Structural Elements of DNA and RNA Eukaryotic Expression Vectors for In Vitro and In Vivo Genome Editor Delivery

Abstract

Gene editing with programmable nucleases opens new perspectives in important practice areas, such as healthcare and agriculture. The most challenging problem for the safe and effective therapeutic use of gene editing technologies is the proper delivery and expression of gene editors in cells and tissues of different organisms. Virus-based and nonviral systems can be used for the successful delivery of gene editors. Here we have reviewed structural elements of nonviral DNA- and RNA-based expression vectors for gene editing and delivery methods in vitro and in vivo.

INTRODUCTION

Currently there are three main variants of genetic editors, i.e., ZFN, TALEN, and CRISPR/Cas systems. The use of these systems involves the creation of vectors for the expression of the above proteins. Currently, viral and nonviral expression vectors are widely used. Nonviral vectors include DNA-based vectors and synthetic mRNAs. In general, the structure of DNA-based vectors for genome editing differs little from the expression vectors of other therapeutic recombinant proteins. In both, there are structural elements, such as promoters, enhancers, poly(A) sequences, selective markers, replication initiation sites, etc. Differences in structure relate to specific elements necessary for the functioning of genome editors, and each editing system has its characteristics. In most cases, a nuclear localization signal is merged with the sequence of the genome editor, and the CRISPR-Cas-carrying vectors contain promoters for RNA polymerases II and III because it is necessary to obtain guide RNA in addition to proteins.

RNA molecules can also be used as expression vectors for the synthesis of therapeutic proteins and genetic editing of cellular DNA. This delivery method has several significant advantages over DNA vectors. When using RNA, the probability of integration of the vector or its parts into the genome is extremely small, which almost totally excludes mutagenic events. Another significant advantage is that RNA in combination with lipid nanoparticles is a biodegradable carrier. In addition, there is no need for RNA to penetrate the cell nucleus because the translation of the transgene occurs immediately after entering the cytoplasm. Finally, the transient nature of transgene expression with RNA is better controlled, which avoids excessive production of transgenes, such as gene editor molecules, and reduces the risk of non-specific editing of the cell genome. It is necessary to note the main disadvantages of using artificial RNA carriers, which are primarily caused by the instability of RNA and the high immunogenicity of foreign RNA, which is associated with the mechanisms of antiviral immunity formed by the evolution of eukaryotes.

The potential for biomedical use of synthetic mRNAs was demonstrated in 1990 in an experiment on the expression of various proteins in skeletal muscle cells of mice. However, problems with extracellular and intracellular stability, RNA immunogenicity, and the complexity of large-scale production have slowed down the spread of the use of synthetic mRNAs for therapeutic purposes. Currently, the synthesis of single-stranded RNA molecules in vitro is a widespread laboratory procedure, which is actively used for both the study of RNA and the preparation of RNA-based drugs. This method is applicable for the biochemical and molecular analysis of RNA, the study of RNA-protein interactions, the structural analysis of complexes, the creation of RNA aptamers, the synthesis of functional mRNAs for expression, and the production of small RNAs that affect gene expression (e.g., guide RNAs for gene editors). In addition, in vitro synthesized RNAs have played an important role in the development of RNA vaccines, the CRISPR/Cas9, ZFN, and TALEN genome editing tools, pluripotent stem cells, and diagnostic methods based on RNA amplification.

In this review, we will consider the main structural elements of DNA- and RNA-based expression vectors including those used in gene editing and the methods for in vitro and in vivo delivery of these vectors.

STRUCTURAL ELEMENTS OF DNA- AND RNA-BASED EXPRESSION VECTORS FOR DELIVERY OF GENOME EDITORS

Promoters

A promoter is a nucleotide sequence that is recognized by an RNA polymerase complex and specifically binds this complex, thus leading to the initiation of transcription of the nucleotide sequence located below the promoter. Mammalian cells contain several types of RNA polymerases, each of which interacts with specific promoters. Most often, researchers are interested in promoters that can interact with RNA polymerase II. This enzyme is responsible for the synthesis of mRNAs, which are templates for the synthesis of polypeptides [1]. The functioning of any promoter is due to the interaction of proteins that are part of RNA polymerase with certain nucleotide motifs in the promoter. The structure of mammalian promoters, which interact with RNA polymerase II, contains about ten different motifs, i.e., MTE, BREu, BREd, XCPE1 and others, and the most well-known motifs are the initiator and TATA box [2]. Most motifs have a fixed position on the promoter relative to the transcription start point. However, CpG islands, for example, can be located at different positions.

Both the position of motifs and their composition are important for the structure of promoters. Based on these two parameters, promoters are divided into two types: scattered and focused promoters. The scattered type most often includes promoters that ensure the expression of housekeeping genes. They often lack an initiator and the TATA box but contain a large number of CpG islands. These promoters provide stable moderate expression of the gene under control. As an example, we can consider the promoter of the human phosphoglycerate kinase (hPGK) gene, which contains a large number of CpG islands but lacks the TATA box and the initiator sequence. In contrast, focused promoters often have few CpG islands but there are motifs, for example, an initiator and a TATA box, which provide strong interaction with RNA polymerase complexes [3]. These promoters usually provide intracellular expression at a high level in a short time. Promoters of this type are often found in the genomes of viruses. The promoters of the focused type include the promoter of the SV40 virus or the promoter of the human EF1a gene. Since researchers want to achieve a high level of expression of recombinant proteins when creating vectors, they use promoters of the focused type.

