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Cloning and Transformation with Plasmid Vectors

• PMID: 34725175
• DOI: 10.1101/pdb.top101170

Plasmids occupy a place of honor in molecular cloning: They were used in the first recombinant DNA experiments and, 40 or more years later, they remain as the carriage horses of molecular cloning. After almost half a century of sequential improvement in design, today's plasmid vectors are available in huge variety, are often optimized for specific purposes, and bear only passing resemblance to their forebears. Here, various features of plasmid vectors and methods for transforming E. coli cells are introduced.

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• Methodology article
• Open access
• Published: 12 October 2011

FastCloning: a highly simplified, purification-free, sequence- and ligation-independent PCR cloning method

• Chaokun Li 1 ,
• Aiyun Wen 1 ,
• Benchang Shen 1 , 3 ,
• Yao Huang 2 &
• Yongchang Chang 1  

BMC Biotechnology volume  11 , Article number:  92 ( 2011 ) Cite this article

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Although a variety of methods and expensive kits are available, molecular cloning can be a time-consuming and frustrating process.

Here we report a highly simplified, reliable, and efficient PCR-based cloning technique to insert any DNA fragment into a plasmid vector or into a gene (cDNA) in a vector at any desired position. With this method, the vector and insert are PCR amplified separately, with only 18 cycles, using a high fidelity DNA polymerase. The amplified insert has the ends with ~16-base overlapping with the ends of the amplified vector. After Dpn I digestion of the mixture of the amplified vector and insert to eliminate the DNA templates used in PCR reactions, the mixture is directly transformed into competent E. coli cells to obtain the desired clones. This technique has many advantages over other cloning methods. First, it does not need gel purification of the PCR product or linearized vector. Second, there is no need of any cloning kit or specialized enzyme for cloning. Furthermore, with reduced number of PCR cycles, it also decreases the chance of random mutations. In addition, this method is highly effective and reproducible. Finally, since this cloning method is also sequence independent, we demonstrated that it can be used for chimera construction, insertion, and multiple mutations spanning a stretch of DNA up to 120 bp.

Our FastCloning technique provides a very simple, effective, reliable, and versatile tool for molecular cloning, chimera construction, insertion of any DNA sequences of interest and also for multiple mutations in a short stretch of a cDNA.

Molecular cloning is one of the most widely used techniques in biomedical research laboratories. Traditionally, molecular cloning joins insert and vector by T4 DNA ligase after restriction digestion to excise insert from a donor vector or from a PCR product with restriction enzyme recognition sites added to the ends [ 1 ]. Although this is a widely used method, it involves multiple steps and is time consuming. This multi-step process also makes it difficult or complicated for troubleshooting. To overcome the difficulties encountered in the original cloning method, many other alternative cloning methods have been developed over the last two decades. These methods include TA cloning [ 2 ], ligation independent cloning with T4 DNA polymerase [ 3 , 4 ], GATEWAY recombinational cloning [ 5 ], and more recent sequence- and ligation-independent cloning kits, such as CloneEZ (GenScript USA Inc., Piscataway, NJ, USA), one step cloning [ 6 ], and overlap extension PCR cloning [ 7 ]. However, each of these techniques has its own limitations. For example, TA cloning uses regular Taq DNA polymerase to add a single 3'-A overhang to the ends of the PCR product. The PCR product is directly cloned into a TA cloning vector with a complementary 3'-T overhang in both ends without restriction digestion. The limitations of this method are low fidelity of Taq DNA polymerase causing unwanted mutations and requirement of subcloning into the final target vector with restriction digestion and ligation. The early ligation independent cloning uses the 3'-exonucnease activity of T4 DNA polymerase to create 15-base 5'overhangs in the ends of insert and complementary 5' overhangs in the ends of vector. This technique requires specific sequences to create 15-base overhangs. Gateway recombinational cloning uses site-specific recombination to transfer cDNAs between donor and destination vectors, which requires additional specific sequences for recombination. The latest ligation-independent cloning, such as CloneEZ and In-Fusion cloning kits, uses some DNA polymerase to generate sticky ends in the vector and insert without specific sequence requirement, except for restriction sites to linearize the vector. However, the new ligation independent cloning still requires purification of the digested vector and PCR-amplified insert, and the purchase of purification and cloning kits. Similarly, overlap extension PCR cloning also requires purification of the first round PCR products (vector and insert) and an additional round overlap extension PCR, which usually generates multiple bands, for producing linked vector and insert. One-step "quick assemble" cloning does not need purification of PCR products. However, it includes two sequential 35-cycle PCRs with a total number of 70 cycles. The first-round PCRs are used to amplify insert and linear vector. The second-round PCR is essentially the overlap extension PCR to assemble vector and insert into a single linear PCR product. Another ligation independent cloning technique, using nick DNA endonuclease to create long single-strand 5' overhangs in the vector and PCR-amplified insert [ 8 ], requires specific sequences for nick DNA endonuclease and purification of the PCR product.

Purification of PCR product not only takes extra time and requires purification kit, but also potentially creates additional problems. For example, to compensate for the loss of PCR product during purification, the number of PCR cycles is generally 25-30. High number of PCR cycles increases the chance of random mutations or, in our experiences, dramatically decreases cloning efficiency for the PCR products generated by high fidelity DNA polymerases. This is especially true for the enzyme with high processability, such as pfuUltraII DNA polymerase. The decrease in cloning efficiency cannot be completely overcome by using state-of-the-art cloning kits, such as CloneEZ. Using this kit (with gel purification of digested vectors and PCR-amplified inserts), the successful rate in our laboratory still varies significantly with overall successful rate less than 50%. In the PCR-based QuickChange mutagenesis, the number of PCR cycles is recommended to be 12-18. Increase in PCR cycles could decrease efficiency according to the QuickChange protocol. Thus, these phenomena made us believe that the proofreading PCR enzymes could potentially damage the ends of PCR products if the number of PCR cycles increases. This could be due to the fact that the high fidelity DNA polymerases with the proofreading ability have 3' exonuclease activity. In the presence of dNTPs, the ends of PCR products can be protected from this 3' exonuclease activity. It is possible that depletion of dNTPs in high number of PCR cycles or dilution of dNTPs in the early stage of purification would weaken the 3' ends protection against 3' exonuclease activity of the DNA polymerase, which results in the damage of PCR products and makes cloning extremely difficult.

To circumvent the above-mentioned problems, we developed a cloning method termed FastCloning. With this method, both insert and vector are amplified by 18 PCR cycles with a high fidelity DNA polymerase. The unpurified PCR products of the vector and insert are then directly mixed at some ratios (see below) and digested by Dpn I restriction enzyme to destroy their methylated DNA templates. Finally, the digested mixture is transformed into competent cells to obtain the target clones. The PCR amplification of vector is designed to make it possible to clone an insert into any position of the vector, to bypass vector digestion (and restriction site limitation) and purification steps, and to be compatible with Dpn I digestion of the insert without further inactivation of the enzyme. Thus, this method can be used to construct cDNAs of fusion proteins or chimera without limitation by available restriction sites. In this study, we experimentally validated all these applications with our new method. In addition, a similar method can be easily adapted for deletion of a DNA fragment.

Figure 1 illustrates the highly simplified procedure of our FastCloning method. Briefly, after gel confirmation of PCR products, the remaining unpurified PCR reactions, containing amplified vector and insert, are mixed and digested with Dpn I for 1 hour at 37°C. The digested mixture is then directly transformed into chemically competent Stratagene XL-10 Gold or NEB 10-beta E. coli cells.

The procedures for FastCloning: Step 1. PCR amplification of vector and insert. Note that the primer pair for insert amplification has 16-base tails overlapping with the PCR-amplified vector ends. Step 2. Dpn I digestion. The parent DNA templates (if in a plasmid) for PCR amplification needs to be methylated in order to be compatible to Dpn I digestion. Although the detailed mechanism is not known, it is likely that the 3' exonuclease activity of the high fidelity DNA polymerase directly creates sticky ends for the overlapped regions of the vector and insert during Dpn I digestion, allowing them to form a circular construct with nicks. Step 3. transformation into competent E. coli . cells. The nicks will be repaired after transformation into the bacteria.

The primers were designed with Oligo Analyzer 1.5 ( http://www.genelink.com ) to have an annealing temperature around 60°C (Nearest Neighbor method). The forward primer for vector amplification is in the 3' side of the polylinker region. The reverse primer for vector amplification is in the 5' side of the polylinker, and its reverse and complementary sequence was generated by Oligo Explorer 1.5. The primers for insert amplification have insert-specific sequences and additional 15-17 bases (depending on the GC content) overlapping with the vector ends. The specific sequences of primers used in this study are listed in Table 1 . All the primers used in this study were synthesized by Invitrogen Corporation (Carlsbad, CA).

The PCR reaction components were: 50 μl total volume, 0.5 μl Phusion DNA polymerase (New England Biolabs, Ipswich, MA), or 0.8 μl Pfu Turbo, or PfuUltra DNA polymerase (Agilent Technologies, Inc, Santa Clara, CA), 5 μl 10× buffer; 5 μl of 2.5 mM dNTPs; 10 ng of plasmid DNA template; 5 pmol of each primer. The PCR cycling parameters were 98°C 3 min, (98°C 10 sec, 55°C 30 sec, 72°C 20 sec/kb) × 18 cycles, 72°C 5 min, and 4°C infinite for Phusion DNA polymerase, and 95°C 3 min, (95°C 15 sec, 55°C 1 min, 72°C 1 min/kb) × 18 cycles, 72°C 5 min, and 4°C infinite for Pfu Turbo or PfuUltra DNA Polymerase. The PCR products (5 μl for each product) were examined with 1% agarose gel electrophoresis with ethidium bromide staining using VWR Mini Gel electrophoresis setup (VWR International, Marietta, GA, USA) running at 100 V for 30 min. The PCR products were then visualized under a UV transilluminator, and gel pictures were taken using an AGFA scanner.

After confirmation of PCR products, 1 μl of Dpn I enzyme (New England Biolabs) was added into the remaining unpurified PCR reactions (45 μl for each product) for vector or insert separately. The vector and insert were then mixed with 1:1 ratio (1:1, 1:2, and 1:4 for α9 nAChR subunit), and digested at 37°C for 1 hour. Two micro-liters (2, 4, and 8 μl for α9 nAChR) of the digested vector-insert mixture were then added to 40 μl of chemically competent XL-10 Gold E. coli cells (prepared with rubidium chloride method) unless indicated otherwise. The mixture was then incubated for 30 min on ice. After heat shock at 42°C for 45 sec, 350 μl of SOC medium was added to the mixture. After 60 min shaking at 37°C and 350 rpm with an Eppendorf Thermomixer, the entire content was plated onto the LB agar plate containing 100 μg/ml ampicillin. The plates were then incubated at 37°C overnight. Next day, colonies from each constructs were picked for PCR confirmation of each construct using GoTaq DNA polymerase (Promega, Madison, WI, USA) and vector specific primers, and also for inoculation in the LB medium (with ampicillin) for overnight culture of each clone for mini-prep. The DNA mini-prep was performed using QIAprep Spin Miniprep Kit (QIAGEN, Valencia, CA, USA). All the cloned sequences were finally confirmed by automated DNA sequencing at the DNA lab of the Arizona State University using primers in the vectors.

Figure 2 illustrates the application of this method to construct cDNAs encoding chimeric or fusion proteins. In this case, the PCR amplification of the vector also includes part of the cDNA in both ends. To insert a cDNA encoding a full-length protein, such as green fluorescence protein, the insert amplification should cover the entire coding region of the cDNA. For homologous domain swap chimera, insert amplification only needs to cover the corresponding region of a cDNA encoding a homologous fragment of protein. The detailed experimental procedures for chimera construction are essentially the same as cloning.

Chimera construction or insertion . Top left : PCR amplification of both ends of the parent cDNA along with the vector. For chimera construction, one fragment of the cDNA needs to be replaced. Thus, the forward primer is immediately downstream of the fragment to be substituted, and the reverse primer will be immediately upstream of the fragment to be substituted. For insertion, however, two primers will be next to each other without skipping a single base. Top right: Insert amplification of the equivalent region of a homologous gene (cDNA) for chimera construction. However, for insertion of a cDNA encoding a full length protein, such as green fluorescent protein, the insert amplification will cover the entire cDNA. The remaining procedure is the same as in Figure 1.

For insertion (or substitution) of a short DNA fragment (< 120 bp), the insert can be directly included in the primer sequences for vector amplification (Figure 3 ). This is a very convenient way to insert a short tag (such as myc tag or FLAG-tag) or to replace a short DNA fragment for chimera construction without limitation of the availability of specific DNA template for the insert. Introducing multiple mutations in a short (up to 120 bp) stretch of a cDNA is equivalent to replacing a short DNA fragment for chimera construction.

Chimera construction or insertion with a short DNA fragment . To replace a short stretch of DNA, such as a DNA fragment encoding a transmembrane region of a nAChR subunit for chimera construction, or to insert a short tag, such as FLAG-tag, into some part of a protein, amplification of the insert is not necessary. In this case, the insert can be directly included in two primers for single PCR amplification of the cDNA along with the vector. The forward primer starts immediately downstream of the insertion site and has a tail with the 3' part of the insert. The reverse primer starts immediately upstream of the insertion site and has a tail with 5' part of the insert. Two primers only require ~16-base overlap. Thus, for a 120 bp insertion, each primer needs to have a 68-base tail for insertion and ~17-22 bases for annealing (depending on the GC content). The total length of each primer will be about 85-90 bases. Dpn I digestion and transformation are the same as in Figure 1.

Results and Discussion

As a proof of principle, we subcloned several cDNAs into different vectors (human nicotinic receptor (nAChR) α4, α9 and β2 subunits, and serotonin receptor type 3A (5-HT 3 A) subunit into the pGEMHE vector, human endothelial cell-specific molecule 2 (ECSM2) into the p3XFLAG-CMV-14 vector, and Akt3v1 or Akt3v2 into pLXSN vector). pGEMHE is a vector containing 5' and 3' untranslated regions (5'UTR and 3'UTR) of Xenopus β-globin, which is highly expressed in Xenopus oocytes. With this vector, the protein expression level of the inserted gene in Xenopus oocyte can be increased by up to 200-fold [ 9 ].

We first optimized cloning conditions by cloning α9 nAChR subunit cDNA into the pGEMHE vector. Figure 4A is an example of agarose gel electrophoresis for verification of PCR products of the vector (pGEMHE) and insert (α9 nAChR). In each lane, 5 μl of PCR products was loaded. Note that amplification efficiency is the highest for Phusion DNA Polymerases and lowest for Pfu Turbo. We then compared cloning efficiency, in terms of number of colonies produced after transformation, for optimal ratio of vector and insert during Dpn I enzyme digestion and optimal amount of the vector-insert mixture used in transformation. Vectors and inserts were mixed with three different ratios (1:1, 1:2, and 1:4 with a total volume of 16 μl), and incubated at 37°C for 1 h. Different amounts (2, 4, and 8 μl) of the Dpn I-digested mixtures were then added into 40 μl of chemically competent XL-10 Gold E.coli cells for transformation. The number of colonies that grew on each plate was counted next day. Figure 4B shows the number of colonies that resulted from different DNA polymerases, vector to insert ratios (1:1, 1:2, and 1:4), and the amount of vector-insert mixture (2 μl, 4 μl, and 8 μl) used for transformation. The results suggest that the best combination for colony formation is Phusion DNA polymerase amplification with 1:1 vector/insert ratio in Dpn I digestion and 2 μl of vector-insert mix for transformation. Four colonies from the plates resulting from each DNA polymerase were picked up for DNA mini-prep. The PCR confirmation of the insert in the target vector (Figure 4C ) was performed with the GoTaq DNA polymerase (Promega Corporation, Madison, WI) using the pGEMHE vector specific primer pair. Our results indicated that > 90% colonies (11 out of 12) were positive. We have also compared colony production efficiency for three different numbers of PCR cycle (12, 18, and 24 cycles) and three different durations (1, 2, and 4 hours) of incubation time for 37°C Dpn I digestion with α9 nAChR cDNA cloned into pGEMHE and transformed into NEB 10-beta high efficiency competent E coli . The result shows that the 18-cycle PCR amplification of vector and insert produced many more colonies than the 12-cycle and 24-cycle amplifications (31, 302, 43 colonies for 12, 18, and 24 cycles respectively). For incubation duration in Dpn I digestion, 1, 2, and 4 hours of incubation had similar colony production efficiency with colony numbers of 281, 263, and 285 for 1, 2, and 4 hour-incubations, respectively). However, shorter incubation time is not recommended. In our QuickChange mutagenesis experiment, 30 min incubation in Dpn I digestion could not adequately remove the wild-type template background and resulted in a significantly higher fraction of wild type constructs.

Optimization of cloning conditions . (A) PCR amplification of a target cDNA (human nAChR α9 subunit) and the pGEMHE vector using different DNA polymerases: Pfu Turbo, PfuUltra and Phusion. (B) Comparison of number of colonies grown on the plates after transformation. Three different vector-to-insert ratios (1:1, 1:2, and 1:4) during Dpn I digestion and three amounts of vector-insert mixtures (2, 4, and 8 μl) for transformation were tested. See text for details. (C) Clone validation by PCR using GoTaq DNA polymerase. Lanes 1 to 12: target clones to be validated; Lane 13: 1 Kb plus DNA ladder; Lane 14: pGEMHE vector control; Lane 15: negative control using pCR4-TOPO-α9 parent plasmid. (D) Clone validation by restriction digestion to exclude unusual constructs. Lane 1: 1 Kb plus DNA ladder, Lanes 2-11: target clones double digested with Kpn I and Nhe I. Note that this digestion resulted in a pGEM vector and an insert with α9 nAChR plus the 5'UTR and 3'UTR of Xenopus β-globin.

In addition, the PCR-based method could produce some unusual constructs. In our > 10-year practice with the QuickChange site-directed mutagenesis, we only encountered an unusual recombination once. With our FastCloning method in the past few months, we have not seen any unusual construct yet. Thus, if our cloning method produces unusual constructs, their occurrence must be very low. Figure 4D is an example of using restriction digestion to screen clones to exclude unusual constructs. It would be a good practice to perform such a screening for all resulting clones before DNA sequencing. With the optimized cloning conditions (Phusion DNA polymerase, 1:1 vector/insert ratio, 18 cycle PCR amplification, 2 μl of vector-insert mix for transformation, and 1 hour Dpn I digestion), we have also successfully cloned other cDNAs into the target vectors, and obtained positive clones at efficiency ranging from 43% to 100%, with an overall efficiency > 70%. The results further validate the versatility and reliability of this new technique.

Because the PCR amplification of vector can be controlled by primers to exact positions, our FastCloning method is truly sequence-independent. Thus, one can put an insert to any position and in any frame. This feature, although with only a small modification of standard cloning protocol, makes it easy to construct cDNAs for fusion proteins or chimeras. Furthermore, a minor variation of this technique can be applied for insertion of a short DNA fragment directly from two relative long primers for PCR amplification of a cDNA along with its vector. As a proof of principle, we have successfully created human nAChR cDNAs encoding β2-β4 chimeric subunit proteins with C-terminal domain swap between the β2 and β4 subunits. We have also successfully inserted DNA fragments into cDNAs using two long primers with a 16-base overlapping region to directly amplify the cDNA. Figure 5 is an example of using a synthesized insert (99 bp), a DNA sequence encoding a fragment of an acetylcholine binding protein (AChBP), to replace a DNA fragment in the cDNA of the human α7 nAChR for chimera construction. Note that each primer contains only slightly more than half of the insert. The primer pairs have a 16-base overlap region. Using similar method, we have successfully replaced other four DNA fragments (105 bp, 90 bp, 69 pb, and 87 pb) in human α7 nAChR with synthetic AChBP fragments and swapped the DNA sequences (66 bp) encoding the second transmembrane domains of human α4 and β2 nAChR subunits.

An example of chimera construction with a short (105 bp) DNA fragment replaced by a synthesized insert of 99 bp (included in two primers) . (A) A region of α7 nAChR subunit sequence with a 105 bp fragment (blue) to be replaced. (B) Amino acid sequence alignment of human α7 nAChR and corresponding region of Aplysia californica AChBP. Segments to be replaced are colored with blue (35 codons) for α7 nAChR sequence and red (33 codons) for AChBP sequence. ( C ) Target chimera construct with the human nAChR α7 subunit sequence (black) and a substituted 99 bp DNA fragment (red) from the mammalian codon-optimized homologous sequence of the AChBP. Two colored arrows indicate two primers with a 15 bp overlapping region (highlighted). Note that the entire insert of the 99 bp fragment is included in the two primers. Thus, there is no need to amplify the insert. The length of each primer is 81 bases for the forward primer and 76 bases for the reverse primer.

Making multiple mutations in a stretch of DNA is equivalent to making short insertions. Figure 6 is an example of 8 amino acid substitutions (8 arginines to 8 glutamines) with multiple nucleotide mutations spanning a 13-codon region in the cDNA encoding human nAChR β2 subunit. With this relatively short primer pair (45-base forward primer and 46-base reverse primer), we have successfully obtained this multiple mutations spanning a 13-codon stretch in the cDNA. In addition, we have successfully made a quintuple mutant (5 consecutive prolines mutated to 5 alanines) of human 5-HT 3 A receptor subunit with a 31-base forward primer and 35-base reverse primer.

An example of making multiple mutations across a wide region . (A) A fragment of the human nAChR α2 subunit cDNA with 8 arginine (R, in bold) codons (top) to be substituted by glutamine (Q) codons (bottom). Mutated nucleotides are indicated in red. (B) Actual forward (45-base) and reverse (46-base) primers with 16 bp overlapping in their 5' ends.

It is important to be aware that with PCR-based cloning, the synthesized primers may not be completely uniform with correct sequences. Random single-base deletions, mutations, or insertions can occur in a small fraction of primers, which result in unwanted deletion/mutation/insertion in a small fraction (< 5%) of clones. This is especially true for short insertion with long primers. Thus, sequencing across the primer region is required for ultimate confirmation of each clone. If random mutation happens in one clone, picking up another clone often solve the problem. The random mutation out of primer region is rare with high fidelity DNA polymerases. However, DNA sequencing of the entire coding region is still necessary. In the past 8 months, we have successfully used our new cloning method to obtain 21 constructs. Among the 63 sequenced clones, we found 2 deletions in 2 chimeras constructed with long primer pairs, but found no mutations in the entire regions of all constructs beyond primers. Finally, the cloning efficiency of our FastCloning method is high. Of the 21 cloning constructs, we obtained 19 desired constructs in single runs. In 2 experiments, we needed to repeat the procedure to get the final clones. With only one additional construct, we have not obtained final clones after two attempts.

We have developed a highly simplified and robust PCR cloning technique termed FastCloning. The new technique eliminates the need of PCR purification/gel purification kit and cloning kit. It is ligation-independent and does not require specific sequence in the vector. Thus, one can insert a DNA fragment into a vector at any desired position without considering restriction sites. This feature also makes it extremely easy to make constructs for fusion proteins and chimeras. In addition, it can be used to make short insertions and multiple mutations spanning a wide region (up to 120 bp) in a cDNA. Finally, it is a highly efficient and reproducible method.

Abbreviations

polymerase chain reaction

nicotinic acetylcholine receptor

acetylcholine binding protein

gene name of the human nAChR α9 subunit

gene name of the human nAChR α4 subunit

gene name of the human nAChR β2 subunit

serotonin receptor type 3 A subunit

the gene name for human 5-HT 3 A receptor subunit

endothelial cell specific molecule 2 (also named ECSCR: endothelial cell-specific chemotaxis regulator)

protein kinase B gamma variant 1 or 2

gene name of the human zinc activated cation channel

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Acknowledgements

We thank Dr. Alan Gibson from the Barrow Neurological Institute for proofreading the manuscript. This work was supported by National Institute of Health (R01GM085237, to YC) and Barrow Neurological Foundation (to YC). This paper is subject to the NIH Public Access Policy.

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Chaokun Li, Aiyun Wen, Benchang Shen & Yongchang Chang

Department of Obstetrics and Gynecology, St. Joseph's Hospital and Medical Center, Phoenix, AZ, USA

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Department of Genetics and Cell Biology, Guangzhou Medical University, Guangzhou, China

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YC designed the experiment and most primers, and wrote the manuscript. CL performed cloning of human α9 and β2 nAChR subunits into pGEMHE vector, and optimized experimental conditions. He also cloned Akt3v1 or Akt3v2 into pLXSN vector and made a multiple mutation (8 amino acid residue substitutions spanning 13 codons) and several chimeras. AW performed cloning of human 5-HT3A receptor subunit and APBB1 into pGEMHE vector and made a quintuple mutant (5 consecutive mutations) in human 5-HT3A subunit. BS performed cloning of human α4 nicotinic receptor subunit and ECSM2 into pGEMHE vector. JL and YH designed primers for ECSM2 and cloned the ECSM2 into p3xFLAG-CMV-14 vector. YH also contributed to manuscript writing and revision. All authors read and approved the final manuscript.

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Li, C., Wen, A., Shen, B. et al. FastCloning: a highly simplified, purification-free, sequence- and ligation-independent PCR cloning method. BMC Biotechnol 11 , 92 (2011). https://doi.org/10.1186/1472-6750-11-92

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DOI : https://doi.org/10.1186/1472-6750-11-92

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ORIGINAL RESEARCH article

Nimble cloning: a simple, versatile, and efficient system for standardized molecular cloning.

• Key Laboratory of Biology and Genetic Resources of Tropical Crops, Ministry of Agriculture, Institute of Tropical Bioscience and Biotechnology, Chinese Academy of Tropical Agricultural Sciences, Haikou, China

Molecular cloning is one of the most fundamental technologies in molecular biology, and has been critical for driving biotechnological advances. In this study, we have developed a novel method for standardized molecular cloning. The cloning technique known as “Nimble Cloning” uses the restriction enzyme, Sfi I, in combination with the T5 exonuclease, to linearize the vector and generate 3′-overhangs simultaneously. Both PCR products and plasmids can be used for the cloning reaction in the Nimble Cloning system. The cloning system is highly efficient, suitable for gene expression in both prokaryotic and eukaryotic expression systems, and enables the reuse of DNA fragments or plasmid entry clones. Nimble Cloning is applicable for the cloning of single or multiple fragments, as well as multi-site cloning. Due also to its simplicity and versatility, the cloning method has great potential for the modular assembly of DNA constructs.

Introduction

Molecular cloning, which is one of the most fundamental procedures available for modern molecular biology research, has been critical for driving biotechnological advances. One of the main objectives in the post-genomics era is to functionally map gene expression data. Thus, developing methods for the rapid and efficient construction of various vectors for transgenic research is more critical now than ever before.

To the best of our knowledge, of the many molecular cloning protocols that have been developed, the following are the main techniques currently used for routine cloning: restriction digestion- and ligation-based cloning ( Cohen et al., 1973 ), Gateway cloning ( Hartley et al., 2000 ), Gibson assembly ( Gibson et al., 2009 ), and Golden Gate cloning ( Engler et al., 2008 ). Each of these methods have specific limitations ( Liang et al., 2017 ). Traditionally, type II restriction endonucleases and DNA ligases have been used to construct recombinant plasmids, and these enzymes are still extensively used for diverse molecular biology applications. However, this method is laborious and time-consuming and is often limited by the relatively few available restriction enzyme sites, especially during the assembly of complex plasmids from multiple elements ( Lampropoulos et al., 2013 ; Wang et al., 2014 ). Gateway cloning is a very popular site-specific recombination system that exploits the lambda phage integration and excision mechanism ( Hartley et al., 2000 ). Although Gateway cloning has been widely used in many experimental systems, the following are the four main disadvantages with this method: (i) the associated two-step (BP and LR recombination reactions) cloning process is labor-intensive and time-consuming; (ii) the recombination site leaves a 25-bp unwanted junk sequence (scar); (iii) the assembly of multiple fragments is relatively inefficient; and (iv) the commercial enzyme mixes available for this method are expensive, especially for laboratories in developing countries. As an alternative, Gibson assembly is a quick and easy method that enables the linking of multiple overlapping DNA fragments in a single isothermal reaction ( Gibson et al., 2009 ). The recombination is based on the homologous ends (15–20 bp). Other in vitro recombination methods may involve the same homologous ends, including the In-Fusion ( Zhu et al., 2007 ; Sleight et al., 2010 ), Gibson-derived Hot Fusion ( Fu et al., 2014 ), and TEDA ( Xia et al., 2018 ) procedures. These methods are mostly sequence-independent, and because of this sequence flexibility, there is currently no standard protocol for designing the overlapping sequences. Additionally, Gibson assembly requires a destination vector that is linearized by enzyme digestion or PCR, making it a method that is not totally free of restriction enzymes. Golden Gate cloning relies on the type IIs restriction enzymes, and is capable of assembling numerous fragments with high efficiency and fidelity ( Engler et al., 2009 ; Engler and Marillonnet, 2011 ), but the DNA fragment to be assembled needs to be free of the recognition sequence of the enzymes used. Type IIs restriction sites are often shorter than 7 bp and are frequently present within DNA sequences to be cloned, thereby limiting the application of Golden Gate cloning, especially for the cloning of long DNA fragments and multiple DNA fragments.

In this study, we have developed a novel method for standardized molecular cloning known as Nimble Cloning. This method, which is based on the Gibson assembly technique ( Gibson et al., 2009 ), requires a simple enzyme mix of the restriction enzyme, Sfi I, and T5 exonuclease. These two enzymes simultaneously mediate the linearization of the plasmid vector and the generation of 3′-overhangs of the insert DNA fragments. We demonstrate that this cloning method is simple and efficient, and has great potential as a versatile tool for assembling DNA constructs.

Materials and Methods

Strains, reagents, and cell cultivation.

Escherichia coli strains DH5α and DB3.1 (Transgen Biotech, Beijing, China) were used for cloning. Agrobacterium tumefaciens strain GV3101 was used for transforming plants. The E. coli cultures were grown at 37°C in Lysogeny Broth (LB) selection medium. The Sfi I, T5 exonuclease, and Phusion polymerase enzymes as well as the Gibson Mix were obtained from New England Biolabs (Ipswich, MA, USA). Primers were ordered from ThermoFisher (Guangzhou, China).

Plasmid Construction

The destination and entry vectors of the Nimble Cloning system were generated by inserting their cloning cassettes into the desired vector via Gibson assembly. The cloning cassette for the destination vector was “adapter 1– Sfi I site– ccdB gene– Sfi I site–adapter 2” (NC frame), whereas that for the entry vector was “ Sfi I site–adapter 1–XcmI– ccdB gene–XcmI–adapter 2– Sfi I site.” Details regarding the primers used to construct these vectors are listed in Supplementary Table S1 . The “ Sfi I–adapter 1–XcmI– ccdB gene–XcmI–adapter 2– Sfi I” fragment was inserted into the cloning sites of the pENTR/D-TOPO (Invitrogen) and pMD-18T (Takara, Dalian, China) vectors to produce the entry vectors with kanamycin and ampicillin resistance genes, respectively.

The pNC-UC destination vector was constructed by replacing the lacZ sequence of pUC19 with the NC frame. The prokaryotic expression vector pNC-ET28 was constructed by inserting the NC frame into pET28 between the NdeI and XhoI sites. The plant expression vector pNC-Cam1304 was constructed by inserting the NC frame into pCAMBIA1304 between the NcoI and PmlI sites, which replaced the GFP/GUS sequences. The pNC-Green vector was constructed by inserting the NC frame into pGreenII 0000 ( Hellens et al., 2000 ) between the HindIII and EcoRI sites. The NC-frame and GFP, and GFP and NC-frame fragments were cloned into pGreenII-35S ( Tuo et al., 2017 ) between the 35S promoter and the CaMV polyA terminator to generate the plant sublocation vectors pNC-GFP-C and pNC-GFP-N, respectively. The plant RNAi vector pNC-RNAi was constructed by inserting the NC frame, Pdk intron ( Yan et al., 2012 ), and the inverted NC frame into pCAMBIA1304 between the NcoI and PmlI sites. To construct the double open reading frame (ORF) expression BiFC vector pNC-BiFC, the NC frame was first inserted into pSAT-nEYFP-N1 and pSAT-cEYFP-N1 ( Citovsky et al., 2006 ) between BglII and BamHI. The two expression cassettes, including the promoter and the terminator, were then amplified and cloned into pGreen 0029 ( Hellens et al., 2000 ) between the HindIII and EcoRI sites. The maps of the destination vectors were listed in Supplementary Figure S1 .

Preparation of Nimble Mix

A 500-μl sample of 2× Nimble Mix was prepared by mixing 200 μl 5× Nimble buffer (25% PEG-8000, 0.5 M Tris, pH 7.5, 50 mM MgCl 2 , and 50 mM DTT), 4 μl T5 exonuclease (1 U/μl), 40 μl Sfi I (2 U/μl), and 256 μl ddH 2 O. The Nimble Mix was divided into 50–100 μl aliquots in 1.5-ml microcentrifuge tubes and stored at −20°C.

Nimble Cloning Reaction

We added 20–100 ng circular destination vector (1–2 μl) and 10–50 ng PCR insert or entry clone plasmid (1–3 μl) to a PCR microtube containing 5 μl 2× Nimble Mix, after which distilled water was added to the tube for a final volume of 10 μl. Multiple fragments were assembled with 1 μl each insert (10–30 ng). Tubes were incubated in a water bath or a thermocycler for 1 h at 50°C, and the reaction mixture was subsequently used for a transformation or stored at −20°C if not immediately used ( Supplementary Methods S1 ).

Escherichia coli Transformation

We added 2 μl Nimble reaction mixture to 50 μl E. coli DH5α competent cells in tubes, which were then incubated on ice for 30 min, heated at 42°C in a water bath for 45 s, and cooled on ice for 2 min. Next, 450 μl LB medium was added to the tubes, which were then incubated at 37°C for 1 h, with shaking at 200 rpm. An 80-μl aliquot of each cell suspension was spread evenly on agar-solidified LB medium supplemented with specific antibiotics for screening.

Nicotiana benthamiana Transient Expression

Wild-type Nicotiana benthamiana was used to analyze transient expression following an agroinfiltration step that was completed according to a slightly modified version of a published procedure ( Sparkes et al., 2006 , Yan et al., 2012 ). Plasmids were inserted into A. tumefaciens strain GV3101 cells by electroporation. A single colony was then used to inoculate 5 mL YEP medium (10 g/l Bacto-Tryptone, 10 g/l yeast extract, and 5 g/l NaCl, pH 7.0) supplemented with 50 mg/l rifampicin and 50 mg/l kanamycin. Bacteria were grown overnight at 28°C, with shaking at 200 rpm, for an optical density at 600 nm (OD 600 ) of 1.0–1.5. The cultures were centrifuged at 2,000 × g for 5 min, after which the pelleted cells were diluted with infiltration buffer (50 mM MES, pH 5.6, 10 mM MgCl 2 , and 100 mM acetosyringone) for a final OD 600 of 0.2–0.4. The cell solutions were incubated for 1–2 h at 25°C in darkness before being used for an agroinfiltration of N. benthamiana leaves with a 1-ml needleless syringe. The GFP fluorescence was observed with a FluoView FV1000 confocal microscope (Olympus, Japan) or a UV lamp (UVP, Upland, CA, USA).

Design of the Nimble Cloning System

The Nimble Cloning system includes recipient expression (destination) vectors and entry vectors. All of the destination vectors contain the NC frame, which comprises the “adapter 1– Sfi I– ccdB gene– Sfi I–adapter 2” sequence in the recombination site. In contrast, the entry vectors contain the “ Sfi I–adapter 1–XcmI– ccdB gene–XcmI–adapter 2– Sfi I” sequence in the cloning site, with kanamycin or ampicillin resistance genes. During a Nimble Cloning reaction, the entry clone should carry a different resistance gene from that in the destination vector. A DNA fragment can be inserted into the entry vector to form the entry clone via TA cloning, or by Gibson cloning with adapters as overlapping sequences, after the entry vector has been digested with XcmI. The DNA fragment in the entry clone or the PCR product flanked by the adapters can be cloned into the circular destination vector with the Nimble Mix during the Nimble Cloning reaction and the transformation of E coli . The Nimble Mix consists of two enzymes, Sfi I and T5 exonuclease ( Figure 1 ).

Figure 1 . Nimble Cloning system. (A) Schematic of the destination and entry vectors in the Nimble Cloning system. The destination vector contains the NC frame comprising the “adapter 1– Sfi I– ccdB gene– Sfi I–adapter 2” sequence, whereas the entry vector contains the “ Sfi I–adapter 1–XcmI– ccdB gene–XcmI–adapter 2– Sfi I” sequence at the cloning site. (B) Schematic of the Nimble Cloning method. The PCR product flanked by the adapters or the DNA fragment in the entry clone can be cloned into the circular destination vector in a one-step Nimble Cloning reaction. The DNA fragment can be inserted into the entry vector to form the entry clone via TA cloning, or by Gibson cloning using the adapters as overlapping sequences, after the entry vector is digested with XcmI. Nimble Cloning involves Sfi I and T5 exonuclease.

The two unique 21-bp nucleotide sequences that form the NC frame adapters required for recombination were designed based on the criteria to increase the likelihood of accurate assembly and minimize the possibility that the expression levels of the nearby genes are affected ( Torella et al., 2014a ). Thus, we computationally designed adapters 1 and 2 comprising the cagtggtctctgtccagtcct and cggtctcagcagaccacaagt sequences, respectively, and used them for the tests of cloning efficiency and gene expression. These two adapters enable efficient assembly and are appropriate for gene expression, as confirmed in the subsequent subsections.

Cloning of a Single Gene Fragment and Assembly of Multiple DNA Fragments With Nimble Cloning

Nimble Cloning is based on Gibson assembly, and both methods involve the enzyme-catalyzed assembly of overlapping DNA fragments. Gibson assembly generally applies three enzymes, namely T5 exonuclease, Phusion DNA polymerase, and Taq DNA ligase; however, Taq DNA ligase can be removed without decreasing the cloning efficiency ( Fu et al., 2014 ; Benoit et al., 2016 ). The more recently developed TEDA method, which requires only T5 exonuclease, results in efficient cloning ( Xia et al., 2018 ). We assessed the enzyme requirements for Nimble Cloning reactions involving one or multiple DNA fragments. The pNC-UC (20 ng) vector used as the destination vector in each reaction was derived from pUC19, with the NC frame replacing the lacZ sequence. The lacZ gene (10 ng) as well as the 35S promotor, GFP gene, and nos termination sequence (Tnos) (10 ng) were used for the cloning of single and multiple DNA fragments, respectively ( Figures 2A,B ). For multiple-fragment cloning, the junctions of the internal fragment do not use the unique adapters, but fragment-specific sequences ( Supplementary Table S1 ). The reaction was completed at 50°C for 1 h. The results indicated that Nimble Cloning with Sfi I and T5 exonuclease worked well for the cloning of single and multiple DNA fragments. Moreover, the cloning efficiency was higher than that of Gibson assembly, which involves the linearized pNC-UC vector and three enzymes. All reactions for the cloning of lacZ yielded >99% positive clones (i.e., blue colonies) on LB plates supplemented with X-gal and IPTG. For the cloning of three fragments, 24 colonies in each group were tested by colony PCR and > 95% had the correct band. Twelve colonies resulting from the Nimble Cloning of single and multiple DNA fragments were further analyzed by sequencing, and no errors were detected.

Figure 2 . Optimization of Nimble Cloning. Effects of enzymes and buffer components (A,B) , reaction temperatures (C,D) , and reaction times (E,F) on the efficiency and fidelity. The pNC-UC vector was used as the destination vector in each reaction. The lacZ fragment was used for the cloning of a single fragment, whereas the 35S promotor, GFP gene, and Tnos fragments were used for the cloning of multiple fragments (A,B,E,F) . The PCR product and the entry clone with lacZ as well as the lacZ PCR product without the unique adapter sequences (scarless) were used in (C,D) . * Gibson assembly with the linearized pNC-UC vector. Fidelity was analyzed based on the ratio of blue colonies to total colonies for the cloning of a single fragment, and the ratio of PCR-positive colonies to tested colonies ( n = 24) for the cloning of multiple fragments. Data are presented as the average (and standard deviation) of three parallel experiments.

Optimal Reaction Temperature and Duration for Nimble Cloning

To determine the optimal reaction temperature, six different temperatures (25, 30, 37, 42, 55, and 55°C) were tested for the standardized cloning (with unique adapters for the PCR product and entry clone) and scarless cloning (without unique adapters) methods ( Figures 2C,D ). The results following a 1-h reaction revealed that the 37–50°C range was appropriate for the standardized cloning with a PCR product and the scarless cloning. However, 50°C was optimal for the standardized cloning with an entry clone.

Six different time points (5, 15, 30, 60, 90, and 120 min) were tested for the cloning of one or three fragments ( Figures 2E,F ) at 50°C. The data indicated that the efficiency and fidelity of the cloning of a single fragment were high even for the shortest time point (5 min), and the efficiency plateaued after 30 min. Regarding the cloning of three fragments, the efficiency and fidelity increased over time and plateaued after 60 min. Three PCR-negative colonies in the three-fragment Nimble Cloning with the reaction time of 5 min were sequenced, and they all were the products of self-ligation of the vector between the two Sfi I sties of NC frame with the overlap sequences “GGCC.”

Application of Nimble Cloning for Prokaryotic and Plant Expression Systems

Unique adapters are used in the Nimble Cloning system for standardized cloning reactions. To confirm that unique adapters do not adversely affect gene expression, we evaluated the expression of sequences from the plasmids constructed by standardized cloning (with unique adapters) and scarless cloning (without unique adapters) in both prokaryotic and plant expression systems.

The pNC-ET28 vector, which is derived from pET28, was used for the prokaryotic expression analysis. The GFP gene was cloned into pNC-ET28 by standardized cloning and scarless cloning to form pNC-ET28-GFP and pNC-ET28-GFP-SL, respectively ( Figures 3A,B ). The plasmids were transformed into E. coli cells, after which GFP fluorescence was observed under a bright light ( Figure 3C ) and a UV lamp ( Figure 3E ). Additionally, GFP was detected in a western blot involving the anti-GFP antibody ( Figure 3D ). There were no differences between pNC-ET28-GFP and pNC-ET28-GFP-SL regarding the resulting protein abundance and activity.

Figure 3 . Nimble Cloning was applied for a prokaryotic expression system. (A) Schematic of the Nimble Cloning system vector pNC-ET28, which is derived from pET28. The primers with the unique adapters as overlapping sequences were applied for standardized cloning, whereas the primers with the sequences outside the adapters as overlapping sequences were applied for scarless (SL) cloning. (B) The sequences around the insertion sites of pNC-ET28-GFP and pNC-ET28-GFP-SL. Unique adapters were used for standardized cloning to generate the pNC-ET28-GFP, while the underline sequences outside the adapters were used as the homologous ends for scarless cloning to generate the pNC-ET28-GFP-SL. (C–E) The GFP fluorescence associated with the pNC-ET28-GFP, pNC-ET28-GFP-SL, and empty pET28 vector was observed under a bright light (C) and a UV lamp (E) . Additionally, GFP was detected in a western blot with an anti-GFP antibody (D) .

For the plant expression analysis, strong GFP fluorescence was observed for plants regardless of whether they were transformed with pCAMBIA1304-GFP or pNC-Cam1304-GFP, which were constructed by Gibson assembly and standardized Nimble Cloning, respectively, with no significant difference between the two plasmids ( Figures 4A,B ). The pNC-Green-GFP plasmid constructed by Nimble Cloning for multiple fragments (35S promotor, GFP, and Tnos) also resulted in a strong GFP signal in N. benthamiana leaves ( Figures 4A,B ). Fusion protein production was assessed with sublocation vectors. The papaya eIFiso4E gene, which encodes a protein that is present in the nucleus and cytoplasm, as well as the papaya ringspot virus VPg gene, which encodes a protein (prsv-VPg) present in the nucleus, were cloned into the sublocation vectors at the N- and C-termini of the GFP gene with the standard and scarless cloning methods of the Nimble Cloning system ( Figures 4A,C,D ). All of the sublocation constructs were expressed in N. benthamiana leaves, and there were no differences between the constructs resulting from the standardized and scarless cloning methods.

Figure 4 . Nimble Cloning was applied for a plant expression system. (A) Schematic of the Nimble Cloning system vectors used for gene overexpression in plants. NC, NC frame for Nimble Cloning. (B–D) The GFP signal in Nicotiana benthamiana leaves was observed with a confocal microscope or a UV lamp. (B) Comparison of the GFP signal associated with pCAMBIA1304-GFP and pNC-Cam1304-GFP, and the expression of pNC-Green-GFP. Comparison of the expression of papaya eIFiso4E (C) and prsv-VPg (D) resulting from different sublocation vectors of the Nimble Cloning system with or without (SL) unique adapters. Papaya eIFiso4E is present in the nucleus and cytoplasm, whereas prsv-VPg is present in the nucleus. Scale bars = 50 μm.

Nimble Cloning for Multiple-Site Cloning

RNA interference (RNAi) constructs contain inverted DNA repeats that express self-complementary RNA sequences. We constructed a plant RNAi vector, pNC-RNAi, consisting of an “NC frame–Pdk intron–NC frame” cassette between the CaMV 35S promoter and nos terminator of pCAMBIA1304. The second NC frame was inverted, which enabled a single PCR product amplified with primers containing the unique adapter sequences to be simultaneously recombined into the vector in the sense and antisense orientations to form RNAi constructs via one-step Nimble Cloning ( Figure 5A ). A 337-bp fragment of the marker gene EGFP was amplified and then ligated to the pNC-RNAi vector with the Nimble Mix during a Nimble Cloning reaction. After the subsequent transformation, 12 colonies were analyzed in a PCR assay to confirm the amplification of the expected band. Six colonies were sequenced, and no errors were detected. We then tested the utility of pNC-RNAi-GFP for gene silencing via A. tumefaciens -mediated transient expression. The A. tumefaciens cells carrying pBIN19-GFP ( Santos-Rosa et al., 2008 ) were mixed with A. tumefaciens cells containing an empty pNC-RNAi vector (control) or the RNAi construct pNC-RNAi-GFP. The cell mixtures then infiltrated different parts of the same N. benthamiana leaves. The leaf areas infiltrated with the control cells produced a strong GFP signal at 3 days post-agroinfiltration, whereas considerably weaker signals were detected for the leaf areas infiltrated with the cells carrying the RNAi construct ( Figure 5B ).

Figure 5 . Nimble Cloning for multiple-site cloning in plant RNAi and BiFC. (A) Schematic of hpRNA construction by a one-step Nimble Cloning method. A single DNA fragment can be simultaneously cloned into the pNC-RNAi vector in the sense and antisense orientations. (B) Silencing of the GFP gene. Agrobacterium tumefaciens cells containing pBIN19-GFP were mixed with A. tumefaciens cells containing an empty pNC-RNAi vector (left) or the hpRNA construct pNC-RNAi-GFP (right). The cell mixtures infiltrated different parts of the same Nicotiana benthamiana leaves. (C) Schematic of the construction of a double ORF expression BiFC vector by Nimble Cloning. Two genes can be cloned into the vector in a single reaction. NC 1, NC frame 1; NC 2, NC frame 2. (D) The double ORF expression BiFC vector containing the genes encoding papaya eIFiso4E and prsv-VPg was infiltrated into N. benthamiana leaves. Fluorescence was observed by confocal microscopy. Scale bars = 20 μm.

A double ORF expression BiFC (bimolecular fluorescence complementation) system enables the coexpression of two fluorescent protein fragments from a single vector ( Grefen and Blatt, 2012 ; Gookin and Assmann, 2014 ). The double ORF expression vector (pNC-BiFC) we constructed for a plant BiFC assay consisted of two NC frames with different adapters. Two genes of interest flanked by the adapters were simultaneously recombined into the vector by Nimble Cloning ( Figure 5C ). The genes encoding eIFiso4E and prsv-Vpg were amplified with primers containing the adapters in NC frames 1 and 2, respectively. The PCR products were inserted into the pNC-BiFC vector during a Nimble Cloning reaction. A PCR analysis of the transformants revealed that all 12 tested colonies produced the expected band, with no sequence errors in the six colonies analyzed by sequencing. An A. tumefaciens -mediated transient expression experiment was used to evaluate the expression of the plasmid sequences. The A. tumefaciens cells containing pNC-BiFC-eIFiso4E/Vpg infiltrated N. benthamiana leaves, after which the infiltration areas were observed by confocal microscopy at 3 days post-agroinfiltration. The EYFP fluorescent signal was observed exclusively in the nucleus ( Figure 5D ). Additionally, fluorescence was undetectable for the negative control leaves that were infiltrated with cells carrying the empty pNC-BiFC vector (data not shown).

The Advantages of Nimble Cloning Over Other Methods

We herein describe the development of a novel method, Nimble Cloning, which is useful for simple and standardized molecular cloning. This new method has several advantages over the widely used Gibson assembly and Gateway cloning techniques. First, unlike Gibson assembly, which requires an additional step to linearize the destination vector, Nimble Cloning uses circular plasmids. The Sfi I and T5 exonuclease enzymes in Nimble Mix simultaneously mediate the linearization of the vector and the generation of 3′-overhangs. Second, Gibson assembly currently lacks a standard primer design method. The Nimble Cloning system involves unique nucleotide sequences (adapters) for standardized cloning, enabling a DNA sequence flanked by the adapters to be cloned into any Nimble Cloning vector. Third, Gibson assembly is limited to PCR products as inserts, and Gateway cloning requires entry clones. In contrast, Nimble Cloning is more flexible because it is appropriate for both PCR products and entry clones. Fourth, Gateway cloning leaves a 25-bp scar sequence at the recombination site, while Nimble Cloning can be applied for scarless cloning, if necessary. T5 exonuclease is not only an exonuclease, but also a flap endonuclease ( AlMalki et al., 2016 ; Xia et al., 2018 ). It can degrade linear dsDNA or ssDNA by the activity of dsDNA specific exonuclease and ssDNA endonuclease. The Sfi I sequence as well as the 21-bp adapters can be degraded by T5 exonuclease for scarless cloning in Nimble Cloning. Finally, Nimble Cloning is relatively inexpensive because it only requires Sfi I and T5 exonuclease. These advantages may make Nimble Cloning an ideal system for molecular cloning, especially for researchers in developing countries.

The recombination mechanism underlying the Nimble Cloning system is similar to that of Gibson assembly, and is mainly based on the T5 exonuclease activity that yields the single-stranded DNA overhangs. The main difference between these two methods is that Gibson assembly requires a linearized destination vector, while Nimble Cloning uses circular plasmids. This is because Nimble Cloning involves a mixture of Sfi I and T5 exonuclease, which mediate the linearization of the vector and the recombination reaction at the same time. Our results indicated that Nimble Cloning is not only more convenient than Gibson assembly, but it also results in a higher cloning efficiency ( Figure 2A ). Previous studies confirmed that Golden Gate cloning, which also involves a one-step digestion/ligation reaction, is more efficient than traditional cloning methods requiring a two-step digestion and ligation reaction ( Engler et al., 2009 ; Hsu and Smanski, 2018 ). The increased efficiency of one-step cloning may be due to the decrease in the number of steps involving the DNA. The manipulation and modification of DNA (e.g., digestion, extraction, column purification, and buffer exchange) are likely to result in DNA damages and a decrease in DNA yield ( Engler et al., 2009 ). The Nimble Cloning vectors are not pre-digested, but are simply added to the reaction mix without the need for a purification step. In addition, removing the Phusion and dNTPs also resulted in increased efficiency ( Figure 2A ) ( Xia et al., 2018 ).

Both Nimble Cloning and Golden Gate cloning include restriction enzymes in their reaction mixes. Golden Gate cloning uses type IIs restriction enzymes, such as BsaI and BpiI, whereas Nimble Cloning applies a relatively rare-cutting restriction enzyme, Sfi I. Specifically, the Sfi I recognition sites occur at low frequencies in the A. thaliana genome, with only three sites per million base pairs, which is in contrast to the 265 BsaI sites per million base pairs ( Ghareeb et al., 2016 ). This provides a clear advantage of Nimble Cloning over Golden Gate cloning. Furthermore, if there are naturally occurring restriction sites in the DNA fragment of interest, Golden Gate cloning often requires site-directed mutagenesis to eliminate them, which is laborious and possibly ineffective. With Nimble cloning, this issue can be circumvented by first linearizing the destination vector with Sfi I followed by the steps of Gibson assembly to clone the DNA fragments of interest. Thus, the occasional presence of the Sfi I recognition site will not limit the utility of Nimble Cloning.

The Unique Nucleotide Sequences and System Vectors Enabled Standardized Molecular Cloning

In vitro recombination methods based on short homologous ends, including Gibson assembly and In-Fusion, are becoming popular in molecular cloning research. These methods are mostly sequence-independent, and because of their flexibility, a standardized method for designing the overlapping sequences has not been proposed. To facilitate standardized molecular cloning, two unique nucleotide sequences were designed for the Nimble Cloning system. All of the entry and destination vectors of this system contain unique nucleotide sequences. A PCR product flanked by the unique nucleotide sequences can be cloned into any entry or destination vector. We also evaluated the cloning efficiency and the effects of the unique nucleotide sequences on gene expression. We observed that these sequences resulted in highly efficient cloning ( Figures 2A,B ), and were suitable for gene expression ( Figures 3 , 4 ). Thus, these unique sequences may be useful as standard homologous arms (primers) for in vitro recombination-based cloning.

Like Gateway cloning and Golden Gate cloning, Nimble Cloning requires system vectors. We constructed two entry vectors, with ampicillin and kanamycin resistance genes. These entry vectors satisfy the requirements for the cloning of a single fragment. For the cloning of multiple fragments, additional entry vectors will need to be constructed, or the PCR fragments for cloning will need to be used directly. We also constructed more than 40 Nimble Cloning destination vectors, some of which were included in this study. Since there are currently hundreds of expression vectors available for molecular biology research, a viable Nimble Cloning system will require the development of many additional destination vectors. To construct these destination vectors, the NC frame will need to be inserted in the cloning sites ( Supplementary Methods S1 ). Thus, the construction of new destination vectors is simple, and will not limit the application of the Nimble Cloning system. Furthermore, entry and destination vectors form the basis of standardized molecular cloning.

Nimble Cloning May Facilitate Modular DNA Assembly and Multi-Level Construct Assembly

In addition to standardization, modularity is another principle associated with DNA assembly. The main methods currently used for modular DNA assembly are Golden Gate cloning and Gibson assembly. Golden Gate-based methods, including MoClo ( Weber et al., 2011 ), GoldenBraid ( Sarrion-Perdigones et al., 2011 ; Vazquez-Vilar et al., 2017 ), and MetClo ( Lin and O'Callaghan, 2018 ), have been applied to construct complex DNA constructs. Gibson-based methods involve unique nucleotide sequences to facilitate the assembly of sequences in the correct order ( Casini et al., 2014 ; Torella et al., 2014a , b ; Halleran et al., 2018 ). However, Gibson-based methods require linearized DNA modules produced via PCR amplification or enzyme digestion in each assembly level. Nimble Cloning enables the direct use of an entry clone for highly efficient cloning, thereby facilitating modular DNA assembly and multi-level construct assembly.

In summary, Nimble Cloning is simple, versatile, and efficient, and is a promising system for the standardized molecular cloning required in diverse applications. The plant expression vectors of pNC system will facilitate the analysis of gene overexpression, protein localization, gene silencing, and protein-protein interactions in plant functional genomics.

Accession Numbers

Sequence data from this article can be found in the GenBank data libraries under accession numbers: pNC-UC, MK720605; pNC-ET28, MK720606; pNC-Cam1304-35S, MK896896; pNC-Green, MK896901; pNC-GFP-C, MK896905; pNC-GFP-N, MK896906; pNC-RNAi, MK896898; pNC-BiFC, MK896893; pNC-KEn, MN604399; pNC-AEn, MN604400.

Data Availability Statement

The datasets generated for this study can be found in the GenBank Nucleotide Database with accession numbers MK720605 for pNC-UC and MK720606 for pNC-ET28.

Author Contributions

PY and PZ conceived the project and designed experiments. PY, YZ, DT, and XL performed experiments. PY, DT, and WS analysis the data. PY wrote the manuscript. All authors commented on the manuscript.

Innovative Research Team of the Natural Science Foundation of Hainan Province, China (2018CXTD343) and Central Public-interest Scientific Institution Basal Research Fund for Chinese Academy of Tropical Agricultural Sciences (1630052018004, 19CXTD-33).

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

We would like to thank Dr. P. Mestre for providing the Agrobacterium clones containing pBIN19-GFP, and Dr. Stanton B. Gelvin for providing pSATN BiFC vectors.

Supplementary Material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fbioe.2019.00460/full#supplementary-material

Supplementary Figure S1. Maps of plasmids used in this study. (A,B) The prokaryotic expression vectors pNC-UC and pNC-ET28. (C,D) The plant expression vector pNC-Cam1304 and pNC-Green. (E,F) The plant subcellular location vectors pNC-SubC and pNC-SubN. (G) The plant RNAi vector pNC-RNAi. (H) The double ORF expression BiFC vector pNC-BiFC.

Supplementary Table S1. Primer sequences used in this study.

Supplementary Methods S1. Standardized molecular cloning by Nimble Cloning.

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Keywords: molecular cloning, standardized cloning, Gibson assembly-derived cloning, vector construction, DNA assembly, prokaryotic expression, plant gene expression

Citation: Yan P, Zeng Y, Shen W, Tuo D, Li X and Zhou P (2020) Nimble Cloning: A Simple, Versatile, and Efficient System for Standardized Molecular Cloning. Front. Bioeng. Biotechnol. 7:460. doi: 10.3389/fbioe.2019.00460

Received: 19 July 2019; Accepted: 19 December 2019; Published: 15 January 2020.

Reviewed by:

Copyright © 2020 Yan, Zeng, Shen, Tuo, Li and Zhou. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY) . The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

*Correspondence: Pu Yan, yanpu@itbb.org.cn ; Peng Zhou, zhp6301@126.com

Disclaimer: All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.

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Unlocking the Power of Molecular Cloning: Revolutionizing Medical Microbiology Procedures

Author: Neelabh Datta

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The revolutionary realm of molecular cloning, encompassing the creation of recombinant DNA molecules, has ignited a wave of progress within the life sciences. The advent of potent tools has facilitated the manipulation of DNA, resulting in an extraordinary surge in the versatility and breadth of applications in recombinant DNA technology. The once complex task of cloning genes has now been simplified, triggering a veritable explosion of insights into gene functionality. This has been achieved through the seamless fusion of multiple DNA fragments or the utilization of interchangeable gene cassettes, culminating in a state of unparalleled agility and expediency. In the 1970s, when restriction endonucleases, enzymes that cut DNA molecules selectively were discovered, molecular cloning technology has grown exponentially in application and intricacy, resulting in influential DNA manipulation tools. Recent decades have seen an explosion in our understanding of gene function due to the simplicity and efficiency of molecular cloning. It is expected that emerging technologies will offer superior potentials, such as stitching together multiple DNA fragments in under a few hours and transforming the resulting plasmids into bacteria, or the use of swappable genes, which can be easily moved between different paradigms, maximizing promptness and flexibility. It has been proved that cloning techniques provide a gold standard technique for polymicrobial infection, recombinant cytokines, antimicrobial peptides, epidemiology and gene therapy due to the limitations of culture-based methods. Due to molecular cloning technique, recombinant antigens are now being used to monitor patients against clinical infections. As a result of laboratory techniques that permit in vitro chemical synthesis of any DNA construct specified in silico, molecular cloning will likely undergo a paradigm shift in the coming future. As a result of these advances, DNA clones can be constructed faster and iteratively, which will speed up the growth of new vaccines, gene therapy vectors, and recombinant proteins. Here I present a detailed overview of the latest applications of molecular cloning techniques in medical microbiology.

Keywords: molecular cloning, medical microbiology, gene therapy, antimicrobial peptides, recombinant cytokines, epidemiology, bioterrorism, polymicrobial infection

Datta, N., (2024) “Unlocking the Power of Molecular Cloning: Revolutionizing Medical Microbiology Procedures”, University of Michigan Undergraduate Research Journal 17. doi: https://doi.org/10.3998/umurj.5509

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Published on 09 mar 2024, peer reviewed, creative commons attribution-noncommercial-noderivs 4.0, introduction.

Have you ever wondered how tiny organisms like bacteria and viruses can wreak havoc on the human body? Medical microbiology holds the key to unlocking the mysteries of these microscopic villains. Since the dawn of germ theory of disease, medical microbiology has progressed considerably at the end of the last century. It was at this time that premature culture techniques were fruitful in quarantining many pathogens and labeling them with their corresponding causative diseases. Medical microbiology has become an essential component of healthcare and society, thanks to its rapid advancement in response to clinical demands. This field of science investigates the relationship between large and small organisms in both normal and disease conditions, as well as the development of disease and the appropriate treatments that result in complete clinical recovery. A wide range of testing methods are employed to accomplish this task ( Savitskaia, 1993 ). The field of medical microbiology continues to rely heavily on culture-based methods for identifying culture organisms despite substantial developments in analysis methodologies. By identifying the useful microorganisms from those that are pathogenic, medical microbiology contributes to the health of the public and helps to control infectious disease epidemics. Medical microbiology has undergone significant changes in the past few decades, with new methods being developed to make the isolation and detection of various microorganisms more efficient. In medical microbiology laboratories, a range of different microscopy and culture techniques are typically used. One example of a technique that can be used to detect bacteria, fungi, parasites, or viruses in infected cells is indirect fluorescent technique. This technique uses polyclonal antibodies raised in animals like mice or horses, as well as monoclonal antibodies produced using hybridization technology ( Sharma et al., 2014 ). Latex agglutination tests are used to detect particulate antigens, while enzyme immunoassay tests are employed for soluble antigens ( House et al., 2005 ). When it comes to life-threatening infections, it is essential to have quick and accurate diagnostic tests to facilitate prompt antimicrobial therapy ( Sharma et al., 2014 ). In recent times, molecular biology techniques have emerged as a more efficient and rapid way to perform microbiological diagnosis. These methods use nucleic acid probes in conjunction with Polymerase Chain Reaction (PCR) amplification techniques ( Ieven et al., 1996 ). There is a possibility that cloning techniques could be useful in clinical settings, as they are already utilized in different areas of our everyday lives. For instance, during the outbreak of severe acute respiratory syndrome coronavirus-2 (SARS CoV-2), laboratories recognized the need for the swift development of diagnostic tests for emerging diseases that have a significant impact on society. It is vital to quickly transfer such technology to laboratories that perform routine diagnostics. A contemporary issue is that scientific reactions to evolving threats are far more prompt than governmental reactions, and new diagnostic tests for use outside research laboratories often take a long time to be sanctioned ( Raoult et al., 2004 ).

From Past to Present: The Evolution of Molecular Diagnostics

Molecular biology has been a vital tool for the medical laboratory, with its roots dating back to Linus Pauling’s discovery of sickle cell anemia as a “molecular disease” in 1949 ( Pauling et al., 1949 ). However, it was not until recombinant DNA technology in the early days of molecular biology that it became usable in medical diagnostics ( Chehab, 1993 ). Molecular diagnostics grew from basic knowledge on the primary sequence of various genes, with DNA probes incorporating radioactive nucleotides allowing analysis via Southern blotting of genomic regions. This led to the concept and application of restriction fragment length polymorphism (RFLP), which helped track variant alleles in the human genome ( Williams, 1989 ). In 1978, molecular diagnostics techniques were used to make the first prenatal diagnosis of α-thalassemia, and the use of RFLP to characterize sickle cell alleles set the foundation for the characterization of other genetic diseases and infectious diseases using molecular diagnostics platforms ( Kan et al., 1978 ). The mid-1980s brought the development of PCR, which quickly became a staple of laboratory medicine with its ability to exponentially amplify a target sequence and identify known mutations or sequences within hours ( Mullis et al., 1986 ). PCR also established the foundation for many variant detection schemes based on the amplification of DNA. Following the publication of the human genome draft sequence, the challenge to improve existing variant detection technologies to achieve robust, cost-effective, rapid, and high-throughput analysis of genomic variation moved to the forefront of molecular diagnostics. Real-time PCR and its numerous variations, DNA microarray-based genotyping and transcription profiling, microbiome sequencing, proteomics, pharmacogenomics, nutrigenomics, forensic medicine, and CRISPR/Cas9 genome editing all represent important and critical advances in the field ( Hendrix and Rohde, 2021 ). Despite the explosion of diverse variant detection assays, DNA sequencing remains the gold standard for pathogen identification and surveillance, especially with breakthroughs in next-generation sequencing (NGS) technology ( Hendrix and Rohde, 2021 ). However, the costs of initial investment and difficulties in standardization and interpretation of ambiguous results continue to limit the use of NGS in clinical laboratories. Physicians and other healthcare professionals are now working with molecular diagnostics professionals to understand the basis of infectious disease pathology and when to use molecular diagnostics like NGS. One example is the use of 16S in-house assay sequencing to identify bacterial pathogens directly from tissue specimens when culture results are negative, but there is evidence of histopathologic pathogen damage ( Hendrix and Rohde, 2021 ). In addition to PCR and sequencing, molecular cloning also played a crucial role in the development of molecular diagnostics techniques. By enabling the replication of specific DNA sequences, molecular cloning made it possible to generate large amounts of identical DNA fragments for further analysis. This technique is based on the use of restriction enzymes to cut DNA at specific recognition sites and then ligating these fragments into a plasmid vector for amplification and manipulation. The ability to produce recombinant proteins using molecular cloning revolutionized the field of medical research and led to the production of many important biological therapeutics, including insulin, growth hormone, and clotting factors. Molecular cloning also allowed for the creation of genetic probes that could be used to detect specific DNA sequences, such as those associated with infectious agents. This enabled the development of highly sensitive and specific diagnostic tests, such as the PCR-based tests that are widely used today for the detection of viruses like HIV and hepatitis C. Molecular cloning techniques have also facilitated the identification of new disease-causing genes and the development of gene therapy approaches for genetic diseases.

An Overview of Molecular Cloning

In the annals of scientific progress, the field of molecular cloning stands as a testament to human ingenuity and perseverance. Throughout history, researchers have endeavored to unlock the secrets of DNA, seeking ways to manipulate and understand its intricate structure. It is within this context that the technique of molecular cloning emerged, revolutionizing the study of genetics and paving the way for a multitude of biological and technological applications. Molecular cloning (a molecular diagnostics technique) refers to a technique that involves isolating a specific DNA sequence and then amplifying short regions of it in vitro. The roots of molecular cloning can be traced back to the mid-20th century, when scientists began unraveling the mysteries of DNA. It was during this time that the concept of isolating specific DNA sequences and amplifying them in vitro took shape. One commonly employed approach involved the use of restriction enzymes, which acted as molecular scissors, cutting DNA at specific sites. By digesting existing DNA fragments or targeting them through PCR, researchers were able to generate short inserts of DNA, typically around 100 base pairs in length (Juliane and Lessard, 2013 ). These resulting short inserts can also be created as complementary single-stranded fragments that are then annealed to form a double-stranded fragment (Juliane and Lessard, 2013 ). Once the DNA of interest has been obtained, it can be inserted into a vector plasmid, which is a circular double-stranded DNA. These vectors are smaller versions of naturally occurring plasmids and contain features such as replication origins, drug resistance genes, and unique restriction sites that enable the insertion of DNA fragments (Juliane and Lessard, 2013 ). The multiple cloning sites of these vectors usually contain different restriction sites, making it easier to select the appropriate enzymes for a variety of inserts. Molecular cloning was typically used to amplify DNA fragments containing genes, but it can also be used to amplify any DNA sequence, including promoters, non-coding sequences, chemically synthesized oligonucleotides, or fragments of randomly generated DNA ( Sharma et al., 2014 ).

There was a wide range of biological applications and technological applications that make use of this method, including the production of recombinant antigens, cytokines, and proteins ( Nguyen et al., 2004 ). If DNA sequences were to be amplified and cloned in vitro and in vivo, they must be linked to primary sequence elements that are capable of directing their own replication and propagation in the desired target host in conjunction with the linked sequence ( Lu et al., 2008 ). Thus, the inclusion of a host-specific origin of replication and a selectable marker became essential sequence elements. In addition, when selecting a cloning vector, researchers had to consider a number of other characteristics, including the ability to express proteins, tag them, and generate single-stranded RNA and DNA ( Sharma et al., 2014 ). As the field progressed, researchers sought to refine and expedite the cloning process. Recombinase-based cloning emerged as a one-step reaction, allowing for high-throughput cloning by inserting a specific DNA fragment into a specific region of target DNA through the interchange of relevant DNA fragments ( Copeland et al., 2001 ). This streamlined approach facilitated the cloning of any DNA fragment, representing a significant leap forward compared to the classical restriction- and ligation-based approach, which involved fragmenting DNA with restriction endonucleases, ligating it to a vector, transfecting it into host cells, and subsequently screening and selecting the desired clones ( Sharma et al., 2014 ). Cloning procedures generally followed these classical steps; however, a number of unconventional routes can be chosen depending on the specific solicitation.

Cloning procedures had traditionally adhered to the classical steps outlined previously, but the field of molecular cloning continued to evolve, offering researchers a growing repertoire of unconventional routes to choose from, based on the specific goals and requirements of their studies. Some advancements had further expanded the possibilities and enhanced the efficiency of molecular cloning techniques. One notable development in molecular cloning was the emergence of advanced DNA assembly methods that enable the construction of large DNA constructs with precise control over their sequence composition. For instance, techniques such as Gibson assembly, Golden Gate assembly, and ligase cycling reaction (LCR) had revolutionized the process by allowing seamless assembly of multiple DNA fragments in a single reaction ( Engler et al., 2008 ; Gibson et al., 2009 ; Li and Elledge, 2007 ). These methods bypassed the labor-intensive steps of traditional restriction digestion and ligation, streamlining the process and reducing the occurrence of unwanted mutations. Gibson assembly was a method that allowed the seamless assembly of multiple DNA fragments without the need for restriction enzymes or DNA ligases. It operated on the principle of in vitro homologous recombination and utilized three key enzymatic activities: exonuclease, polymerase, and DNA annealing. In this technique, the DNA fragments to be assembled were designed with overlapping regions called “homology arms.” These homology arms enabled the fragments to anneal to one another in the presence of the exonuclease and polymerase enzymes, which trimmed back the ends of the fragments and filled in the gaps, respectively. The resulting annealed fragments were then extended and ligated, producing a seamless composite DNA construct ( Gibson et al., 2009 ). Gibson assembly offered several advantages, including its simplicity, efficiency, and the ability to seamlessly assemble multiple fragments with high fidelity, minimizing the introduction of errors. Golden Gate assembly was another powerful technique used for the modular assembly of DNA fragments. It relied on the activity of type IIS restriction enzymes, such as BsaI, which cut DNA sequences outside their recognition sites. The DNA fragments to be assembled were designed with specific recognition sequences for the type IIS enzyme at their ends, along with overlapping regions. During the assembly process, the type IIS restriction enzyme cuts the DNA fragments at the specific recognition sites, generating cohesive ends. The cohesive ends from different fragments were then ligated together, creating a composite DNA construct ( Engler et al., 2008 ). Golden Gate assembly offered advantages such as modularity, versatility, and compatibility with high-throughput applications. It allowed the construction of complex DNA constructs by assembling multiple fragments in a single reaction, enabling researchers to efficiently generate diverse genetic constructs with precise control. LCR was a technique that combined the principles of PCR and DNA ligation to facilitate the assembly of DNA fragments. In LCR, short DNA oligonucleotides, called splints, were designed to anneal to specific regions on the DNA fragments to be ligated. These splints hybridized to their complementary sequences, bringing the DNA fragments in close proximity for ligation. The ligation step was mediated by a DNA ligase enzyme, which catalyzed the formation of phosphodiester bonds between the adjacent DNA fragments. The ligated DNA fragments were then amplified by PCR using primers that hybridized to the ends of the assembled construct ( Li and Elledge, 2007 ). These advanced molecular cloning techniques had transformed the field by offering streamlined processes for the assembly of multiple DNA fragments. They eliminated the need for laborious traditional methods, such as restriction digestion and ligation, and provided advantages such as increased efficiency, reduced errors, and the ability to construct complex DNA constructs with precision.

Another significant advancement was the utilization of site-specific recombinases, such as the Cre-Lox system and the Flp-FRT system, to enable precise DNA rearrangements. These recombinases catalyzed the exchange, inversion, or deletion of DNA segments at specific target sites, facilitating the generation of complex DNA constructs and genetic modifications ( Sauer and Henderson, 1988 ; Buchholz et al., 1996). The Cre-Lox system, derived from the bacteriophage P1, consisted of two key components: the Cre recombinase enzyme and the LoxP sites. In the Cre-Lox system, the Cre recombinase recognized and bind to LoxP (locus of crossover [X] P1) sites present in the DNA. The LoxP site consisted of two 13-base-pair inverted repeats flanking a central 8-base-pair spacer region. The Cre recombinase binds to the LoxP site and catalyzes a strand cleavage and exchange reaction. This reaction involves the cleavage of the DNA strands at the specific LoxP sites, followed by the exchange or rearrangement of the cleaved DNA segments. The result was the excision, inversion, or rearrangement of DNA segments depending on the orientation and arrangement of the LoxP sites ( Sauer and Henderson, 1988 ). The Cre-Lox system had been widely used for conditional gene knockout, gene expression control, and the generation of tissue-specific or inducible genetic modifications. This system offered precise control over DNA modifications, as the recombination events occurred exclusively at the LoxP sites and did not affect surrounding genomic regions. Similarly, the Flp-FRT system, derived from the yeast Saccharomyces cerevisiae , operated through the interaction between the Flp recombinase enzyme and the FRT (Flippase Recognition Target) sites. FRT sites are DNA sequences that serve as recognition sites for the Flp recombinase. The FRT site consisted of two 13-base-pair inverted repeats separated by a central 8-base-pair spacer region. The Flp recombinase recognized the FRT site and catalyzed a similar strand cleavage and exchange reaction as the Cre-Lox system. By cleaving the DNA strands at the FRT sites and facilitating the exchange or rearrangement of the cleaved segments, the Flp-FRT system enabled the excision, inversion, or rearrangement of DNA segments, depending on the arrangement and orientation of the FRT sites (Buchholz et al., 1996).The Flp-FRT system offered high precision and versatility in genetic manipulations, which allowed researchers to precisely control the outcome of the recombination events. The benefits of these site-specific recombinase systems are manifold. First, they offered precise control over DNA modifications, allowing researchers to manipulate specific regions within a genome without affecting surrounding genetic elements. This targeted approach minimized unwanted off-target effects and preserved the integrity of the genome. Second, these systems provided flexibility and versatility in creating complex genetic modifications, such as gene knockouts, gene insertions, and conditional gene expression systems. The ability to precisely control DNA rearrangements at specific target sites opened up new avenues for functional studies and understanding gene regulation. Moreover, the reversible nature of these recombinase systems allowed for the creation of conditional genetic modifications, where DNA segments can be switched between different configurations, providing temporal and spatial control over gene expression.

In recent years, CRISPR-Cas9 technology has revolutionized molecular cloning and genetic engineering. Originally developed as a precise genome editing tool, CRISPR-Cas9 has been adapted for DNA cloning applications, allowing for efficient and targeted insertion of DNA fragments into specific genomic loci (Kleinstiver et al., 2014; Nishimasu et al., 2014 ). The CRISPR-Cas9 system has simplified and accelerated the process of creating transgenic organisms, generating knock-in or knock-out models, and facilitating the study of gene function ( Doudna and Charpentier, 2014 ). The working principle of the CRISPR-Cas9 system involves two main components: the Cas9 enzyme and a guide RNA (gRNA). The Cas9 enzyme acts as a molecular scissor and can be programmed to target specific DNA sequences with the help of the gRNA. The gRNA is designed to recognize a complementary DNA sequence adjacent to the target site, guiding the Cas9 enzyme to that specific location. Once the Cas9 enzyme binds to the target DNA sequence, it generates a double-strand break (DSB) at that site. This DSB triggers the cellular DNA repair machinery, which can be harnessed to introduce desired DNA fragments into the genome ( Doudna and Charpentier, 2014 ). CRISPR-Cas9 technology offers several significant benefits in molecular cloning and genetic engineering. First and foremost, it provides unparalleled precision in DNA targeting. The gRNA can be easily customized to recognize any desired DNA sequence, allowing researchers to selectively edit or insert DNA fragments at specific genomic loci. This level of precision enables the creation of highly specific genetic modifications, facilitating the study of gene function and the elucidation of molecular mechanisms. Another advantage of CRISPR-Cas9 technology is its efficiency and versatility. The Cas9 enzyme, guided by the gRNA, efficiently generates DSBs at the target site, triggering the cellular repair mechanisms. Researchers can exploit these mechanisms to introduce exogenous DNA fragments into the genome. This enables the precise insertion of DNA fragments, such as genes, regulatory elements, or reporter constructs, at desired genomic locations. In addition, the system can be used for gene knockouts by introducing DSBs that lead to gene disruptions or deletions. CRISPR-Cas9 technology has significantly streamlined the process of creating transgenic organisms and generating genetically modified cell lines, providing researchers with a powerful tool for studying gene functions and disease mechanisms. The advancements in synthetic biology have opened new avenues for molecular cloning. DNA synthesis technologies have improved significantly, enabling the de novo assembly of entire genes, pathways, or even genomes ( Kosuri and Church, 2014 ). This has paved the way for the creation of synthetic DNA constructs with tailored functions, such as metabolic pathways for bioproduction, novel enzymes, or even synthetic organisms with redesigned genomes ( Gibson et al., 2010 ; Hutchison et al., 2016 ). These recent developments in molecular cloning techniques have not only increased efficiency but also expanded the possibilities for genetic manipulation, DNA engineering, and biotechnological applications. By embracing these cutting-edge approaches, researchers can push the boundaries of what is achievable in the field of molecular cloning.

The Role of Molecular Cloning in Polymicrobial Infections

Polymicrobial infections, where multiple microorganisms are involved in an infection, are a common occurrence in clinical settings. Molecular cloning had become an important tool in identifying diseases caused by multiple microorganisms and understanding the exchanges that occur within microbial communities. These diseases, as described by Kim et al. (2005) , can be acute or chronic and are caused by combinations of viruses, bacteria, fungi, and parasites ( Sharma et al., 2014 ). When a single pathogenic microorganism creates a niche, other pathogenic microorganisms can inhabit it, leading to colonization or the emergence of disease by two or more non-pathogenic microorganisms ( Kim et al., 2005 ). Culture-based routine diagnostic testing is one treatment strategy, but it has limitations since it may not isolate all significant microbe species present in a sample. New bacterial community profiling techniques have revealed a greater diversity of microbes in infections caused by these bacteria than previously thought ( Sharma et al., 2014 ). It is the result of these findings that polymicrobial infections are increasingly being perceived as complex communities of interacting organisms, whose pathogenicity is determined by dynamic processes ( Rogers et al., 2010 ). One of the primary applications of molecular cloning in polymicrobial infections is the identification and characterization of individual pathogens. Traditional microbiological methods involve culturing the microorganisms from clinical samples, which can be time-consuming and may not always yield accurate results. In contrast, molecular cloning techniques such as PCR and NGS can identify and differentiate multiple pathogens in a single sample rapidly and accurately. These techniques can identify pathogens that are difficult or impossible to culture, making them a valuable tool in the diagnosis of polymicrobial infections. However, even these techniques have certain drawbacks ( Datta, 2023 ).

In the past few decades, advances in cloning and NGS technology had provided us with opportunities for gaining such insights about viable microbial cells ( Wheat, 2010 ). An approach based on sequence homology allowed identification of bacteria in polymicrobial infections by cloning and sequencing the 16S ribosomal gene ( Amann et al., 1995 ). With this method, it was possible to identify bacteria that died in the course of transportation or due to antibiotic treatment and to discover bacteria with specific growth requirements ( Kommedal et al., 2009 ). However, it may not be possible to detect rare members of a community with divergent target sequences with rRNA gene-based cloning and characterization ( Petrosino et al., 2009 ). Some of the limitations for it are due to primer bias and low sampling depth, which could be solved by 454 sequencing, pyrosequencing, or whole genome shotgun sequencing ( Petrosino et al., 2009 ). Hence, molecular cloning techniques, such as Loop-mediated isothermal amplification (LAMP), Multi-Plex PCR-Based Reverse Line Blot Hybridization (mPCR-RLB), Target Enriched-Multiplex PCR (Tem-PCR), Gene chip technology, and Multiplex Ligation-Dependent Probe Amplification (MLPA), are utilized, which play a crucial role in identifying the genes and pathways involved in polymicrobial infections ( Datta, 2023 ). These techniques enable researchers to gain a comprehensive understanding of the mechanisms underlying the interactions between pathogens, which is essential for developing novel treatments that target these interactions. LAMP is a powerful nucleic acid amplification technique that allows for the detection and quantification of specific DNA sequences at a constant temperature. LAMP amplifies DNA by exploiting the properties of Bst polymerase and the unique structure of the target DNA. It involves the design of specific primers for different regions of the target gene, resulting in the formation of a loop structure. The amplification reaction produces a characteristic ladder-like structure (amplicon), which can be visualized using fluorescence or turbidity changes ( Datta, 2023 ). LAMP is particularly useful for the prompt recognition of pathogenic microorganisms in laboratories with limited resources and experimental conditions. mPCR-RLB combines the sensitivity and the specificity of PCR with the high-throughput capabilities of reverse line blot hybridization. It involves the use of multiple primer sets that specifically bind to conserved regions of the microbial genome ( Datta, 2023 ). These primers are used in a multiplex PCR reaction to amplify multiple target DNA sequences simultaneously. The amplified DNA is then hybridized to a reverse line blot membrane containing probes for different microbes. The presence or absence of hybridization signals allows for the identification of specific microbial types in the sample. Tem-PCR combines the specificity of PCR with target enrichment to detect and identify microbes involved in polymicrobial infections ( Datta, 2023 ). The target enrichment step involves the selective amplification of specific regions of interest from the sample DNA using specific probes. This is followed by a multiplex PCR reaction, where multiple sets of primers amplify different targets in the same reaction. Tem-PCR allows for the simultaneous detection of multiple targets with high sensitivity and specificity. This technique is particularly useful for the detection of low-abundance targets in clinical samples. Gene chip technology, also known as DNA microarray, is a powerful tool for the detection and characterization of microbes. It allows for the simultaneous detection and analysis of multiple microbial genes, providing a comprehensive view of viral load and genetic diversity ( Datta, 2023 ). Gene chips consist of a glass slide or silicon wafer coated with thousands of DNA probes complementary to specific regions of the microbial genome to be tested. When a sample of genomic DNA or cDNA is labeled and hybridized to the chip, the binding of target sequences to the complementary probes generates fluorescent complexes. The resulting signals provide information on the relative expression levels of the genes in the sample, enabling the identification of differentially expressed genes and grouping of samples based on their gene expression profiles. MLPA is a technique used to detect and quantify specific sequences in a sample. It combines the specificity of PCR with ligation and probe amplification. MLPA involves the ligation of two probes, one specific for the target sequence and the other for a control sequence, followed by PCR amplification using universal primers ( Datta, 2023 ). The resulting pool of amplified fragments, proportional to the amount of DNA in the sample, can be quantified and analyzed to determine relative copy numbers of target sequences.

Molecular cloning also plays a crucial role in understanding the interactions between multiple pathogens in polymicrobial infections. For example, studies have shown that certain bacterial species can facilitate the growth of other pathogens, which can lead to more severe infections ( Doron and Gorbach, 2008 ). Molecular cloning techniques can help identify the genes and pathways involved in these interactions, allowing researchers to better understand the mechanisms behind polymicrobial infections. This understanding can lead to the development of new treatments that target the interactions between pathogens rather than just targeting individual pathogens. Once the key genes and pathways responsible for pathogen interactions have been elucidated through molecular cloning techniques, researchers can delve deeper into understanding the underlying mechanisms. Armed with this knowledge, they can develop innovative treatments specifically designed to disrupt the cooperative or synergistic relationships between the pathogens involved. Several novel treatment strategies have emerged from this approach, each targeting different aspects of pathogen interactions. One such strategy involves interfering with quorum sensing, a cell-to-cell communication system employed by many bacteria to coordinate their activities ( Rutherford and Bassler, 2012 ). Quorum sensing is a cell-to-cell communication system utilized by many bacteria, enabling them to synchronize their gene expression and coordinate their activities as a group ( Rutherford and Bassler, 2012 ). This coordinated behavior often leads to the production of virulence factors, biofilm formation, or the regulation of key processes required for the survival and colonization within the host. Researchers have been able to identify the genes and molecules involved in quorum sensing through molecular cloning techniques, and by understanding the specific components of the quorum sensing system, they can develop therapeutic agents that interfere with this communication process and disrupt the signaling and coordination among pathogens ( Vadakkan et al., 2018 ). There are different strategies employed to interfere with quorum sensing. One approach involves blocking or inhibiting the production or activity of quorum sensing signaling molecules, such as autoinducers ( Vadakkan, 2020 ). These signaling molecules are produced by bacteria and accumulate in the environment as the population density increases and they act as chemical signals, and when their concentration reaches a threshold, they trigger specific gene expression programs, coordinating behaviors among the bacterial community ( Vadakkan, 2020 ). Therapeutic agents targeting quorum sensing can be designed to mimic or block the action of these signaling molecules ( Milly and Tal‐Gan, 2023 ). For example, small molecules can be developed that either competitively bind to the receptor sites of bacteria, preventing the binding of natural signaling molecules, or mimic the signaling molecules themselves, leading to false or ineffective coordination among the pathogens ( Milly and Tal‐Gan, 2023 ). Another strategy involves inhibiting the production or activity of the enzymes responsible for synthesizing the signaling molecules, which results in the disruption in communication and coordination among pathogens ( Paluch et al., 2020 ). This interference can be achieved through the development of enzymatic inhibitors or by targeting the genes encoding the enzymes involved in the synthesis process ( Paluch et al., 2020 ). Such an approach holds promising potential in combating pathogenic infections by impeding their ability to communicate and coordinate their detrimental actions.

The Role of Molecular Cloning in Antimicrobial Peptides

In order to combat the rapidly increasing antimicrobial resistance, more effective antimicrobial peptides are being developed. These antimicrobial peptides will kill target cells promptly and be effective against antibiotic-resistant and clinically pertinent pathogens. Advances in molecular genetics of antibiotic biosynthesis offer new opportunities to improve antibiotic production. The DNA of antibiotic makers is enriched with genes coding for antibiotic biosynthesis enzymes. There have been two types of vectors developed to clone antibiotic genes: low-copy and high-copy plasmids ( Fakruddin et al., 2013 ). Many manufacturers are actively registering molecular clones of antibiotic synthesis genes. By manipulating the cloned genes encoding the enzymes involved in the biosynthetic pathway, new antibiotics can be produced as structural deviations of existing ones (Muller et al., 2007). These structural variants have diverse spectra and potency of activity against innumerable bacteria ( Sharma et al., 2014 ). Moreover, the production of highly purified antimicrobial peptides at competitive costs is one of the biggest challenges facing the development of antimicrobial peptides.

Chemical peptide synthesis can be used to yield either native or modified cationic peptides, but it is more expensive than isolating peptides from natural sources. In vivo synthesis in host cells using recombinant technology is a more effectual and cost-effective method of synthesis (Muller et al., 2007). Recombinant expression of antimicrobial peptides is a promising area of research despite the challenges posed by their toxicity to bacteria and susceptibility to degradation by proteases ( Sharma et al., 2014 ). This technology is widely recognized as an effective method for enhancing protein, peptide, and enzyme production. It offers advantages such as reduced time, well- established protocols, cost-effectiveness, and scalability ( Sinha and Shukla, 2019 ). Bacteria and yeast are the most commonly used host systems for expressing recombinant products, including antimicrobial peptides (AMPs) ( Gupta and Shukla, 2017 ). Escherichia coli , particularly the strain E. coli BL21 (DE3), is a popular choice due to its fast growth rate, high yields, established expression protocols, and commercial availability of expression vectors. Other bacterial systems like Bacillus subtilis have also been used although to a lesser extent. Among yeast, Pichia pastoris has been employed as a potential host. Scientists have developed strategies to address these issues, such as fusing peptide genes with larger proteins and then using enzymes or chemicals to cleave them and release active peptides ( Kuhnel et al., 2003 ). Promoter probe vectors are also being employed to clone DNA sequences with transcriptional control signals. One interesting approach involves neutralizing the positive charge of the peptide to enhance its activity ( Ali and Murrell, 2009 ). Another promising application of molecular cloning is the amplification of genes involved in biosynthesis pathways. By increasing the production of limiting enzymes, researchers have shown that it is possible to boost antibiotic production. This represents a significant step forward in the fight against antibiotic-resistant bacteria, which pose a major threat to public health. With the help of molecular cloning techniques, scientists are now better equipped to identify and develop novel antimicrobial agents that can be used to combat infectious diseases. Recombinant expression of AMPs is a popular method to produce large quantities of these peptides for further study or therapeutic use. However, the toxicity of some AMPs against bacteria and their susceptibility to proteolytic degradation can present challenges in the expression process ( Nizet, 2006 ). To overcome these issues, researchers have used molecular cloning techniques to fuse the AMP genes with larger proteins. The fusion proteins can then be cleaved by enzymes or chemicals to release active AMPs ( Ingham and Moore, 2007 ). To mitigate the toxicity of AMPs to the host strain, fusion proteins consisting of the target AMP and a carrier protein are often used. Carrier proteins possess anionic properties that neutralize the cationic charge of AMPs, reducing their toxicity and increasing solubility ( Li, 2009 ). Common fusion partners include thioredoxin, Small Ubiquitin-like Modifier (SUMO), Glutathione S-transferase (GST), Biotin Carboxyl Carrier Protein (BCCP), and Green Fluorescent Protein (GFP). Cleaving the carrier protein from the target AMP typically requires the use of chemicals or enzymes. Chemical cleavage, although less specific, is considered more efficient than enzymatic cleavage, but it can introduce modifications in the side chain ( Sinha and Shukla, 2019 ). Cleavage processes often leave one or two non-native residues at the N-terminus of the cleaved AMP ( Sinha and Shukla, 2019 ). Affinity tags are sometimes conjugated to fusion partners to facilitate easy purification using affinity chromatography methods ( Meiyalaghan et al., 2014 ). Examples include His6-thioredoxin tagged GSL1 fusion protein, which can be purified by affinity chromatography, and fusion partners like glutathione S-transferase, which have inherent affinity properties and eliminate the need for additional tags ( Schäfer et al., 2015 ). Self-cleaving tags with inducible proteolytic activity have also been used to simplify the separation of fusion tags. Thioredoxin and SUMO are the preferred fusion partners for the expression of recombinant AMPs ( Xia et al., 2013 ). For instance, cecropin XJ, an insect AMP, was highly expressed in E. coli as a fusion peptide along with thioredoxin ( Sinha and Shukla, 2019 ). Similarly, LsGRP1C protein production was assisted by a yeast SUMO tag in the E. coli host system, resulting in a high yield of the soluble SUMO-LsGRP1C fusion protein ( Lin et al., 2017 ). BCCP has been used as a fusion protein for AMP expression, and B. subtilis has been employed for the recombinant expression of cathelicidin-BF from snake venom ( Luan et al., 2014 , Orrapin and Intorasoot, 2014 ). Another method for cloning AMPs is through the use of promoter probe vectors. These vectors contain transcriptional control signals that can be used to clone DNA sequences with transcriptional control signals by neutralizing the positive charge of the peptide ( Muriana and Klaenhammer, 1991 ). This allows for more efficient expression of AMPs in various hosts. Furthermore, molecular cloning has also been used to amplify genes coding for limiting enzymes in biosynthesis pathways, which has been shown to increase the production of antimicrobial compounds ( Adrio and Demain, 2010 ). This approach has led to the discovery and production of new antimicrobial compounds with potent activity against a number of microorganisms. Yeasts, particularly the methylotrophic yeast Pichia pastoris , have become increasingly important in genetic engineering and recombinant protein production due to their ease of genetic manipulation, ability to perform complex post-translational modifications, and rapid growth in cost-effective media ( Kim et al., 2015 ). P. pastoris is a popular host for heterologous expression of recombinant AMPs ( Ahmad et al., 2014 ). It offers advantages over E. coli , including the presence of a methanol-induced alcohol oxidase promoter, absence of endotoxins, correct protein folding capability, and suitability for large-scale production ( Ahmad et al., 2014 ). In one study, the NZ17074 gene was synthesized and fused with SUMO3 in P. pastoris X-33, and the carrier protein was subsequently cleaved using formic acid ( Wang et al., 2014 ). AMPs from plants, fruits, and chicken have also been expressed in P. pastoris but without the use of fusion proteins ( Meng et al., 2017 ). The emergence of advanced gene editing tools, including Zinc-Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeat-CRISPR-associated protein (CRISPR-Cas), has opened up new possibilities in the field of gene editing ( Dangi et al., 2018 ). These tools make it easier to manipulate the genomes of expression hosts, enabling targeted modifications of specific genes to achieve desired outcomes. By harnessing gene editing technology, it is now possible to revolutionize the production of AMPs, particularly in response to the increasing demand for industrially and therapeutically valuable AMPs ( Sinha and Shukla, 2019 ).

The Role of Molecular Cloning in Recombinant Cytokines

There are several fundamental homeostatic mechanisms that are modulated by cytokines ( Huang et al., 2005 ). They include fever, acute phase infections, wound healing, inflammation, immune responses at the cellular levels, and tumor deterioration. Recombinant DNA technology has allowed us to clone the genes that encode these proteins, making it possible to use unrestrained measures of these cytokines to treat disease. There can be changes in amino acid sequence, absence of glycosylation ( E. coli ), and changes in glycosylation pattern (yeast, mammals, and insects) ( Bandaranayake et al., 2011 ). Proteins expressed in the mature form in different host cells can also differ in their specific activities for several reasons ( Sharma et al., 2014 ). Studies show that cytokines expressed in different host cells can have different pharmacokinetics, biologic properties, and immunogenicity due to physiochemical differences ( Descotes, 2009 ). To explore the structure-function relationship of cytokines, expression vectors can be used to hypothesize recombinant forms. In addition to cytokines engineered for improved clinical efficiency or novel specificities, heterologous expression systems have also been used to create streamlined cytokines ( Bermúdez-Humarán et al., 2011 ). Nowadays, recombinant cytokines are available as therapeutic agents. There has been evidence that GM-GSF or G-CSF can reduce the period and risk of infectious complications associated with chemotherapy-induced neutropenia ( Sharma et al., 2014 ). Molecular cloning techniques are utilized to produce recombinant cytokines and enable their mass production. One of the most commonly produced cytokines through molecular cloning is interferon- alpha (IFN-α), which has potent antiviral and antiproliferative properties (Kumar et al., 2018). Recombinant IFN-α has been used in the treatment of hepatitis B and C infections, as well as certain types of cancer, including melanoma and leukemia. Molecular cloning is also used to produce other cytokines, such as interleukins and tumor necrosis factor (TNF). Recombinant TNF has been used in the treatment of bladder cancer, while interleukins have been used to enhance the immune response against cancer cells (Muller et al., 2003). In addition, the use of cytokines produced through molecular cloning has been explored in the treatment of autoimmune diseases, such as rheumatoid arthritis and multiple sclerosis. Moreover, the investigation into the therapeutic potential of molecularly cloned cytokines has extended to encompass the management of autoimmune disorders, including rheumatoid arthritis and multiple sclerosis. These chronic conditions, such as rheumatoid arthritis, are characterized by persistent joint damage caused by an overactive immune response targeting the synovial membrane, cartilage, and bone ( Zhang, 2021 ). While the precise cause of rheumatoid arthritis remains elusive, significant progress has been made in understanding its complex pathogenesis involving various cell types and signaling pathways ( Verhoef et al., 2019 ).

Autoimmune processes and cytokines have emerged as key players in the initiation and perpetuation of rheumatoid arthritis. Notably, the extensively studied tumor necrosis factor (TNF), interleukin-6 (IL-6), and interleukin-1 (IL-1) have been implicated in the disease progression ( Upchurch and Kay, 2012 ). Consequently, therapeutic strategies have focused on combating inflammation, with non-steroidal anti-inflammatory drugs and glucocorticoids serving as primary symptomatic management options ( Zhang, 2021 ). In recent years, disease-modifying antirheumatic drugs (DMARDs) have revolutionized rheumatoid arthritis treatment by specifically targeting pro-inflammatory cytokines and their respective receptors ( O’Dell, 2004 ). These biological DMARDs include TNF-α inhibitors (such as infliximab, etanercept, adalimumab, golimumab, and certolizumab pegol), IL-1 inhibitors, monoclonal antibodies against the IL-6 receptor (tocilizumab), T cell signaling inhibitors (abatacept), and monoclonal antibodies targeting CD20 (rituximab) ( Klarenbeek et al., 2011 ). When combined with conventional synthetic DMARDs, early initiation of these treatments has shown promising results, leading to improved clinical outcomes and reduced joint damage ( Zhang, 2021 ).

Along with using recombinant immunomodulators to enhance the host’s defense mechanism, attempts have been made to repackage defective immune genes ( Miyake et al., 2001 ). The gene therapy of adenosine deaminase deficiency (ADA) deficiency has successfully treated patients with a defective T cell immune response ( Aiuti and Roncarolo, 2009 ). It is possible to transfer a cloned ADA gene into lymphocytes of a patient using a retroviral vector; enzymatic and immune functions are thus reinstated ( Aiuti and Roncarolo, 2009 ). Many ongoing research projects focus on gene therapy’s use in HIV infection and oncogenic virus-associated cancer, even though there is no clear evidence of its impact on infectious disease. Through HBV- or HHV-8-encoded surface receptors, the thymidine kinase gene has been inserted into the cancer cell in HBV-associated hepatocellular carcinomas and HHV68-associated Kaposi’s sarcomas ( Sharma et al., 2014 ). The patient is given ganciclovir when the thymidine kinase gene is integrated into their cancer cells, where the drug amasses and becomes toxic ( Kieback et al., 2008 ). Future developments in the creation of cytokine receptor agonists show promise in a number of ways. One avenue of exploration involves using de novo protein design in addition to the existing combinatorial ligand engineering strategies. De novo design involves a computational approach to design analogues of interleukin-2 (IL-2) and interleukin-15 (IL-15) that can signal independently of IL-2Rα/IL-15Rα ( Saxton et al., 2023 ). These synthetic analogues, known as “neoleukins,” have unique structural features compared to natural cytokines. They exhibit enhanced stability and improved effectiveness in mouse tumor models. Currently, neoleukins are undergoing phase I clinical trials for multiple cancer types, either alone or in combination with immune checkpoint blockade therapies. Advancements in artificial intelligence and machine learning-based protein structure prediction are expected to facilitate the development of even more sophisticated de novo designed cytokines ( Saxton et al., 2023 ). These cytokines may have entirely distinct structural topologies that can modulate receptor geometry and composition in novel ways, potentially revolutionizing our understanding of cytokine signaling. While de novo cytokines offer advantages such as enhanced stability and potential ease of manufacturing, they may carry a risk of increased immunogenicity due to their non-human amino acid sequences ( Saxton et al., 2023 ). However, ongoing efforts focus on minimizing immunogenicity by reducing the number of mutations and utilizing databases to assess potential neo-epitopes. Improvements in the pharmacokinetic and pharmacodynamic properties of cytokines are also crucial for their clinical success and strategies like half-life extension through Fc fusions or PEGylation, as well as local cytokine production via engineered T cells, aim to enhance cytokine safety and efficacy ( Saxton et al., 2023 ). However, these modifications can influence cytokine signaling activity, necessitating a thorough understanding of their effects. Experimental models for screening and testing novel cytokine activities include in vitro cell culture and mouse models, but these have limitations in predicting human efficacy. Human patient-derived organoid cultures with diverse immune cell populations may provide valuable insights into the effectiveness of cytokine-based drugs in humans ( Saxton et al., 2023 ). Immunogenicity is another significant consideration for protein therapeutics, including engineered cytokines. Non-human proteins, like de novo designed cytokines, may carry a higher risk of eliciting immune responses due to their divergent amino acid sequences, so efforts to minimize immunogenicity, especially for mutant cytokines, include reducing the number of mutations and monitoring for cross-neutralizing antibodies ( Saxton et al., 2023 ).

The Role of Molecular Cloning in Gene Therapy

An individual can be treated with gene therapy by introducing a regular gene into their genome in order to mend the mutation that is the origin of genetic disease. In addition to possibly repairing the mutation, insertion of a regular gene into another functional gene may result in a new mutation if the normal gene integrates into another functional gene’s chromosomal site ( Khan et al., 2016 ). Transformed cells may proliferate if normal genes replace mutant genes, leading to the restoration of the non-disease phenotype of the entire body. To date, human gene therapy has only been tested on somatic cells to treat cancer and severe immunodeficiency syndromes ( Khan et al., 2016 ). It is possible for gene therapy to inverse the signs of disease in somatic cells, but the adjustment does not pass on to future generations. By placing corrected cells inside the germ line (e.g., cells of the ovary or testis), gene therapy will ensure that the next generation of cells undergoes meiosis and contributes to standard gametic development ( Khan et al., 2016 ). In the field of health services, gene therapy is a progressive technique that has therapeutic prospect. The first successful report in gene therapy for the cure of genetic diseases gave physicians a promising approach to treating the lethal genetic disorders ( Cavazzana-Calvo et al., 2000 ). Treatment for the primary immunodeficiency adenosine deaminase-deficiency (ADA-SCID) shows good results with this approach. An improved gene transfer protocol and myeloablative conditioning regime, however, were later used to achieve fruitful results by aiming the hematopoietic stem cells (HSCs) ( Aiuti et al., 2002 ).

The expression of specific genes by lentiviral vectors can correct X-linked disorders and adrenoleukodystrophy (X-ALD) ( Cartier et al., 2009 ). Based on HIV-1 genes, X-ALD protein expression indicates gene-correction of true HSCs. As part of the treatment of metastatic melanoma through immunotherapy, lentiviral vectors were used for the first time in the cure of genetic human diseases ( Montini et al., 2012 ). By expanding the field of health sciences through immunotherapy, new opportunities were unlocked for treating serious diseases that cause death ( Morgan et al., 2006 ). In two patients, continuous levels of T cells engineered to recognize tumors in the blood after infusion resulted in the recession of metastatic melanoma lesions up to a year after infusion. The bioengineered T cells were later studied for the treatment of chronic lymphocytic leukemia and metastatic synovial cell carcinoma where autologous T cells were innately altered to express Chimeric Antigen Receptors (CARs) that specifically bind to B cell antigen CD19 ( Robbins et al., 2011 ). It has shown remarkable results for incorrigible autosomal recessive dystrophies, such as congenital blindness and Leber congenital amaurosis (LCA), in which gene transfer to a small number of cells at anatomically discrete sites has the potential to confer therapeutic benefit. Through gene therapy, a variety of cancers have been treated, including lung, gastrointestinal, hematological, gynecological, skin, urological, and neurological tumors ( Khan et al., 2016 ). In order to treat diverse types of cancer, tumor suppressor genes have been inserted into immunotherapy, oncoturlytic virotherapy, and gene-directed enzyme prodrugs. In some cancer treatment strategies, p53 gene transfer is combined with chemotherapy or radiotherapy to increase the effectiveness of the tumor suppressor gene. An effective new anticancer agent (Ad5/35-EGFP) is being developed from fiber chimeric adenovirus vectors for the improved cure of hepatocellular carcinoma ( Khan et al., 2016 ). In hepatocellular carcinoma (HCC), these vectors were established to improve transduction and produce more virus progeny as a consequence of proper assaying. As a result of complex transgenic expression, in vitro HCC cells were found to possess enhanced antitumor activity while normal cells remained cytotoxic-free. As a result of the use of this technology, tumor growth was also repressed ( Zhang et al., 2011 ). Recently, cancer gene therapy has gained more advanced technology and expanded its effectiveness ( Lam et al., 2013 ). A mutation of the ABCA1 gene in high-density lipoproteins can cause the cells to discriminate into macrophages. Knockouts of this gene in embryonic stem cells augment the capability of these cells to differentiate into macrophages and precisely aim abnormal cells. A study of these allele replacements will provide insights into the regulatory variants that alter macrophage transcription and stability of mRNA ( Smith, 2016 ).

Yan et al. (2020) have spearheaded a groundbreaking advancement in molecular biology by introducing a cutting-edge technique called “Nimble Cloning.” This pioneering method revolutionizes standardized molecular cloning, a fundamental technology in the field. By harnessing the combined power of the restriction enzyme SfiI and the T5 exonuclease, Nimble Cloning enables simultaneous vector linearization and generation of 3’-overhangs. Notably, this novel cloning system accommodates both PCR products and plasmids as inputs for the cloning reaction, rendering it highly efficient and adaptable to gene expression in both prokaryotic and eukaryotic systems. What sets Nimble Cloning apart is its remarkable versatility and simplicity. It empowers researchers to reuse DNA fragments or plasmid entry clones, facilitating efficient and streamlined workflows. This innovative method proves to be equally adept at cloning single or multiple fragments, as well as facilitating multi-site cloning. Consequently, the possibilities for modular assembly of DNA constructs are greatly expanded. In their groundbreaking research, Yan et al. (2020) introduced Nimble Cloning, a novel technique inspired by Gibson assembly, for the seamless assembly of DNA fragments. By leveraging the power of enzyme-catalyzed reactions, they demonstrated that Nimble Cloning surpasses traditional Gibson assembly in terms of cloning efficiency. Excitingly, this method eliminates the need for Taq DNA ligase, streamlining the process without compromising effectiveness. Through meticulous experimentation with single and multiple DNA fragments, Yan et al. successfully validated the superior performance of Nimble Cloning, consistently achieving over 99% positive clones. To further emphasize its versatility, they employed unique adapters in standardized cloning reactions, confirming that gene expression remained unaffected in both prokaryotic and plant systems. These findings not only establish Nimble Cloning as an efficient and reliable method for genetic engineering but also pave the way for future advancements in DNA fragment assembly.

In recent years, the CRISPR-Cas9 system has revolutionized the field of gene therapy. This revolutionary gene editing tool enables precise and efficient modification of specific DNA sequences within the genome. Molecular cloning plays a critical role in the construction of CRISPR-Cas9 vectors, which are used to deliver the Cas9 nuclease and guide RNA sequences into target cells. The ability to edit genes with unprecedented precision has opened up new possibilities for treating genetic diseases by correcting or modifying disease-causing mutations. Advancements in DNA synthesis technologies have facilitated the design and construction of synthetic genes and gene circuits for gene therapy applications. Molecular cloning techniques allow the assembly of these synthetic genes into expression vectors, enabling the production of therapeutic proteins or the regulation of gene expression in a controlled manner. In terms of gene delivery, tissue-specific targeting has become an area of active research, and by engineering viral and non-viral vectors to possess tissue-specific promoters or ligands, researchers aim to enhance the specificity and efficiency of gene delivery to target tissues or cells. This targeted approach minimizes off-target effects and improves the overall safety and efficacy of gene therapy. Innovations in genome engineering techniques, such as base editing and prime editing, have expanded the possibilities of gene therapy. These advanced tools enable precise modification of individual bases within the genome, offering the potential to correct disease-causing mutations without the need for introducing foreign DNA.

The Role of Molecular Cloning in Epidemiology

The rise of multidrug-resilient pathogens necessitates early molecular epidemiology outlining, both for comprehensive public-health reconnaissance and for timely treatment of infested patients. Due to the inoculum size and culturing conditions’ inconsistency, conservative tests of this type require extended culturing times (48–72 h) ( Yang and Rothman, 2004 ). As genetic mechanisms of drug resistance are explained, nucleic-acid-based assays are being developed to report these inadequacies ( Yang and Rothman, 2004 ). Three examples of how molecular epidemiology can be applied clinically are provided next. Although the absence of a resistance gene does establish a lack of resistance through that particular genetic mechanism, the presence of a resistance gene does not ineludibly infer its expression and conferment of phenotypic resistance: for example, the mecA gene is responsible for methicillin resistance ( Yang and Rothman, 2004 ). With its high sensitivity and specificity, the mecA-PCR has become the most consistent method for detecting methicillin-resistant staphylococcus aureus (MRSA) due to its high detection sensitivity (Tenover et al., 1999) The detection of rifampicin resistance in M tuberculosis can also be performed using PCR-based resistance testing. RNA polymerase resistance in M tuberculosis is well characterized and is conferred by mutations within the DNA-directed RNA polymerase subunit beta ( rpoB ) gene, which result in amino acid substitutions in the rpoB subunit (Telenti et at.,1997). The Line Probe assay (LiPA; Inno-Genetics) targets the mutation-prone rpoB gene segment ( De Beenhouwer et al., 1995 ). A dramatic waning in detection time is provided to clinicians with this method, which is acute for treatment decisions, with an association of over 90% with typical resistance-detection methods ( Marttila et al., 1999 ). There are many mutations and genetic loci involved in other M tuberculosis drug resistances, which makes genotyping more challenging ( Yang and Rothman, 2004 ). Technical innovations like multiplex PCR or DNA microarray allow concurrent extension and analysis of multiple target sequences and will likely be able to overcome this future challenge ( Elnifro et al., 2000 ) A PCR followed by nucleotide sequencing method is currently the most habitually used process to identify drug-resistant mutations in HIV-infected patients with ever-increasing indication supporting their prediction value ( Demeter and Haubrich, 2001 ). Even though genotypic tests are more multifaceted than typical antimicrobial susceptibility tests, they provide perception into resistance development by perceiving mutations at concentrations too low to affect phenotypic assay susceptibility ( Yang and Rothman, 2004 ). Also, they have the benefit of detecting mutations that do not cause drug resistance but do designate selective drug pressure, which could influence treatment decisions for individual patients.

A recent study by Zhang et al. (2021) showed that the pathogen Carbapenem-Resistant Acinetobacter baumannii (CRAB) poses a significant threat to both health and economy. It haunts hospitals, using them as breeding grounds for its insidious transmission. The vulnerable victims of CRAB’s infections are often those who find themselves confined within hospital walls, with open wounds and lengthy stays. The elderly, whose natural defenses have weakened, are especially susceptible. Strikingly, most patients in the study suffered from multiple diseases simultaneously, with pulmonary infections, such as pneumonia and respiratory failure, being the most common. This aligns with the well-known association between CRAB and pneumonia and bloodstream infections. The key to CRAB’s resistance lies in its ability to produce carbapenemases, such as blaOXA-23 and blaOXA66, which render it impervious to carbapenem antibiotics ( Zhang et al., 2021 ). This study by Zhang et al. (2021) confirmed that all strains harbored these carbapenem resistance genes. The mechanisms behind CRAB’s increased antibiotic resistance are multifaceted, involving mobile genetic elements, chromosomal β-lactamases like blaADC, and the presence of efflux pumps ( Zhang et al., 2021 ). The study also highlighted the presence of various genetic structures, including ISs and AbaR-type genomic resistance islands, which facilitate the spread of antibiotic resistance determinants among pathogens, compromising treatment efficacy. But CRAB is not merely resistant; it is also armed with virulence factors that allow it to thrive in the inhospitable environment of a hospital. The study also revealed that all CRAB strains possessed a repertoire of virulence genes, such as bap, csuABCD, pgaABCD, bfmRS, entE, ompA , and plcD . These genes contribute to the formation and maintenance of biofilms, which are strongly associated with MDR strains and clinical infections. In addition, CRAB isolates carried genes involved in the production and uptake of the acinetobactin siderophore, further enhancing their infectious prowess ( Zhang et al., 2021 ). When it comes to the origins of CRAB outbreaks, molecular cloning holds the key to unraveling the mystery. Previous reports have linked clonal outbreaks to international clone lineages, with European clones I, II, and III being the most prominent. In the study conducted by Zhang et al. (2021) , all CRAB isolates belonged to ST2, which is associated with outbreaks and harbors the blaOXA-23 gene. Interestingly, certain clone types, such as ST2, ST25, and ST78, exhibited enhanced biofilm formation, likely contributing to their success in colonizing the clinical environment ( Zhang et al., 2021 ). Through the lens of core genome phylogenetic analysis, it became clear that specific CRAB strains dominated the scene. Cluster 1 and cluster 2 emerged as the primary players, with clone 1 and clone 2 persisting throughout the study period ( Zhang et al., 2021 ). However, changes in clone groups were observed over the years, indicating the dynamic nature of CRAB populations. These clones, particularly clone 1, underwent population expansion and exhibited distinct resistance, virulence, and insertion sequence patterns compared to clone 2 ( Zhang et al., 2021 ). These findings emphasize the genetic diversity within CRAB outbreaks and the importance of monitoring and controlling the spread of specific clones.

The Role of Molecular Cloning in Bioterrorism

Biological warfare, in its essence, is the strategic utilization of living organisms or their by-products to cause harm or destruction. It involves the deliberate release or deployment of pathogens, toxins, or biological agents with the intent to incapacitate, debilitate, or even kill living organisms, including humans. This form of warfare relies on exploiting the natural abilities of organisms to produce toxins or propagate diseases, harnessing their destructive potential as a means of achieving military or political objectives. Biological warfare encompasses a range of tactics, from the use of infectious diseases as weapons to the manipulation of toxic substances derived from a number of organisms. Its history intertwines with humanity’s understanding of the natural world and our capacity to wield its forces for destructive purposes. An anthrax outbreak that occurred has drawn much attention to the growing threat of bioterrorism. Responsibility of the clinicians will be crucial in instigating applicable response actions if they can recognize and diagnose real or suspected bioterrorism events quickly and accurately ( Pavlin et al., 2002 ). It is difficult to discriminate between a bioterrorism victim’s symptoms and symptoms of an ordinarily encountered disease process, as was the case at the time of the 2001 anthrax episode ( Pavlin et al., 2002 ). In suspected clinical outbreaks, conventional culture-based assays cannot detect bioterrorism agents due to the formerly designated restrictions. As a result of the prolonged incubation required by conventional microbiological methods, the laboratory is exposed to increased biohazard risks because of the redundant proliferation of bioterrorism pathogens ( Yang and Rothman, 2004 ). Many bioterrorism agents have, of late, been studied using PCR-based assays, including Variola major, Bacillus anthracis, Yersinia pestis , and Francisella tularensis ( Espy et al., 2002 ). Bioterrorism agent PCR diagnostics was used for both screening preclinical victims for early prophylactic treatment and diagnosing symptomatic individuals ( Yang and Rothman, 2004 ). Despite the similarity between most bioterrorism-induced illnesses and natural outbreaks, it was possible that the contributory agents of bioterrorism may have been genetically engineered to be more virulent, impervious to antibiotics or vaccines, or to produce phenotypic characteristics resembling multiple infections via the insertion of recombinant genes ( Alibek, 1999 ). Since DNA-based methodologies are more easily adjustable and proficient in uncovering more comprehensive information embedded within genetic sequences, they are likely to be more valuable than conventional detection methods in such cases ( Yang and Rothman, 2004 ).

The potential role of genetic engineering in enhancing the lethality of infectious agents is a topic often discussed in relation to biological warfare. While there is some validity to this claim, it is crucial to consider the following perspective: Imagine a scenario where a bioterrorist endeavors to genetically modify a harmless laboratory bacterium, such as E. coli . The aim would be to render the bacterium “invisible” to the human immune system upon entry into the body ( Clark and Pazdernik, 2016 ). In addition, the bacteria could be engineered to release toxins, thwarting immune cells and introducing genes to impede the vital iron supply from blood cells. Lastly, the bacterium could be modified to possess high levels of infectivity. Such a biological agent would undoubtedly pose a formidable threat. Astonishingly, this bacterium already exists in nature—it is known as Yersinia pestis , the notorious causative agent of bubonic plague ( Clark and Pazdernik, 2016 ). Endemic in different parts of the world, including China, India, Madagascar, and the United States, this pathogen demonstrates how nature has provided an exceptionally dangerous biological weapon. Consequently, the notion of enhancing infectious diseases through genetic engineering appears to be a minor concern. Although information regarding the Soviet germ warfare facility’s modification of the smallpox virus and the creation of artificial mutants and hybrids remains largely undisclosed, recent experiments involving mousepox (Ectromelia virus) have yielded alarming outcomes. Mousepox, a virus primarily infecting mice, exhibits varying degrees of virulence depending on the mouse strain. Genetically resistant mice rely on cell-mediated immunity rather than antibodies to combat the virus, with natural killer (NK) cells and cytotoxic T cells effectively eliminating infected cells and clearing the virus from the body. In an attempt to improve and balance the immune response, researchers introduced the human gene for the cytokine interleukin-4 (IL-4) into the mousepox virus. IL-4 stimulates B cell division and antibody synthesis, which theoretically should have led to an enhanced immune response ( Clark and Pazdernik, 2016 ). Surprisingly, the results were contrary to expectations, as the engineered virus displayed significantly heightened virulence. It not only caused mortality in all genetically resistant mice but also claimed the lives of 50% of vaccinated mice. Excessive IL-4 expression suppressed NK cells and cytotoxic T cells while failing to enhance the antibody response ( Clark and Pazdernik, 2016 ). Similar results have been observed with different strains of Vaccinia virus, which is utilized for smallpox vaccination. The repercussions of inserting IL-4 or other immune regulators into smallpox itself and the potential to undermine the immune response and increase virulence remain uncertain ( Clark and Pazdernik, 2016 ). Poxviruses possess genes called cytokine response modifier ( crm ) genes, designed to hinder the action of NK cells and cytotoxic T cells, further complicating the assessment of smallpox’s virulence ( Clark and Pazdernik, 2016 ). Nevertheless, genetic engineering allows for the concealment of a potentially hazardous virus within a harmless bacterium, a phenomenon already witnessed in nature when bacteriophages incorporate their genomes into bacterial chromosomes or plasmids, subsequently re-emerging to infect other hosts. Theoretically, cloning the complete genome of a small animal or plant virus into a bacterial plasmid could serve as the basis for a biological weapon, while larger viruses could be accommodated using bacterial or yeast artificial chromosomes. In the case of RNA viruses, the viral genome must first be reverse transcribed into complementary DNA (cDNA) before cloning into a bacterial vector. Viruses harboring toxic sequences, which are not stably maintained on bacterial plasmids, could potentially be cloned as separate fragments. This approach has proven successful with yellow fever virus albeit requiring in vitro ligation of the fragments to generate a complete, functional cDNA ( Clark and Pazdernik, 2016 ). Different cell types, including bacteria and eukaryotes, are capable of taking up DNA or RNA under specific conditions through transformation. Consequently, the naked nucleic acid genomes of numerous DNA and RNA viruses retain their infectivity even without their protein capsids or envelopes. Once a viral genome is cloned, the DNA molecule containing it becomes infectious itself. Alternatively, cDNA versions of RNA viruses can infect host cells, giving rise to new virus particles containing RNA. Poliovirus, influenza, and coronavirus are among the RNA viruses that have demonstrated this capability. An ingenious strategy for generating an RNA virus involves cloning the cDNA version of its genome downstream of a strong promoter in a bacterial plasmid. Transcription within the induced bacterial cell results in the production of a large number of infectious viral particles. Combining a dangerous human RNA virus with a harmless intestinal bacterium, controlled by a promoter responsive to intestinal conditions, could pose a significant threat ( Clark and Pazdernik, 2016 ). While certain pathogenic bacteria exhibit slow growth or are challenging to the culture outside their host organisms, advancements in biotechnology have facilitated the identification of infectious microbes through molecular diagnostics. Instead of relying on classical microbiological techniques to grow and identify disease-causing agents, molecular diagnostics employ the analysis of molecules, primarily DNA but also RNA, proteins, and volatile organic compounds. These molecular techniques offer advantages in terms of speed, accuracy, and sensitivity.

Fluorescent in situ hybridization (FISH) is one such diagnostic method, involving the direct probing of biopsies or patient samples with fluorescent DNA oligonucleotides specific to a target pathogen ( Clark and Pazdernik, 2016 ). If the pathogen is present, the probe binds to complementary DNA in its chromosome, enabling visualization of fluorescence under a microscope. An innovative approach utilizing peptide nucleic acid (PNA), which replaces the negatively charged sugar-phosphate backbone of DNA with a neutral peptide backbone, enhances the binding affinity of PNA probes to complementary DNA and facilitates their entry into bacterial cells. PCR amplification of target DNA sequences, unique to a particular pathogen, is another commonly used method in molecular diagnostics ( Clark and Pazdernik, 2016 ). The ability to design primers specific to a pathogen allows for the amplification of target DNA, enabling PCR to serve as a diagnostic tool. PCR’s versatility stems from its potential to detect a single molecule of target DNA and its applicability to microbes that are challenging to culture in the laboratory. However, PCR is susceptible to contamination and false positives. Randomly amplified polymorphic DNA (RAPD), a PCR variant, distinguishes different strains of the same bacterial species, aiding in epidemiological studies to track the spread of infectious diseases. Each microorganism species possesses a unique small-subunit ribosomal RNA (SSU rRNA) sequence, such as 16S rRNA in bacteria and 18S rRNA in eukaryotes ( Clark and Pazdernik, 2016 ). Therefore, clinicians can employ PCR to amplify the gene encoding the microbe’s SSU rRNA when faced with an unknown infection. By sequencing and comparing the PCR fragment to a database of known DNA sequences, the pathogen can be identified. Checkerboard hybridization is a technique that utilizes SSU rRNA as a basis, allowing for the simultaneous detection and identification of multiple bacteria in a single sample. Horizontal lines on a hybridization membrane contain probes specific to different bacteria, while PCR amplification of the SSU rRNA gene from clinical samples, potentially harboring a mixture of pathogens, generates fluorescently labeled fragments that are applied vertically to the membrane. Fluorescent spots indicate samples that have hybridized with the probes. Abbott Laboratories has developed a potentially revolutionary technology called PLEX-ID, which combines traditional PCR with mass spectrometry to identify unknown microbes in patient samples ( Clark and Pazdernik, 2016 ). By analyzing the mass of amplified DNA fragments using a mass spectrometer, the DNA sequence can be deduced, enabling the identification of the pathogen. PLEX-ID has the capability to provide a diagnosis within eight hours. In the future, disease diagnosis may be achievable using an “electronic nose,” a device that detects volatile organic compounds released by pathogens or by the body in certain diseased conditions ( Clark and Pazdernik, 2016 ).

The Role of Molecular Cloning in SARS-COV 2 Treatment

B cell receptor (BCR) repertoires display remarkable sequence diversity resulting from somatic recombination and hypermutation processes occurring during B cell development. BCRs are transmembrane receptors situated on the surface of B cells, and their variable regions interact with specific antigen epitopes to initiate an immune response through antibody production ( Zhou et al., 2021 ). This variable region shares identical gene sequences with the corresponding antibody produced by the B cell. The variability in the variable region is generated by somatic recombination of three gene segments in the heavy (H) chain locus (V, D, J) and two gene segments in the light (L) chain locus (V, J) ( Zhou et al., 2021 ). The variable region of an antibody, also known as immunoglobulin (Ig), determines its specificity for binding to a specific viral antigenic epitope. Somatic hypermutation occurs during B cell proliferation in the germinal center, introducing random mutations in the genes encoding the variable region of individual monoclonal antibodies (mAbs) ( Zhou et al., 2021 ). This process is crucial for the development of high-affinity antibodies, a phenomenon referred to as antibody affinity maturation. Importantly, no two B cells possess identical BCRs. Therefore, the cloning of a functional mAb requires the analysis and cloning of one B cell at a time to ensure the native pairing of antibody heavy and light chains ( Bassing et al., 2002 ).

Advancements in antibody gene cloning techniques, such as hybridoma technology, human B cell immortalization, antibody phage display, human immunoglobulin transgenic mice, and single B cell antibody technology, have revolutionized the process of cloning functional human neutralizing antibodies (HuNAbs) ( Tiller, 2011 ). Among these techniques, single B cell-based antibody cloning has been extensively employed to isolate SARS-CoV-2-specific antibodies during the ongoing COVID-19 pandemic. This method involves amplifying auto-paired Ig heavy- and light-chain RNA sequences from a heterogeneous population of memory B cells, followed by their in vitro construction into functional mAbs ( