural and enzymatic data on the tRNA repair enzyme from Bacteriophage T4 Rnl1 show that its C-terminal domain is not required to catalyse RNA ligation in general but confers specificity for tRNA substrates (El Omari et al. 2006). Based on these observations and the structural similarities between the adenylyltransferase domains of CtTrl1-LIG and T4Rnl1, a similar substrate specificity-conferring function of the C-terminal domain of CtTrl1-LIG is conceivable but remains to be uncovered.

Figure 3:

Fungal Trl1 and archaeal RtcB tRNA ligase active site structure and reaction equations of the ligation step.

(A) Chaetomium thermophilum Trl1 (PDB: 6N67) active site with bound AMPcPP and SO₄²⁻. (B) Pyrococcus horikoshii (PDB: 4IT0) RtcB active site with bound GMP and Mn²⁺. Active site amino acids (Lys148 in Trl1 and His404 in RtcB) are depicted in yellow. Nucleotide interacting amino acids are presented in grey. Sulfate ion (yellow) and Mn²⁺ (green) interacting amino acids are presented in cyan. The respective phosphodiester bond formation reactions during the ligation step are summarised below the active site panels (see main text for details).

The covalent LIG-(lysyl-Nζ)–AMP·Mn²⁺ intermediate and a LIG·ATP·(Mn²⁺)₂ complex reveal a two-metal mechanism. A metal complex stabilises the transition state of the ATP α-phosphate and a second metal bridges the β-and γ-phosphates to help orienting the pyrophosphate leaving group. Interestingly, in vitro assays show ligation activity without addition of Mn²⁺ ions. In addition, the available LIG·AMPcPP (adenosine-5′-[(α,β)-methyleno]triphosphate) structure was crystallised with Mg²⁺ in the active site (Peschek and Walter 2019). Two possible explanations are that Mg²⁺ and Mn²⁺ can substitute for each other or that trace amounts of Mn²⁺ co-purify from heterologous expression. Two conserved arginine residues (C. thermophilum: Arg334 and Arg337) within an α-helix (a11) of the LIG-C subdomain coordinate a single well-ordered sulfate ion via salt bridges near the active site pocket. In addition, His227 is within distance to form a third salt bridge, and Asn150 forms a hydrogen bond with the sulfate ion (Figure 3A). Notably, Arg337 and His227 were identified as essential residues by alanine scanning mutagenesis (Wang and Shuman 2005). This sulfate ion might represent a surrogate for one of the phosphate groups of Trl1’s RNA substrate.

Trl1-LIG differs from other ATP-dependent polynucleotide ligases by its dependence on a 2′-PO₄ at the splice site of the 5′ exon. In order to ligate the 3′-OH and 5′-PO₄ ends, it requires a terminal 2′-PO₄, which is left over at the splice junction after ligation. Structural and biochemical studies of Trl1-RNA complexes will help us to understand this unique enzymatic property. The remaining 2′-PO₄ at the splice junction is removed by the 2′ phosphodiesterase enzyme Tpt1 (Banerjee et al. 2019c; Culver et al. 1997; McCraith and Phizicky 1991, 1990; Spinelli et al. 1997). Currently, there are no structures of fungal Tpt1. An apo-enzyme structure was published from the archaeum Aeropyrum pernix (Kato-Murayama et al. 2005). Further mechanistic insights into the NAD⁺-dependent phospho-ADP-ribosylation reaction stem from Tpt1 structures from the bacterium Acetivibrio thermocellus (Banerjee et al., 2019c).

Taken together, the available crystal structures of fungal tRNA splicing enzymes have expanded our knowledge about their mechanisms and interplay. However, we still do not know how the three Trl1 modules act together in passing along the substrate or how the exon ends are coordinated during the final ligation step. All three Trl1 modules are essential for cell viability, but in vivo S. cerevisiae experiments revealed that they individually fulfil their functions, when expressed separately from each other (Sawaya et al. 2003).