In practice, it turns out that many promoters of this type have a viral nature, for example, the SV40 or cytomegalovirus (CMV) promoters. Sometimes, there is a need for an RNA product whose structure differs from that of mRNA. In particular, this product is necessary for obtaining a guide RNA for the CRISPR/Cas genome editing system. For this purpose, a promoter that interacts with RNA polymerase III is used, e.g., the U6 promoter [4]. In addition to promoters that provide stable expression, there are also induced promoters, which can be both scattered and focused types and include a regulatory element to control expression [5]. Examples are the Tet-On inducible system or the thermoinducible Hsp70 promoter. These promoters can be advantageous when the synthesized protein is cytotoxic or it is necessary to give time to the cell culture to grow before expressing the target gene.

Another type of promoter, which have much in common with inducible promoters, are tissue-specific promoters. They can belong to both the scattered and focused types but contain a motif or motifs, which specifically interact with some protein. In most cases, the protein is a tissue-specific transcription factor. Since gene expression under the control of these promoters occurs strictly in a certain type of tissue, they are used for genetic editing in vivo [6]. For example, the SLPI promoter, which is active in some types of carcinomas, ensures a low level of expression of the gene encoding the inhibitor of leukoprotease secreted in the liver. Promoters come in different ‘strengths’, i.e., they provide different levels of gene expression. They can promote both stable gene expression and expression in a certain type of tissue under the action of a certain inductor. They can have different sizes ranging from hundreds to thousands of nucleotides. It is necessary to select these parameters based on the needs of the experiment.

Enhancers

Enhancers increase the level of gene expression by increasing the local concentration of transcription factors. Like promoters, enhancers perform their function using a variety of motifs that attract transcription factors [7]. Enhancers are most often located in the genome far from the controlled promoter; the promoter and enhancer may even be located on different chromosomes [8]. At the same time, due to the DNA architecture, the enhancers are located near the controlled promoter that is a prerequisite for enhancer function. In vectors, however, enhancers are located before the promoter. Eukaryotic enhancers most often have a fairly large size; therefore, viral variants of these elements with a small size have found wide application. For example, the CMV enhancer [9] is widely used, which is merged with both viral and eukaryotic promoters. Most of the currently used vectors contain an enhancer for increasing the level of expression of the target gene.

Untranslated Sequences

Translation of the polypeptide begins with the AUG initiator codon (start codon). For successful initiation, the start codon must be in a certain nucleotide surrounding, i.e., the Kozak sequence. This motif is quite conservative; in higher eukaryotes, it is represented by the GCC(A/G)CCATGG sequence, which contains especially important nucleotides at positions –3 and +4 [10]. The Kozak sequence functions through interaction with the 40S subunit of the ribosome and translation initiation factors. During mRNA scanning, the ribosome is delayed on the secondary structure formed by the Kozak sequence, thus increasing the probability of translation initiation [11]. When creating a vector, the Kozak sequence is embedded directly before the gene because the absence of this sequence significantly reduces the translation efficiency. The poly(A) tail on the 3' end of mRNA significantly increases both the stability of mRNA and the efficiency of translation. To attach the poly(A) tail in the mRNA structure, the plasmid should include the polyadenylation signal, i.e., the conservative sequence with the AATAAA motif followed by a GT-rich region. mRNA without the poly(A) tail undergoes rapid degradation by cellular nucleases, which negatively affects the overall efficiency of transgene expression [12]. Therefore, the inclusion of the polyadenylation signal in the expression cassette after each protein-coding sequence is also a mandatory step in creating a vector that expresses a recombinant protein.

Selective Marker

A selective marker helps to successfully select transformed cells. Often, antibiotic resistance genes are used because this allows one to quickly select transfected cells by their survival after adding the appropriate antibiotic to the nutrient medium. Moreover, the presence of the antibiotic resistance gene makes it possible to be sure that the plasmid will not be eliminated by the cell over time because the cell needs to express the antibiotic resistance gene for survival under constant antibiotic pressure. To save space, the selective marker can be expressed along with the transgene in the same reading frame using the 2A peptide or IRES element [13]. However, it should be taken into account that the expression level under the control of the IRES element is significantly lower than under the control of the promoter. In addition to resistance genes, fluorescent proteins, such as GFP, are also used as a marker of selection in eukaryotic cells. In this case, transformants are selected using cellular sorters. This approach is appropriate when it is necessary to temporarily express a transgene, followed by elimination (preferably) of the plasmid from the cell to prevent load on the translational apparatus [14]. In particular, this approach is used when creating plasmids that carry gene editors. The fluorescence proteins make it possible to select transformants; genetic editors work for a short time, followed by degradation of the plasmid. There is also a ‘bacterial’ part in the plasmids, which is necessary to obtain a large amount of the plasmid before transfection of eukaryotic cells. A prokaryotic selective marker is present in the bacterial part; most often it is the ampicillin resistance gene. This gene allows Escherichia coli cells to be cultured on an ampicillin-containing medium and, more importantly, to retain the plasmid in the cell.

Signal Peptide

It is often not enough to express the target gene; it is also necessary to ensure the secretion of a protein from the cell or its entry into the appropriate compartment for functioning. For this purpose, there are signal peptides encoded by the 5'-terminal region of the nucleotide transgene sequence. As a result, the synthesized protein is delivered to the cellular compartment, determined by the signal peptide. Genetic editors are usually merged with nuclear localization signals, which leads to the delivery of the synthesized editor to the cell nucleus, where it exhibits its activity [15].

Replication Start Point

Another important component of the bacterial part of the vector is the replication start point (origin) of replication. This is the start point of the synthesis of plasmid copies in a bacterial cell. In fact, this is a regulatory site that influences the number of copies of the plasmid in the cell. There are low-copy and high-copy replication origins. Researchers most often need to obtain a large number of copies of the plasmid; therefore, the high-copy ColE1 origin of replication has become widespread [16].

