The first reported method for the removal of total tRNA from S30 extracts is based on ethanolamine agarose (Figure 3A). It was discovered by accident that 90% of native tRNA in rabbit reticulocyte lysates could be separated by covalent interactions using the chemical groups of ethanolamine anchored to the resin. (Jackson et al., 2001). For this method, a column of epoxy-activated Sepharose^® 6B is more sensitive than ω-aminododecyl-Sepharose. This lysate system depends on supplementation with tRNA and is only applicable for the synthesis of smaller and medium-size proteins, but not for large proteins. The ethanolamine-Sepharose column procedure can expand opportunities for manipulating the tRNA population for reorganizing the genetic codon. However, this method is not universally applicable for all extracts. The most critical step is to optimize the concentration of different salts in the reaction, which influence the interaction between the tRNAs and the ethanolamine-Sepharose resin. Furthermore, there are still several challenges redesigning sense codon for ncAAs owing to the residual total tRNAs. Subsequently, the buffer used to equilibrate the resin was substituted with pure water, resulting in the elimination of about 95% of the total endogenous tRNAs in E. coli extracts (Ahn et al., 2006). Although this treatment process is simple and the removal efficiency is improved, a small amount of tRNA is still present, which increases the risk of residual tRNA coupling with amino acids. To alleviate the negative effects, reaction conditions including salts, ionic strength, temperature and retention time could be optimized to increase the interaction between tRNA and ethanolamine agarose so as to facilitate the construction of a tRNA complement protein synthesis system.

The reassignment for sense codons requires the complete removal of native tRNAs from the cell extract. A promising emerging approach is based on RNase-coated magnetic beads and a phenylmethylsulfonyl fluoride (PMSF)-treated cell extract, which results in near-complete tRNA depletion (Salehi et al., 2017). For this approach (Figure 3A), ribonuclease A (RNase A) attached to superparamagnetic beads was more convenient than an immobilized RNase A Resin in controlling the RNase degradation ability and subsequently removing the RNase from the extract (Kunda et al., 2000). In addition, this method can make full use of the protective effect of nucleoproteins, so that it not only degrades tRNA but also ensures the activity of rRNA (Suhasini and Sirdeshmukh, 2006). PMSF treatment has a two-sided effect, since it inhibits proteases fivefold less than using RNase inhibitor, while also preventing RNaseA from leaching into the cell extract. Notably, the rate of tRNA removal was directly measured using quantitative real-time PCR (qPCR) with tRNA-specific primers (Kralik and Ricchi, 2017). The average removal ratio for all assessed native tRNAs was 99.3% following RNase A treatment for 60 min. However, there are still some problems that need to be addressed. The cell extract treated with RNase A-beads was able to produce a designer peptide containing 40 Val residues with the addition of a synthetic tRNA, but it remains unclear if proteins with larger molecular weights can be synthesized. The removal of tRNA by the magnetic bead method is only applicable for the cell extract of E. coli, and must be further optimized for other bacteria or eukaryotes. In general, this method is more practical than most current methods for tRNA removal. The direct quantification by PCR is a valuable tool to assess cell extracts for high-fidelity codon reassignment. Looking forward, this robust platform has the potential to be applied in emerging synthetic biology and biomedical engineering applications.

For specific tRNA removal, the primary issue is codon choice, while excluding Trp and Met with only one encoding codon. According to previous studies, the tRNase colicin D (ColD) can specifically recognize and degrade four different tRNA^Arg species of E. coli, including the tRNA_ICG, tRNA_CCG, tRNA_UCU, and tRNA_CCU (Tomita et al., 2000). This method (Figure 3B) takes full advantage of this property to inactivate all the tRNA^Arg from the E. coli cell extract (S12) (Kim et al., 2006), resulting in a inability to incorporate Arg in this system. It was further demonstrated that ColD-treated lysates did not affect the ability of protein synthesis, under the condition of supplementing tRNA^Arg. More importantly, this creates favorable conditions for rearrangements of the remaining Arg codons to expand the genetic codon (Lee et al., 2016). There are two reasons for choosing AGG as the sole codon encoding Arg. Firstly, this is a rare codon, while tRNA^UCU and tRNA^CCU are relatively resistant to ColD digestion, resulting in a residual amount remaining in the treated S12 lysates. For specific sense codon rearrangement, it is also important to take into account the flexible effect of wobble base pairs in the in vivo decoding process (Das and Duncan Lyngdoh, 2014). Although this method is simple and feasible in reconstructing translation systems, it can only be used to rearrange Arg codons and not for other types of tRNAs. Therefore, there are more other tRNases to be explored for use in similar approaches, such as the tRNA^Lys anticodon nuclease PrrC (Blanga-Kanfi, 2006). Overall, this method expands further implementations for building a platform for a small number of sense codon rearrangements, and it creatively broadens new horizons and ideas for reconstructing a more general platform.