The human strategy

While all subunits of the splicing endonuclease are homologous proteins in both, yeast and human, the respective tRNA ligases are completely unrelated. Metazoa use RTCB-type ligases to seal tRNA exon halves (Popow et al. 2012, 2011). The phylogenetic origin of RTCB-type RNA ligases is intriguing. While RTCB homologs are present in bacteria, archaea and metazoa displaying a high level of conservation, they are completely absent in fungi and plants (Englert 2005; Englert et al. 2011; Popow et al. 2011; Tanaka et al. 2011; Zillmann et al. 1991). In 2011, three laboratories reported the identification of bacterial, archaeal, and mammalian RTCB proteins as novel RNA ligase enzymes capable of sealing 2′,3′-cP and 5′-OH ends (Englert et al. 2011; Popow et al. 2011; Tanaka et al. 2011). Javier Martinez and co-workers discovered the human tRNA ligase enzyme using an elegant biochemical reconstitution approach (Popow et al. 2011). Employing biochemical fractionation of human cell lysates, they isolated RTCB (also termed HSPC117) as the essential tRNA splicing ligase enzyme. They also identified additional protein subunits that form a stable tRNA ligase complex together with RTCB. This work defines the human ligase as a 228 kDa hetero-pentameric complex consisting of the subunits DDX1, CGI-99, ASW, FAM98B and RTCB (Figure 4).

Figure 4:

The human tRNA ligase complex and activity regulators (competitors and cofactors).

Blue: RTCB ligase complex components. Red: competing factors. Green: promoting factors. AlphaFold (Jumper et al., 2021) structures are shown in grey.

Human tRNA ligase catalyses the sealing of the tRNA exon halves via a different mechanism compared to Trl1-type ligases (Figure 3B). Unlike Trl1, which requires the aforementioned end-healing steps, the human tRNA ligase RTCB directly ligates a 2′,3′-cP or a 3′-PO₄ at the 5′ exon to the 5′-OH at the end of the 3′ exon (Englert et al. 2011; Popow et al. 2011; Tanaka et al. 2011). Regarding its co-factors, RTCB depends on GTP and Mn²⁺ for the ligation of the RNA exon halves (Englert et al. 2011; Popow et al. 2011; Tanaka et al. 2011). Numerous structural models of Pyrococcus horikoshii RtcB (Banerjee et al. 2021; Desai et al. 2013; Englert et al. 2012) have contributed to our understanding of the catalytic mechanism and modes of RNA substrate recognition. RTCB-catalysed ligation occurs in three distinct steps: The first step is the guanylylation of the active-site histidine (H404) via GTP hydrolysis (Figure 3B). The archaeal PhRtcB structures show two Mn²⁺ ions in the active site with different coordination. The octahedrally coordinated Mn²⁺ is involved in the guanylylation step and the tetrahedrally coordinated Mn²⁺ takes part in the further ligation reaction (Englert et al. 2012). The phosphate of the GMP-His intermediate is in contact with the octahedrally coordinated Mn²⁺. The successive step is the transfer of GMP to 2′,3′-cP at the 5′ exon half, which leads to an RNA 3′-P-GMP intermediate. The tetrahedrally coordinated Mn²⁺ interacts with the cyclic phosphate and is involved in the last step of the exon-exon sealing. The guanylylated 3′-end is joined to the 5′-OH of the 3′ exon half while GMP is released (Englert et al. 2012). The phosphate that is used in the exon-3′-P-5′-exon junction is the 2′,3′-cP or 3′-PO₄ of the 5′ exon and does not stem from GTP. RTCB does not require any additional end-modifying enzymes. The 2′,3′-cP is hydrolysed into a 2′-OH and a 3′-PO₄, which is ligated directly to the 5′-OH (Englert et al. 2012; Tanaka et al. 2011).

A crystal structure of PhRtcB in complex with a 5′-OH-containing 6-mer DNA oligonucleotide (Banerjee et al. 2021) shows a possible binding mode of the 3′ RNA exon. The position of the 5′ exon is still unknown although the SO₄²⁻ in another PhRtcB structure (Desai et al. 2013) possibly mimics the 2′,3′-cP terminus of the other RNA substrate half (Banerjee et al. 2021; Englert et al. 2012; Maughan and Shuman 2016). The GMP-, Mn²⁺- and nucleotide-interacting amino acids of RTCB are conserved in all domains of life (Nandy et al. 2017). In a recently reported human RTCB structure with GMP, there are two bound Co²⁺ instead of Mn²⁺ ions in the active site (Kroupova et al. 2021). The same study identified several physical interactions between tRNA ligase core subunits using mass spectrometry/crosslinking suggesting that C-terminal regions of DDX1, FAM98B, and CGI-99 mediate the assembly of the core tRNA ligase complex. These results represent a first step towards understanding the complete molecular architecture of the human tRNA ligase complex.

The analysis of RTCB-containing gene clusters in archaea revealed an additional protein, named archease, that enables full activity of the human tRNA ligase complex. Initial co-immunoprecipitation experiments did not detect archease as component of the RTCB complex, indicating weak binding affinity or a transient interaction (Popow et al. 2011). Nevertheless, our current understanding of its function indicates an important role in conferring multiple-turnover capacity to the RTCB ligase (Desai et al. 2015 2014; Popow et al. 2014). The core enzyme RTCB does not require archease or any of the other known subunits for exon-exon ligation (Englert et al. 2011, 2011; Tanaka et al. 2011). The ligation of tRNA halves in an in vitro reconstitution assay remains incomplete in the absence of archease despite an access of available RNA substrate. Only upon addition of archease, the stalled exon-exon ligation reaction continues to completion by virtue of RTCB functioning as multiple-turnover enzyme. Desai et al. showed that PhRtcB and archease from P. horikoshii work together in a very similar manner, allowing for multiple turnovers and accelerating of the reaction rate (Desai et al. 2014). Archease promotes guanylylation of RTCB’s active site histidine, which represents an activated intermediate along the ligase reaction pathway (Desai et al. 2014, 2013; Englert et al. 2012; Popow et al. 2014). However, the detailed mechanism of RTCB activation by archease remains elusive.

An enhanced effect of archease on RTCB-mediated ligation was observed for archease in combination with DDX1. This effect is increased in the presence of hydrolysable ATP but not with the non-hydrolysable AMPPcP. DDX1 requires ATP for its effect on archease (Popow et al. 2014). The human RTCB ligation complex subunit DDX1 is a DEAD-box helicase. DEAD-Box proteins are the largest family of superfamily two helicases (Linder et al. 1989). However, the SPRY domain (from SPla and the RYanodine receptor) of DDX1 distinguishes it from other DEAD-box proteins (Godbout et al. 1994; Kellner and Meinhart 2015). SPRY domains usually function as protein-protein interaction domains (D’Cruz et al. 2013; Ponting 1997; Woo et al. 2006). The role of CGI-99, ASW and FAM98B as well as the molecular architecture of the holo-complex are still unknown. A recent study provides an interaction map of the core subunits by combining co-immunoprecipitation assays and cross-linking mass spectrometry (Kroupova et al. 2021).

The ligation of the pre-tRNA exon halves to a mature tRNA is not the only possible fate for the transcripts. The 3′ end of the 5′ exon half for example can be modified after cleavage and thus render it non-ligatable for RTCB. RTCB recognises 3′-PO₄ or 2′,3′-cP, thus, the modification of the 3′ end can prevent tRNA ligation. Recent studies with human embryonic kidney (HEK) cells revealed a competitor for the 2′,3′-cP of tRNA 5′ exon halves. This competitor is Angel2, which is a mammalian 2′,3′-cyclic phosphatase and hence dephosphorylates the cyclic phosphate, resulting in a 3′-OH (Pinto et al. 2020). Another competitor is the 2′,3′ cyclic nucleotide phosphodiesterase (CNP). CNP performs the same reaction as the essential healing step of yeast Trl1 (Schwer et al. 2007). It hydrolyses the 2′,3′-cP to a 3′-OH, 2′-PO₄ which cannot be ligated by RTCB (Lee et al. 2001; Olafson et al. 1969; Unlu et al. 2018). CNP is not essential in mice, however, it is involved in neurodegeneration and CNP-deficient mice died prematurely (Lappe-Siefke et al. 2003). In contrast, RTCA (also named Rtc1) modification of the 3′ end reopens the possibility of RTCB mediated ligation. It circularises 2′-PO₄ to 2′,3′-cP and is therefore the antagonist of CNP (Das and Shuman 2013; Filipowicz et al. 1983; Unlu et al. 2018).

The human tRNA ligase RTCB does not require any tRNA end modifications after pre-tRNA cleavage by the SEN complex to ligate the tRNA exon halves. However, the polynucleotide kinase HsCLP1, which is associated with the SEN complex, is responsible for the ATP-dependent phosphorylation of the 3′ exon’s 5′ terminus. This function is very curious since the Homo sapiens tRNA ligase does not require a phosphorylated 5′ end at the 3′ exon for the ligation of the exon halves. In fact, RTCB catalysed ligation is diminished when the 5′ end is phosphorylated (Popow et al. 2011; Tanaka et al. 2011). The function of HsCLP1 during tRNA splicing is therefore still a mystery. Interestingly, the yeast homologue ScCLP1 does not show any kinase activity due to a mutation in the active site (Ramirez et al. 2008). Nevertheless, ScCLP1 is essential in S. cerevisiae and a part of the mRNA 3′-end-forming machinery (Haddad et al. 2012; Holbein et al. 2011). Experiments in which HsCLP1 complemented Trl1 kinase-defective mutants showed that the 5′ phosphorylation during tRNA splicing might be performed by HsCLP1 (Ramirez et al. 2008). It is therefore possible that the kinase activity of ScCLP1 in tRNA splicing was replaced during evolution by the kinase domain of Trl1.

Enzymes, such as RTCB, which contain metal ions coordinated by cysteines at their active site are prone to oxidative stress. Hence, there is a protection mechanism in place to secure their functionality in aerobic conditions. Some aerobic living eukaryotes (e.g. metazoa) use RTCB as tRNA ligase (Popow et al. 2011), other (e.g. plants and fungi) have replaced RTCB with another tRNA ligase, Trl1 (Greer et al. 1983). The enzyme that protects RTCB in humans from oxidative inactivation is PYROXD1 (Figure 4). PYROXD1 is an oxidoreductase that has co-evolved with the proteins of the RTCB complex and contains a pyridine nucleotide-disulfide oxidoreductase domain. It catalyses the NAD⁺/NADH-dependent reduction of oxidised cysteine residues, most importantly C122 in the HsRTCB active site (Argyrou and Blanchard 2004; Asanović et al. 2021; O’Grady et al. 2016).

Other cellular roles and clinical relevance of tRNA splicing enzymes

There are several known examples of tRNA splicing components functioning in other cellular pathways. Probably, the most prominent case of another eukaryotic splicing mechanism is the IRE1-mediated stress signalling during the unfolded protein response (UPR). Upon protein folding stress in the endoplasmic reticulum (ER), the cytosolic endonuclease domain of the transmembrane stress sensor IRE1 is activated (Mori et al. 1993; Nikawa 1996) and cleaves HAC1 mRNA in fungi or XBP1 mRNA in metazoans at the splice sites (Calfon et al. 2002; Cox and Walter 1996; Yoshida et al. 2001). The HAC1/XBP1 mRNA exons are ligated by Trl1 (Sidrauski et al. 1996) and RtcB (Jurkin et al. 2014; Kosmaczewski et al. 2014; Lu et al. 2014), respectively. Thus, despite their independent phylogenetic origin, both types of tRNA ligase moonlight in the UPR rendering them essential components of ER homeostasis. The non-conventionally spliced mRNAs are translated to active transcription factors promoting the expression of UPR target genes [The UPR has been reviewed elsewhere (Ron and Walter 2007)].

The dual functionality in tRNA splicing and non-conventional mRNA splicing during the UPR shows the versatility of tRNA ligases. Biochemical characterisation of various tRNA ligases established that they ligate a broad range of “artificial substrates” such as nucleolytic RNA fragments and synthetic RNA if they exhibit suitable functional groups at the termini (Chakravarty et al. 2012; Popow et al. 2011; Tanaka et al. 2011). Thus, it is not surprising that homologous RNA ligases are involved in RNA repair pathways in bacteria (Manwar et al. 2020; Temmel et al. 2016). A well-studied example is the bacteriophage T4 RNA ligase (T4Rnl1), which repairs an endonuclease cut in Escherichia coli tRNAs (Amitsur et al. 1987). T4Rnl1 is a prototypical member of the same enzyme class as fungal Trl1. Intriguingly, there is also an RTCB homologue in E. coli where it functions as tRNA ligase in ribosome repair. E. coli RtcB repairs stress-induced cleavage of 16S ribosomal RNA (rRNA) by the endonuclease MazF to restore translational functionality of bacterial ribosomes (Temmel et al. 2016). Moreover, human RtcB was described as part of the mammalian “ribo-interactome” (i.e. ribosome-associated protein), which might indicate a ribosome-related ligase function (Simsek et al. 2017). The identification of additional tRNA ligase substrates and protein interaction partners harbours great promise for fascinating insights into RNA biology.

Because of its essential functionality for cell viability, the tRNA splicing machinery is interesting from a clinical perspective. tRNA processing was found as a basic cellular impairment in neurological disorders. Mutations in tRNA splicing endonuclease cause pontocerebellar hypoplasia (PCH), a neurodegenerative autosomal recessive disorder (Budde et al. 2008). A recent study has shown that PCH-associated mutants of human TSEN cause reduced complex stability and accumulation of pre-tRNA processing intermediates (Sekulovski et al. 2021). The cellular role of the interaction between TSEN and the RNA kinase CLP1 and its relevance for human disease remains to be elucidated. Previous studies already pointed out a correlation of a homozygous mutation in CLP1 to tRNA maturation and neuronal development resulting in neurodegeneration through defective CLP1 function in humans (Hanada et al. 2013; Schaffer et al. 2014). [The roles of CLP1 and the tRNA splicing machinery in other RNA pathways and their link to neurodegenerative disorders are reviewed elsewhere (Weitzer et al. 2015).]

The fungal tRNA splicing enzymes Trl1 and Tpt1 are present and essential in all fungi and thus also in all clinically relevant fungal pathogens. Because their structure and mechanism are distinct from the human RTCB-type tRNA ligase in metazoans (as well as archaea and bacteria), they are often discussed as promising targets for antifungal therapy. The development of new fungal drugs has stalled in recent years, which is especially problematic due to the emergence of drug resistances. One major challenge is the identification of novel targets due to the high degree of conservation between fungal enzymes and their human equivalents (Denning and Bromley 2015). There is no known human homolog of Trl1, which renders it particularly suited for antifungal intervention. The cellular function of TRPT1, the human homologue of Tpt1, has remained unknown. While Tpt1 is an essential enzyme in yeast, TRPT1 knock-out mice are viable and display no measurable effect on tRNA splicing (Harding et al. 2007).