DNA-Based Expression Vectors to Deliver Genome Editors

Summarizing the above, the map of the base vector for the expression of recombinant therapeutic proteins can be schematically represented as follows. The base vector must have a bacterial part, which should consist of a replication origin to obtain a large number of copies of the plasmid in E. coli and a prokaryotic selective marker of the vector resistance in E. coli cells. The ‘eukaryotic’ part of the vector should contain a fused enhancer and promoter to control the expression of a certain target protein, the Kozak sequence at the 5' end of the sequence encoding the protein, and the polyadenylation signal with a terminator on the 3' end. If the vector is to be used in vitro, it is necessary to have a eukaryotic selective marker (such as an antibiotic resistance gene) controlled by a second promoter because this marker is also a protein. The Kozak and poly(A) sequences are also needed (Fig. 1).

Fig. 1.

The structure of the base vector for eukaryotic expression of therapeutic proteins. (1) Replication origin; (2) prokaryotic selective marker; (3) prokaryotic promoter; (4) eukaryotic enhancer; (5 and 9) the eukaryotic promoters specifically interacting with RNA polymerase II; (6 and 10) Kozak sequences; (7) target protein gene; (8 and 12) polyadenylation signals; (11) eukaryotic selective marker.

The vectors presented in the work of Zou J. et al. can be considered as examples of the structure of vectors that carry the ZFN-based gene editor [17]. The authors proposed a vector that contains a target gene, i.e., a sequence encoding a chimeric protein that consists of the FokI nuclease domain and zinc fingers that direct the nuclease to the desired part of the genome. In addition, there is a nuclear localization signal, which is located on the 5' end of the protein-coding sequence. As a marker of selection, the vector contains the gene for resistance to blasticidin, which provides rapid selection of transfected cells. In other respects, it is a typical vector for the synthesis of recombinant proteins (Fig. 2a). Editors can be directed not only to the cell nucleus but also to the mitochondria [18]. Since the compartment in which the editing should take place was not the nucleus, the transport signal into the mitochondria was placed at the N-end of the protein. The synthesized protein entered the mitochondria, where FokI endonuclease activity was manifested.

Fig. 2.

The structure of vectors for the expression of gene editors in eukaryotic cells. (a) The expression vectors for genome editing using ZFN and TALEN are very similar, only element 8 differs. Targeting is carried out by zinc fingers in the first case and the effector DNA-binding domain in the second case. (1) Replication origin; (2) prokaryotic selective marker; (3) prokaryotic promoter; (4) eukaryotic enhancer; (5 and 11) eukaryotic promoters which specifically interact with RNA polymerase II; (6 and 12) Kozak sequences; (7) nuclear localization signal peptide; (8) zinc fingers/effector DNA-binding domain; (9) FokI endonuclease; (10, 14) polyadenylation signal; (13) eukaryotic selective marker; (b) Genome editing using CRISPR/Cfs. (1) Replication origin; (2) prokaryotic selective marker; (3) prokaryotic promoter; (4) eukaryotic enhancer; (5 and 10) eukaryotic promoters, which specifically interact with RNA polymerase II; (6 and 11) Kozak sequences; (7) signal peptide of nuclear localization; (8) Cas protein with endonuclease activity; (9 and 13) polyadenylation signals; (12) eukaryotic selective marker; (14) U6 promoter, which specifically interacts with RNA polymerase III; (15) guide RNA sequence.

The targeting to the target site in the TALEN-based technology is provided by effector DNA-binding domains, and the cleavage is performed by the FokI nuclease domain. Effector DNA-binding domains are rather conservative small regions that consist of 33–34 amino acid residues with variable residues at positions 12 and 13 that promote interaction with DNA. The sequential arrangement of these domains makes it possible to recognize different motifs in DNA. In this case, a chimeric protein is synthesized that consists of the TALEN domain and FokI nuclease connected by a spacer. Consequently, the vector for the expression of the TALEN-based gene editor will also not differ from the base vector that provides the synthesis of recombinant proteins because the entire editor is one large protein [19] (Fig. 2a). As an example, we can consider the work of Kim Y.H. et al., who tried using the TALEN-based gene editor to change the set of antigens on the surface of erythroid progenitor cells, thus obtaining universal group I blood from the blood of any other group [20]. A nuclear localization signal was attached to the TALEN-encoding nucleotide sequence; on the whole, however, this vector can be called standard, except for one feature, i.e., it lacks a eukaryotic selective marker but this is due to the specificity and requirements of the experiment. The disadvantages of ZFN- and TALEN-based systems include the complexity of creating recognition domains and, sometimes, the inability to choose modules for recognizing the nucleotide sequence. The advantage of these systems is the relatively small size of the expression vector.

Unlike ZFN and TALEN, the CRISPR/Cas-based genetic editor is a complex consisting of a Cas endonuclease and a guide RNA, which is the main feature of the expression vector of the CRISPR/Cas editor [21]. The structure of synthesized guide RNA significantly differs from RNAs synthesized by RNA polymerase II. The guide RNA lacks the cap and poly(A) tail, characteristic components of mRNA. The synthesis of guide RNA is provided by RNA polymerase III; therefore, the vector must include a promoter that can interact with this polymerase. The human U6 promoter is most often used for this purpose [22]. As to the rest, the composition of the vector should be the same as that of a typical vector for the synthesis of recombinant proteins (Fig. 2b). As an example, we can consider a vector from Gabriel C.H et al. [23]. The authors used a CRISPR/Cas-based editor to knock out genes. They used GFP as a selective marker, which was expressed from the same reading frame as the Cas9 protein. To obtain two proteins, the 2A peptide was placed between Cas9 and GFP. Another difference was that the promoter, which controlled the expression of Cas9, was not merged with the enhancer, and, instead of that, an intron was placed next to the promoter. This element can also enhance the expression of the target product.

Thus, it can be stated that the expression vector may include various functional elements in addition to the standard elements. If it is necessary to place the plasmid DNA into viral particles, the vector must contain sites that specifically interact with the viral capsid. Transgene expression can also be modulated by certain regulatory elements. Ultimately, the choice of elements depends on either the requirements of the experiment or the therapeutic application.