Emerging attempts have been made to free up more sense codons for the incorporation of multiple ncAAs in the genetic codon simultaneously. Alexandrov et al. demonstrated that isoacceptor tRNAs can be removed through DNA-hybridization chromatography of the standard E. coli S30 lysate (Cui et al., 2017). For this approach (Figure 3B), DNA oligos with 3′-amine modifications were designed complementary to the sequence spanning the D-arm down to the anticodon loop of the targeted native isoacceptor tRNA (tRNA^Arg _UCU), and were immobilized on an NHS-activated matrix. Then, the treated DNA oligos are mixed with the extract to carry out the hybridization reaction for the chromatographic depletion. The authors introduced a novel criterion for tRNA removal efficiency as follows: tRNA   removal   efficiency = 1 − ( RFU ( Depleted tRNA ) RFU ( Depleted tRNA + t7 tRNA ) )

According to this formula, there are many factors contributing to the more precisely quantified tRNA removal. First of all, the feasibility of this method was demonstrated since the protein expression level could be restored with the addition of the targeted synthetic tRNA in the tRNA-depleted lysate. The depletion efficiency is also related to the number of corresponding decoded codons contained in the template of protein synthesis. For example, using super-folder green fluorescent protein (sfGFP) as the template, the deletion rates of one and six AGG codons were nearly 100 and 90%, respectively. Therefore, the depletion efficiency is determined depending on codon composition in different templates, which further illustrates the role of a specific tRNA and the ability of protein synthesis. Early studies identified a biased distribution of tRNA abundance for bacteria growing at different rates (Dong et al., 1996). This could directly affect the quality of the prepared cell lysate. To ensure basic levels of protein synthesis, several new technical means can be used to accurately measure the amount of specific tRNAs to determine the optimized time for collection of bacteria, such as the modification-induced misincorporation tRNA sequencing (mim-tRNAseq) method (Behrens et al., 2021).

Subsequently, several implemented effective measures were taken to abrogate the targeted tRNA activity and liberate the corresponding sense codons (Cui et al., 2018). This strategy offers several improvements over the above-mentioned methods. Firstly, the antisense oligonucleotides are not affected by the high-level complex molecular structures in tRNAs that reform the L-shaped conformation. Secondly, 2′O-Me modifications of the antisense oligonucleotide perform better than other types in the entropic stabilization to the hybrid base pairs (Yildirim et al., 2014). It should be noted that methylated oligonucleotides display slow dissociation kinetics from the target tRNA, resulting in better RNA strand displacement. This is particularly important for the designed sequence targeting a specific tRNA in the E. coli S30 lysate. The hybridization ability shows inconsistencies in the sequence of the same tRNAs between species (E. coli and Leishmania tarentolae), even though they are located in the anticodon or variable loop region in the same cell. A few heterologous factors need to be further explored to improve the general applicability of the method, such as the input ratio of tRNA and oligonucleotides, incubation time, and temperature, which requires further work in the future. In addition, the two-step protein synthesis reaction system was divided into the treated extract and the total RNA. The described standard L. tarentolae extract (LTE) (Mureev et al., 2009) was generated by removing the total RNA by the ethanolamine-sepharose column method. The total RNA was isolated from the lysates. Then, individual isoacceptor tRNAs were selectively sequestered from the purified total RNA by the treatment with DNA oligos with 2′O-Me modifications. Finally, the above two parts are added to the whole reaction system to minimize the background level. Although a somewhat cumbersome, this method is worth learning and considering to reduce the background activity. Treatment with the RNase-related compound angiogenin (ANG) (Su et al., 2019), which can cleave tRNA anticodons to inactivate specific codons, may be a feasible strategy for freeing up sense codons. More effective strategies for the tRNA denaturation approach are expected to be developed in the near future.

In protein translation systems, the requirement for purifying individual tRNAs in vivo has become increasingly prominent. A prominent approach relies on the introduction of a hydrophobic tag for the specifically charged aminoacyl-tRNAs (aa-tRNA) followed by hydrophobic interaction chromatography (Figure 4A) (Kothe et al., 2006). The used hydrophobic tag was 9-fluorenylmethyl-succinimidyl carbonate (FmocOSu), which can react with the free amino group of a selected aa-tRNA. Therefore, the hydrophobicity of this complex is significantly increased compared with other tRNAs, which in turn increases the retention time in the chromatographic column, and finally results in its isolation from total tRNA. Although the used raw material is available at a low cost, and the operation is relatively simple, the entire purification process requires multiple steps, including aminoacylation, modification, purification, and deacylation. In addition, this hydrophobic tagging method has broader applicability for diverse tRNAs than previous approaches (Cayama et al., 2000), because it solely depends on the specific aa-tRNA synthetases and the specific amino acids. The aaRS can only aminoacylate tRNA with one specific amino acid, but it cannot recognize and distinguish the isoacceptor tRNAs. According to this principle, the purified species are a complex of tRNAs encoding the same amino acid. This method is therefore not suitable for the fine-tuning of transfer RNA (tRNAs) in the cell-free lysate. Recent research found that a two-dimensional liquid chromatography (2D-LC) integrating a weak anion-exchange method could be used to isolate tRNA^Val _UAC and tRNA^Leu _CAG (Cao et al., 2020). These purified individual tRNAs might be applied to the minimal codon protein synthesis system. In terms of the cost and feasibility of purification, these methods will open new avenues in the process of simplified chromatographic purification.

In an early study on probe-elution, biotinylated DNA oligonucleotides were immobilized onto streptavidin agarose beads to isolate individual tRNAs from the Leishmania tarentolae (Kaneko et al., 2003). Recently, Söll et al. incubated 5′biotinylated oligo-beads with yeast tRNAs in the equilibrated buffer, and this complex was washed several times to remove the nonspecifically bound or loosely bound molecules. The pure tRNA_m ^Glu was eluted by a low-salt buffer at high temperature (Rinehart et al., 2005). Although the principle and operation of this procedure are relatively simple, verification of the tRNAs from a cell by northern blot may be necessary (Figure 4B). However, the current purified volume and purity are relatively low for the distribution and regulation of different species of tRNA for in vitro protein synthesis, which hinders its application and development. The following different factors are worth exploring to produce high-quality tRNA. First of all, the individual tRNAs could be over-expressed by constructing recombinant plasmids with a strong constitutive E. coli promoter (Cervettini et al., 2020). The main purpose of this process is to enrich enough tRNA for initial extraction and purification, greatly reducing the proportion of non-target tRNAs. In addition, the combination of a chromatographic spin column and streptavidin-coated agarose beads may have more obvious advantages in improving the binding and purification efficiency for bulk purification. The continuous circulation process in the Chaplet Column Chromatography (CCC) method circumvents these drawbacks to a certain extent (Suzuki and Suzuki, 2007). Similarly, the choice of synthetic 3′-biotinylated DNA probes and chromatographic columns can be changed according to the amount of the Bos taurus mitochondrial tRNA mixture, exhibiting great flexibility and scalability.

With easy availability, biotinylated DNA oligonucleotide probes complementary to the targeted tRNA were successfully used to extract the tRNA-derived stress-induced RNAs (tiRNAs) fraction by the gel purification process (Nilsen, 2013). The hybridized tiRNA-oligo complex was pulled down using streptavidin agarose beads (Figure 4C). With the addition of the Trizol reagent, the targeted tRNA fraction can be freed via digestion with DNase I (Akiyama et al., 2020). While initially met with some skepticism, this approach was developed into feasible methods for the isolation of endogenous mature tRNAs, such as tRNA^Ala _AGC and tRNA^Gly _GCC. Since tRNAs have a complex molecular structure and are heavily modified, this approach conquered the current limitation of tRNA purification from host cells and opened up a new Frontier in tRNA biology. Nevertheless, several disadvantages of this approach should be addressed. To obtain high-purity target tRNA, toxic Trizol was used repeatedly throughout the purification process, which should be circumvented by seeking alternative reagents. The designed sequences of DNA probes need to be optimized due to the high sequence similarity between the selected tRNA and its isoacceptor tRNAs. Finally, there is a lack of scientific evaluation criteria for the purity and activity of purified tRNA. Emerging technologies, including high-throughput tRNA sequencing (Shigematsu et al., 2017; Erber et al., 2020) and immuno-northern blotting (Mishima et al., 2015; Mishima and Abe, 2019), may be applied to push forward tRNA purification technology.