Taken together, there is an emerging appreciation of the existence of RNA repair pathways that rely on RNA ligases to maintain RNA structure and integrity. Eukaryotic tRNA ligases are essential RNA processing enzymes and represent in their respective host species the only (currently) known ssRNA ligase enzyme (Phizicky et al. 1992; Popow et al. 2011). In combination with the possibility of targeting the fungal enzymes for antifungal therapy lies the exciting potential for tRNA ligases as physiologically versatile and clinically relevant RNA repair enzymes.

Future perspectives and open questions

The evolutionary origin as well as the biological function of tRNA introns has remained elusive. Introns preclude tRNAs from their essential task during translation. Why do organisms possess introns in some of their tRNA genes? What is their function? How does tRNA splicing intersect with other tRNA maturation steps? Several studies indicate specific effects of tRNA introns on certain modifications (Grosjean et al. 1997; Yoshihisa 2014). However, considering the large number, chemical variety, and importance of nucleotide modifications within tRNAs, little is known about their interplay with processing pathways, including tRNA splicing.

The existence of competitors for the cleaved tRNA substrate such as ANGEL2 and CLP1 shows that there is more to discover. Why are the exon ends modified in a manner that interferes with ligation by tRNA ligase? Incorrect processing of intron-containing tRNAs as well as the accumulation of intermediate tRNA fragments, are specifically linked to human neurological disorders (Budde et al. 2008; Karaca et al. 2014; Schaffer et al. 2014). Thus, a better understanding of the role of tRNA introns and splicing might harbour medical relevance.

While we have a good grasp on the “division of labour” between SEN and tRNA ligase during tRNA splicing, we know surprisingly little about their interplay in the cell. How is the tRNA splicing reaction spatially organised? Do endonuclease and ligase interact (either directly or mediated by an adaptor protein)? Is there a hand-off mechanism of the cleaved pre-tRNA to shield it from exonucleolytic degradation? Is there a specific targeting mechanism for intron-containing pre-tRNAs towards the enzymatic centres?

The existence of two unrelated tRNA ligase enzymes within the main eukaryotic groups is peculiar. Despite the mechanistic differences between Trl1-type and RTCB-type ligases, many of the open questions are similar. To date it is not known how ligase enzymes coordinate the two RNA exons for the final sealing step. To this end, we will probably need structures of RNA substrate complexes. Other questions regarding tRNA ligases remain: Is there any coordination between the three Trl1 modules? What are the functions of the different components of the RTCB ligation? What are their respective roles outside of tRNA splicing?

Recent studies have uncovered fascinating insights into the structure and function of the eukaryotic tRNA splicing machinery. Several additional components and interactions have been identified over the past few years. Now, we have available atomic-resolution structures of tRNA splicing enzymes, albeit mostly of single domains or subunits. It will be interesting to see how our structural understanding of the eukaryotic tRNA splicing machinery will expand in the wake of modern structural biology methods and breakthroughs in protein structure prediction.

Corresponding author: Jirka Peschek, Biochemistry Center (BZH), Heidelberg University, D-69120 Heidelberg, Germany, E-mail: jirka.peschek@bzh.uni-heidelberg.de

Janina L. Gerber and Sandra Köhler contributed equally to this article.

Funding source: Deutsche Forschungsgemeinschaft

Award Identifier / Grant number: Emmy Noether/442512666

Award Identifier / Grant number: TRR 319/439669440

Acknowledgments

J.P. acknowledges funding from the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) via an Emmy Noether grant and as part of the Collaborative Research Centre TRR 319 TP A06.

Author contributions: All the authors have accepted responsibility for the entire content of this submitted manuscript and approved submission.
Research funding: None declared.
Conflict of interest statement: The authors declare no conflicts of interest regarding this article.

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Received: 2021-10-25

Accepted: 2022-05-10

Published Online: 2022-05-30

Published in Print: 2022-07-26

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enzyme catalysis; ligase; nuclease; protein structure; RNA processing

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