STRUCTURAL ELEMENTS OF SYNTHETIC mRNAs FOR DELIVERY OF GENOME EDITORS

Structure of Natural mRNAs

mRNA is synthesized in the nucleus and undergoes various modifications, followed by translation in the cytoplasm. After modification of the 5' and 3' termini, mRNA becomes functionally active. The transcribed mRNA undergoes splicing and two significant modifications. 7-Methylguanosine is attached to 5'-triphosphate in pre-mRNA through a 5'–5' bond with the formation of a so-called cap structure. This structure protects mature mRNA from degradation and promotes nuclear transport and efficient translation. The second modification is the post-transcriptional addition of the poly(A) tail (100–250 adenosine residues) to the 3' end of the RNA molecule. The addition of the poly(A) tail imparts stability to the mRNA molecule, promotes the export of mRNA to the cytosol, and participates in the formation of a translationally competent ribonucleoprotein complex along with the 5'-cap structure. Mature mRNA forms a ring structure (closed loop) and connects the cap to the poly(A)tail through the cap-binding eIF4E protein (eukaryotic translation initiation factor 4E) and poly(A)-binding protein (PABP), which interact with eIF4G (eukaryotic translation initiation factor 4G). Thus, the main requirements for functional mRNA are the presence of the cap (7-methylguanosine) at the 5' end and the poly(A) tail at the 3' end [24].

Transcription and Capping of Synthetic mRNAs

Transcription is one of the main biological processes underlying the central dogma of molecular biology. DNA-dependent RNA polymerases, which perform this stage of transmission of genetic information, are common in all forms of life. As a rule, these enzymes are intricate multisubunit complexes, which transcribe prokaryotic and eukaryotic genes in vivo. mRNA is synthesized in vitro by bacteriophage polymerases (T7, T3, and SP6), one-unit enzymes whose activity requires only Mg²⁺ ions. The DNA molecule (PCR fragment or linearized plasmid), which contains the sequence of the corresponding promoter, acts as a template for in vitro transcription. The enzyme promotes the synthesis of a complementary RNA molecule, followed by dissociation of the transcription complex at the end of the process. The DNA template and the enzyme can be reused in the following reactions. The yield of synthesized mRNA can reach milligram quantities in this process. After transcription, RNA is treated with DNase I to remove the DNA template and is purified before capping. This method makes it possible to obtain RNA with 5'-triphosphate termini in a high yield. The resulting RNAs need to be supplied with the cap structures [25].

For effective translation of synthetic mRNA, it must transformed into mature mRNA by attaching the cap and poly(A) tail during or after the RNA synthesis in separate reactions catalyzed by capping enzymes and poly(A)-polymerase, respectively (Fig. 3). Additionally, modified internal bases or modified cap structures can be included in artificial RNA molecules, which can increase the stability and translational activity of the final molecule. Depending on the chosen copying strategy, two variants of in vitro transcription are used. The standard synthesis with enzymatic capping after the transcription reaction (post-transcriptional capping) or the inclusion of a cap analog during transcription (co-transcriptional capping). The strategies of the in vitro mRNA synthesis depend on the desired scale of synthesis.

Fig. 3.

The structure of synthetic mRNA. (1) Cap structure; (2) 5'-untranslated regions; (3) open reading frame encoding the transgene; (4) 3'-untranslated area; (5) poly(A) tail.

Post-transcriptional mRNA capping is often performed using enzymes of the smallpox vaccine virus. This enzyme complex converts the 5'-triphosphate ends of transcripts in vitro into m7G-cap structures. The cap system of the smallpox vaccine virus includes three active parts (RNA triphosphatase, guanylyl transferase, and guanine-N7 methyltransferase), which are necessary for the formation of the complete cap structure (m7Gppp5'N) using GTP and S-adenosyl methionine, the donor of the methyl group. Additionally, 2'-O-methyltransferase can be included in the same enzymatic reaction, which leads to the formation of the cap 1 structure by methylation of the 2'‑O-position of the nucleotide following the cap. This reaction is a natural modification of many eukaryotic mRNAs [26].

During transcriptional copying, the cap analog is embedded in RNA as the first nucleotide of the synthesized molecule. The cap analog is introduced into the transcription reaction along with four standard nucleoside triphosphates in a 4 : 1 ratio of optimized cap to GTP. This makes it possible to initiate the formation of a significant number of cap-containing transcripts among the synthesized RNA molecules. As a result, a mixture of transcripts is formed, of which ~80% are capped and the rest have the 5'-triphosphate termini. The decrease in the total yield of RNA products is caused by a lower concentration of GTP in the reaction. In the case of co-transcriptional capping, several synthetic cap analogs are used. The most common analogs are the standard cap, 7-methylguanosine (m7G), and the symmetric cap analog (ARCA), also known as 3'-O-me-7-meGpppG. The standard m7G cap analog can be embedded in the RNA molecule in both the forward and reverse orientation, which significantly reduces the overall level of translation of mRNA. ARCA is methylated at the 3'-position of m7G, which makes it possible to embed this cap analog only in the direct orientation and obtain a pool of capped mRNAs with a translational activity of 100%. The yield of the products in this reaction is lower than in the transcription reaction without synthetic cap analogs. The ARCA cap structure can be converted to the cap 1 structure using cap-dependent 2'-O-methyl transferase and S-adenosylmethionine in the subsequent enzymatic reaction [27]. It should also be noted that several other synthetic cap analogs with improved properties have been synthesized including the cap 1 analog from Trilink Biotechnologies (United States).

Modified Nucleotides and Optimization of the Nucleotide Composition of Synthetic mRNAs

Natural RNAs, as a rule, tend to mature not only from the 5' and 3' ends but also by modifying certain nucleotides inside the molecule. The use of modified nucleotide analogs in synthetic mRNAs is considered the most effective way to avoid activation of cellular sensors (TLR3, TLR7, TLR8, PKR), which trigger an immune response to foreign (usually viral) RNAs. This means that RNA can be synthesized in vitro using a mixture of modified nucleoside triphosphates instead of natural triphosphates of A, G, C, and U. Modified nucleotides, such as naturally occurring 5‑methylcytosine and/or pseudouridine (including N1-methylpseudouridine) are most often used instead of C and U, respectively [28]. It has been shown that the use of modified nucleoside triphosphates for mRNA synthesis significantly increases the stability of mRNAs in the cell, the efficiency of their translation, and reduces the cellular immune response to these mRNAs. This is especially important in some therapeutic applications of mRNA, e.g., in gene editing, protein replacement therapy, or differentiation of stem cells using mRNA-encoded transcription factors. For example, the expression of the ZFN gene editor in mouse lung tissues was significantly higher when using synthetic mRNAs that contain modified nucleotides than when using unmodified mRNA [29]. It is important to note that the choice of nucleotides can affect the overall yield of the mRNA synthesis in vitro. It is possible to obtain mRNA with the desired modification. Transcripts with the replacement of one or more nucleotides can be capped post-transcriptionally or co-transcriptionally by including ARCA or another cap analog. It should also be noted the importance of chromatographic purification of the final mRNAs to minimize the nonspecific immune response [30]. Another factor affecting the efficiency of synthetic RNA translation is the optimization of the composition of the transgene codons taking into account both the organism and the type of tissues and cells. During optimization, some rare codons are replaced by synonymous ones, which are translated more efficiently. Most often, this leads to increased translation and enhances the stability of RNA. However, this approach is not always suitable for therapeutic proteins. In some cases, for the correct folding of proteins encoded by synthetic mRNAs, it is necessary to reduce the translation rate at certain regions of the sequence; therefore, the value of codon optimization in such situations is ambiguous [31].

5'- and 3'-Untranslated Regions in Synthetic mRNAs

The stability and translational activity of mRNA in the cell are largely determined by the 5'- and 3'-untranslated regions (UTR). These regulatory regions flank the coding sequence of the transgene mRNA. They contain various motifs and form secondary structures that determine the stability of mRNA, recognition of mRNA by ribosomes, and interaction with components of the translational apparatus [32]. The inclusion of these sequences in synthetic mRNA can improve the translation and stability of mRNA. Many UTRs, which enhance mRNA translation, are natural. For example, UTR in mRNAs of alpha and beta globins are widely used to create synthetic mRNAs. Moreover, the stabilizing effect can be enhanced by sequentially placing two copies of 3'-UTR of beta-globin. UTRs of human heat shock protein 70, albumin, and alphavirus proteins have similar effects on synthetic mRNAs [33].

Synthetic UTRs developed in the laboratory can be used as an alternative to natural UTRs. For example, the de novo constructed 5'-UTR sequence with a length of only 14 nucleotides provides an expression level comparable to that characteristic for 5'-UTR of human alpha-globin [34]. Modern screening technologies allow experimental selection of UTRs with improved properties [35].

Poly(A) Tail of Synthetic mRNAs

The polyadenylated 3'-terminal region of mRNA, the so-called poly(A) tail, is an important structural element that determines the lifespan of mRNA molecules. With some exceptions, the poly(A) tails of most natural mRNA molecules in mammalian cells are up to 250 nucleotides long, and they gradually shorten during the lifetime of mRNA in the cell. In addition, the poly(A) tail plays an important role in translation, specifically in the formation of a translationally active mRNA complex with translation factors [36]. The size of the tail affects the stability of mRNA, thus preventing 3'-exonuclease degradation. Therefore, it is desirable to include poly(A) tails of approximately 100–120 nucleotides in length in synthetic mRNAs. It has been experimentally shown that a length of the poly(A) tail of more than 120 nucleotides significantly increases recombination events in bacterial cells involving this sequence. As a result, this affects the stability of the plasmid, which is used as a template for the mRNA synthesis in vitro and reduces the yield of the plasmid during its development in bacterial culture. Polyadenylation of mRNA in vitro can be carried out by either the enzymatic attachment of a poly(A) tail to the capped mRNA using recombinant poly(A) polymerase or the encoding of this sequence in a plasmid vector [25]. Polyadenylation by a separate enzyme adds an extra stage to the process of the mRNA synthesis and does not allow accurate control of the number of nucleotides included in all molecules during the reaction. Therefore, the second approach, which involves the use of a plasmid for encoding poly(A) tail, is more suitable for the industrial production of therapeutic mRNAs.

Synthetic mRNAs to Deliver Genome Editors

The safe nonviral delivery of genome editors opens new prospects for the therapeutic application of gene editing of therapeutic mRNAs. This approach worked well during the COVID-19 pandemic. The combination of mRNA and lipid nanoparticles ensures the safe expression of antigenic or therapeutic proteins in vivo, which has been confirmed by the results of clinical and preclinical trials. In recent years, successful experiments on genomic editing have been carried out with the use of synthetic mRNAs. The transtiretin (Ttr) gene was edited in the liver of mice, which led to a decrease in the level of the TTR serum protein by more than 97% [37]. The authors used an original delivery system based on lipid nanoparticles, which made it possible to simultaneously pack Cas9-encoding mRNAs along with guide RNAs [37]. Intravenous co-delivery of Cas9 mRNA and guide RNA led to DNA editing in liver, kidney, and lung tissues in mice [38]. The use of nanoparticles of a different composition ensured the effective release of RNA inside the cells in the reducing medium. The authors of [39] have shown effective knockout of the reporter gene in embryonic kidney cells, accumulation of intravenously injected mRNA-bearing nanoparticles in liver tissues, and successful knockout of the target gene (up to 20% of the serum level). Recently, the possibilities of gene editing using the RNA delivery platform have been shown by the example of correction of muscular dystrophy in muscle cells obtained from a wide range of donors. The safe delivery of gene editors using the synthetic RNA platform in terms of embedding the transgene into the genome has been confirmed, and the high efficiency of the editing with the possibility of dosing SpCas9 activity has also been shown [40]. The therapeutic potential of delivering gene editors using lipid nanoparticles that contain synthetic RNAs has been shown. In this case, knockdown of the Angptl3 gene in the mouse liver, which was performed to reduce the level of the ANGPTL3 protein, led to a decrease in blood lipid levels and correction of hypercholesterolemia. Researchers noted a significant duration of the therapeutic effect (up to 100 days after a single injection), the absence of nonspecific editing in several of the most possible places, and a lack of hepatotoxicity [41].

METHODS OF DELIVERY OF DNA- AND RNA-BASED EXPRESSION VECTORS IN VITRO AND IN VIVO

The successful application of genome editing tools requires not only correctly designed expression vectors, but also effective and safe delivery methods for carriers of genetic information to cells in vitro or to certain tissues in vivo. These methods are divided into three main types (Table 1). The first is biological transfection with the use of viral vectors or virus-like particles. The second and the third methods are the use of physical and chemical transfection. In this review, we do not consider viral transgene delivery vectors; we will focus only on physical and chemical transfection methods for the delivery of nucleic acid-based expression vectors.

Physical Transfection Methods

Electroporation. Electroporation, or electrotransfection, is the most common physical transfection method. Traditionally, electrotransfection is carried out in vitro in a cuvette with suspended cells; however, this method is also applicable in vivo. A cellular suspension that contains the DNA plasmid or mRNA of interest is placed between two electrodes. A medium with cells is subjected to a series of short electrical pulses, which leads to a sharp change in the voltage on the cell membrane (reaching a critical threshold between 250 and 500 mV) and the appearance of pores in the cell membrane for penetration of nucleic acid into the cytosol. DNA and RNA move in an electric field from the anode to the cathode [42]. Interestingly, larger pores in the cell membrane are formed from the side of the anode, which also contributes to the penetration of fairly large nucleic acid molecules into the cell. When the electrical impulses are terminated, the pores gradually relax and close. Electrotransfection is a very effective method for DNA and RNA delivery; unfortunately, a large number of cells die during this procedure.

It should also be noted that there are differences in the efficiency of transfection of different cell lines. The classical version of transfection can be used only for suspensions of cells; adhered cells cannot be modified. However, Maschietto et al. recently proposed an electrotransfection method that circumvents this limitation [43]. The authors proposed the use of plates for cell cultivation, the bottom of which is covered with microelectrodes. The plates are chips for electroporation, which are supplied with arrays of thin-film capacitive microelectrodes. Each individual microelectrode is an octagonal structure formed by highly conductive p-type silicon, which is covered with a layer of silicon oxide 15 nm thick. Cells grow on the surface of these electrodes, which makes it possible to use significantly lower currents without losing the efficiency of transfection. It is claimed that, in this case, the efficiency of transfection of CHO-K1 cells, reaches 60–80% without a high mortality rate of cell culture after the procedure. It was also possible to transfect differentiated neurons after six days of cultivation, although with low efficiency of ~10%. To apply this approach in vivo, it was necessary to develop several electrodes suitable for use in various organs and tissues [44]. In general, this method is well accepted for use in clinical practice due to its simplicity and relatively few side effects. Kawasaki et al. proposed electroporation for the successful delivery of the CRISPR/Cas genetic editor in utero [45]. Thus, transfection by electroporation is at the moment a very effective and cheap approach to the delivery of nucleic acids to cells both in vitro and in vivo, which is limited only by the availability of equipment.

Gene gun: the principle of operation and application. Like electroporation, this approach is applicable to the delivery of both plasmid DNA and mRNA. Initially, this method was used to modify plant cells; however, its potential is much wider. Using a gene gun, various cell types can be modified both in vitro and in vivo. The method is based on the high-speed bombardment of cells with complexes of heavy metal particles of about 1 µm in size coated with nucleic acids. At first, tungsten particles were used; however, this metal is toxic to cells, so it is often replaced with biologically inert gold particles [46]. Particles can be accelerated in different ways. Currently, the most common acceleration method is the use of a short pulse of an inert gas, e.g., helium. The dispersed particles penetrate the cell membrane and deliver genetic material to various cellular compartments. Nucleic acids can enter either the cytosol, nuclei, or other organelles. Plastids and mitochondria can be modified using this approach. The main limitations when using biolistics for in vivo transfection are the size of the equipment and the depth of penetration of particles into the body. This approach is used mainly for the transfection of epithelial and subcutaneous muscle tissues because of their accessibility [47].

The main advantages of this method include its high efficiency, the possibility of delivering several plasmids or mRNAs simultaneously, a wide range of modification objects, and the possibility of modifying differentiated cells and cells growing in adhesive culture. But there are also a number of disadvantages. The first is the high cost of the gene gun with relatively cheap further operation of the device. The second significant drawback is the high cellular mortality during the transfection procedure. However, as reported by O’Brien and Lummis, the use of lower pulse pressure (about 345 kPa) significantly reduces the degree of cell damage [48]. The gene gun delivery method continues to evolve. The important advantages of this method include versatility and applicability to cells of different types, while its use is constrained by the high cost of equipment. It is likely that reduction in cost will attract more attention to this method.

Sonoporation for DNA and RNA delivery in vitro and in vivo. The sonoporation method promotes the transfer of nucleic acids by exposing cells to ultrasound waves, which results in effects such as cavitation, radiation pressure, and micro-flows. Cavitation, i.e. the appearance and collapse of micro-bubbles of air, provides a very high local pressure and an increase in temperature, thus leading to a violation of the integrity of the cell membrane if the bubble collapses at the cell surface. Nucleic acid penetrates through these pores under the influence of other effects that occur during low-frequency ultrasound treatment [49]. There is also the method of plasmid DNA delivery using a combination of sonoporation and microfluidic technologies, hereinafter referred to as the method of acoustofluidic sonoporation. Belling et al., the authors of this method, state that it is possible to transfect a large number of cells at a high speed. In this method, a mixture of cells and plasmid DNA is treated with ultrasound while passing through a capillary. Researchers report that they managed to achieve a transfection rate of about 200 000 cells/min in the case of the Jurkat cell line, with cell viability after the procedure being about 80% [50]. This method is an effective and affordable analog of physical transfection methods, although it is necessary to study the effects of ultrasound on cells, especially on cell nuclei, which are also susceptible to local damage as a result of cavitation. Sonoporation is also used for transfection in vivo but its effectiveness, in this case, is significantly lower than in vitro. Nevertheless, the use of this approach made it possible to successfully deliver a CRISPR/Cas-based genetic editor for the treatment of male pattern baldness. The researchers directed the genetic editor to the SRD5A2 gene using a combination of nanoliposomal particles as a vehicle and sonoporation as an ‘activator’ of particles, for delivery. Under the action of cavitation, the particles burst and delivered plasmid vectors directly to the cells of the hair follicles [51].

Phototransfection. Laser transfection works by the local destruction of the cell membrane. The laser beam is focused under a microscope on the cell membrane in a 1–2 µm area, followed by a series of gentle high-intensity pulses, which leads to local destruction of the membrane. In this case, plasmid DNA from the nutrient medium penetrates into the cell [52]. The duration of radiation is femtoseconds, the used wavelength can vary significantly and depends, as a rule, on the available equipment, and power in the range of 50–100 mW is recommended [53]. However, the method has a low transfection efficiency (~25%) and a significantly lower throughput compared to the previously considered methods. Therefore, it is more suitable for point transfection. One of the advantages of phototransfection is the applicability of the method to cells that grow in both suspension and adhesive culture. The low cytotoxicity of this approach is also noted, which may be important for the transfection of single cells [54].

Chemical Transfection Methods

Transfection with calcium phosphate. Transfection with calcium phosphate was one of the first methods of cell transfection in vitro [55]. The method is based on the cellular absorption of complexes that consist of plasmid DNA and calcium phosphate. These complexes are prepared by mixing CaCl₂ with DNA, followed by the addition of a buffer that contains phosphorus compounds, for example, HEPES [56], which leads to precipitation of the desired complexes. The resulting complexes are adsorbed on the cells from the nutrient medium. The main advantages of this method are its simplicity and low cost. The disadvantages include the relatively low efficiency of transfection, especially of differentiated cells, the relatively high level of cytotoxicity, and applicability only in vitro [57]. The method was proposed in the 1970s but is still widely used, and attempts are being made to optimize it to increase its effectiveness [58].

Transfection with poly-L-lysine. This method is also based on the creation of complexes of nucleic acids and a carrier, which are absorbed by the cell. The carrier is a polymer consisting of lysine amino acid residues. A molecule that contains ~10 lysine residues has a positive charge, which makes it possible to form complexes with negatively charged nucleic acid molecules. The complex of poly-L-lysine with nucleic acid also has a total positive charge, which provides its interaction with the cell membrane. In such the ‘classic’ version, this method of DNA delivery has a number of disadvantages. First, the microparticles are absorbed by lysosomes, followed by degradation, which negatively affects the overall transfection efficacy. Second, the poly-L-lysine complex itself has pronounced cytotoxicity [59]. However, these disadvantages can be compensated for by using additional agents. Thus, the addition of an endosomolytic agent, e.g., glycerin, to the cellular medium before transfection solves the problem of the release of poly-L-lysine complexes from endosomes. Modification of poly-L-lysine with PLGA (polylactide-co-glycolide) reduces cytotoxicity and increases the overall efficiency of transfection. If necessary, poly-L-lysine complexes can be targeted to certain cell types by attaching signal sequences to the polymer for binding to surface cell receptors. Similarly, some studies solve the problem of absorption of the poly-L-lysine complex by endosomes [60]. One of the main advantages of this method is its easy modification for the needs of the experiment and application both in vivo and in vitro. However, this method does not provide the highest level of transfection. In addition, the resulting complexes have different sizes, which can also affect the effectiveness of transfection.

Transfection with polyethylenimine. Polyethylenimine (PEI) is a polymer with high cationic potential due to its large number of amino groups. It is possible to synthesize linear or branched PEI molecules [61] of a given size and with various modifications. PEI interacts with negatively charged nucleic acids, thus forming complexes that are ready for transfection. The interaction with a negatively charged cell membrane is caused by the total positive charge of the complex. Unlike poly-L-lysine complexes, PEI complexes leave endosomes quite easily due to the ‘proton sponge’ or buffering effects [62]. All this leads to a high efficiency of transfection, which is the main advantage of this method. The main disadvantage of PEI is its cytotoxicity, which is directly proportional to the particle size and the efficiency of transfection. Neutral hydrophilic copolymers, such as polyethylene glycol, are added to the complexes to reduce their cytotoxicity [63]. Transfection with PEI has found wide application in vitro as an effective approach to transgene delivery. However, this approach is of little use for the delivery of genetic material in vivo.

Transfection with chitosan. Chitosan is a polysaccharide that consists of repeating units of D-glucosamine and N-acetyl-D-glucosamine. At pH 6.5, chitosan is protonated and becomes soluble in water. After processing, it acquires a high positive charge. Like other cationic polymers, chitosan forms complexes with negatively charged nucleic acids. This polymer contains two hydroxyl groups, which are easily modified for the tasks of the researcher [64]. The mechanism of chitosan penetration into cells is the same as that of PEI or poly-L-lysine complexes. Unlike other polymers, chitosan has low cytotoxicity. However, the efficiency of transfection with this polysaccharide is significantly lower than in other transfection methods. Transfection with chitosan is considered a promising alternative to the use of PEI. Its low cytotoxicity and the possibility of modification make chitosan a fairly convenient platform. Although studies are being performed to improve the efficiency of transfection using this polymer, an acceptable level of efficiency of PEI has not yet been achieved [65].

Lipofection. Lipofection is based on the delivery of DNA and RNA in lipid complexes. Positively charged lipids and negatively charged nucleic acids are used to create complexes. As an example, we can consider the first commercially available drug Lipofectin®, which consists of the positively charged DOTMA lipid and the neutral DOPE lipid. To date, transfection is performed using various lipids, which differ in properties and degree of impact on cells. In particular, lipids, such as DOTAP and DODAP for the preparation of the YSK05 liposomes are significantly less cytotoxic than Lipofectin® components, and the MVLBG2 lipid dendrimer has a high positive charge, which promotes its effective interaction with DNA [66]. Lipid vesicles that contain DNA or RNA are formed when mixing nucleic acid and cationic lipids. The total charge of the vesicle should be positive to provide interaction with the cell membrane due to electrostatic forces. Vesicles are absorbed by cells by endocytosis.

Lipofection provides a high level of cell transfection in vitro, but lipid vesicles have high cytotoxicity, especially if we consider a classic variant, such as Lipofectin® [67]. Another disadvantage is the high heterogeneity of the size of the vesicles. It is believed that this heterogeneity negatively affects the transfection efficacy. However, the simplicity of the chemical synthesis and modification of cationic lipids make this approach very flexible. For example, it is possible to obtain lipid nanoparticles (LNP), which are ‘dissolved’ in the lysosomes at a decreased pH value, thus increasing the efficiency of transfection. These LNPs are produced using ionizable lipid, polyethylene glycol, auxiliary phospholipid, and cholesterol. The key component is an ionizable lipid that is neutral at physiological pH values, but after penetration into the endosome with internal acidic pH, acquires a positive charge. A change in the particle charge leads to the destruction of endosomes and the release of nucleic acids [68]. This approach was used in vivo for the knockdown of the Angptl3 gene in the liver of mice [41]. Lipofection, despite its pronounced cytotoxicity, allows transfection to be performed with high efficiency. In addition, the possibility of modifying each unit of the cationic lipid and the complex as a whole makes this method very flexible.

Table 1.  .

Methods of delivery of DNA- and RNA-based expression vectors in vitro and in vivo

Method	Principle	Advantages	Disadvantages
Electroporation	Penetration of nucleic acids occurs under the influence of a series of short electrical pulses	Simplicity of the procedure; low cost; wide range of electrodes; applicability both in vitro and in vivo	Specialized equipment is required; the procedure is accompanied by cell/tissue damage
Biolistics	Metal particles coated with nucleic acid molecules are dispersed by an inert gas pulse and penetrate the cell membrane	High transfection efficiency; possibility of simultaneous delivery of different vectors; a wide range of modification objects; applicability both in vitro and in vivo	High cost of the gene gun and subsequent operation; high level of cell damage as a result of the transfection procedure; limited number of tissues available in vivo
Sonoporation	Transfer of nucleic acids due to exposure of cells to ultrasound waves	High rate and efficiency of transfection in vitro; high cell survival; applicability both in vitro and in vivo	Availability of equipment; limited use and lower efficiency in vivo
Phototransfection	Local destruction of the cell membrane by short laser pulses	The method is applicable to cells growing both in suspension and in adhesion; low level of cytotoxicity	Applicable only in vitro; low efficiency of transfection relative to other physical methods; low bandwidth; labor intensity
Calcium phosphate	Absorption by cells of complexes that consist of nucleic acids and calcium phosphate	Simplicity of the procedure; low cost	Low transfection efficiency; pronounced cytotoxicity; applicability only in vitro
Poly-L-lysine	Absorption by cells of complexes that consist of nucleic acids and poly-L-lysine	Easily modified for the needs of the experimenter; applicability both in vivo and in vitro	Low transfection efficiency; different sizes of the resulting complexes; pronounced cytotoxicity of unmodified poly-L-lysine complex
Polyethyleneimine	Absorption by cells of complexes that consist of nucleic acids and polyethyleneimine	High efficiency of transfection in vitro	High cytotoxicity; applicability mainly in vitro
Chitosan	Absorption by cells of complexes that consist of nucleic acids and chitosan	Low cytotoxicity; applicability both in vivo and in vitro	Low transfection efficiency
Lipofection	Absorption by cells of complexes that consist of nucleic acids and a mixture of lipids	High transfection level; applicability both in vivo and in vitro; easy modification for the experimental needs	High cytotoxicity; different sizes of the resulting complexes

CONCLUSIONS

In this paper, we have considered the structural elements of expression vectors based on DNA and synthetic mRNAs and the methods for their delivery both in vitro and in vivo. Optimization of the structural components of these vectors makes it possible to effectively express therapeutic proteins and gene editing tools both in cells and in tissues for successful implementation of gene therapy. The choice of appropriate vector platforms, their structural elements, and transgene delivery methods make it possible to solve a wide range of experimental biomedical and therapeutic tasks, which is confirmed by a growing amount of scientific data and the results of preclinical and clinical trials on the use of gene editors for the correction of hereditary and acquired pathologies.

FUNDING

The work was supported by the program of Ministry of Higher Education and Science of the Russian Federation (agreement no. 075-10-2021-113, ID of the project RF-193021X0001).

COMPLIANCE WITH ETHICAL STANDARDS

The authors state that there is no conflicts of interest. This article does not contain any studies involving humans or animals as objects of research.