Among bacteriophages T3, T7, and SP6, T7 RNA polymerase (T7RNP) offers the largest yield of RNA transcription. Since the length of tRNA is between 75 and 95 nt, the T7RNP transcriptional synthesis of small RNA has been established in initial reports (Milligan and Uhlenbeck, 1989). Originally, there were two major flaws in the T7 transcription process. One defect is that its transcription efficiency mainly depends on the specific recognition of its cognate promoter sequence, incorporating an unfavorable sequence at the 5′-end. On the other hand, the runoff of T7 transcription results in a mixture of products with 3′-end heterogeneity, which usually results in multiple consecutive A nucleotides at the end of the product (Helm et al., 1999). Three different transcription patterns for tRNA transcription will also be discussed based on their applications in the field of transcriptional product heterogeneity (Figure 5A). In a direct pattern, the transcription template starts with the T7 promoter sequence, followed by the encoded tRNA gene sequence, which is directly transcribed (Li et al., 1999). This approach works for most mature tRNAs within cells, but it is limited to tRNA sequences that do not begin with a G, such as tRNA^Asn and tRNA^Pro. The ribozyme pattern is a fusion between the T7 promoter sequence and the tRNA gene, encoding a ribozyme called hammerhead, which has a cis-acting, self-cleavage ability (Huang et al., 2019). This design intends to circumvent the above-mentioned deficits and release a tRNA transcript with the desired 5′- sequence (Korencic et al., 2002). The main advantage of this approach is that pure transcripts can be obtained directly in large quantities for all tRNAs. Although the 5′-OH tRNA transcript can be aminoacylated in this reaction, they proved to be active. Additionally, RNase P can catalyze tRNA maturation with the generation of tRNAs with homogeneous 3′and 5′ends (Fukunaga et al., 2006). RNase P has a catalytic RNA subunit, which can cleave homogeneous 3′-OH ends in tRNAs of interest (Gossringer et al., 2012). The 2′-methoxy modification of the second nucleotide 5′end at the primer prevents additional nucleotide amplification of the 3′-terminal transcripts. Therefore, the latest research fully integrated the above two characteristics, to achieve the production of transcripts of high quality (Hibi et al., 2020). Using this approach, 21 different tRNAs from E. coli were analyzed by urea-PAGE, and the homogeneity of the major synthesis was successfully verified. What’s more, the purified iVTtRNA can be sufficiently aminoacylated by the corresponding aaRS. This breakthrough demonstrates that despite lacking any modifications in vivo, tRNA transcribed using RNase P was capable of protein synthesis comparable to the native tRNA mixture. The in vitro transcribed transfer RNA (iVTtRNA) lacks the modifications at position 37, which possibly allow undesirable wobbling at codon first positions and frameshifts. Indeed, generating the complex base modifications in the anticodon loop regions of iVTtRNA will be an important perspective in decoding accuracy required for the appropriate modification enzymes like the TsaD and TrmD modification enzymes. The introduction of modified nucleotides into the anticodon loop of iVTtRNAs shows a significant increase in the aminoacylation, the protein yield, and the fidelity. Overall, this is a fast, inexpensive and efficient method for the production of specific tRNAs of good purity, providing favorable conditions for in vitro protein synthesis based on the iVTtRNA system.

In early studies, solid-phase chemical synthesis was used for the production of 77 nt tRNA^Met (Ogilvie et al., 1988). The whole production cycle involves de-blocking, coupling, capping, and oxidation (Figure 5B). According to the sequence of chemical synthesis from the 3′- to the 5′-terminus, the synthetic target tRNA is finally obtained through reversed-phase high-performance liquid chromatography (HPLC) purification (Baronti et al., 2018). From the cost perspective, the upper limit of the synthesizable length is approximately 80 nt. In this approach, the synthesis mainly depends on the used phosphoramidite monomers and the spatial structure of target sequences. Accordingly, synthetic production is a challenge for all tRNAs, which are generally structurally complex molecules. However, it is still feasible and effective to adopt chemical methods for the synthesis of tRNAs with specifically modified bases (Vare et al., 2017). Chemical synthesis has been used for decades in the production of small RNAs, and it still plays a major role. For future studies, there is an urgent need to develop a lower-cost, high-efficiency protocol, focusing on the manipulation of tRNAs.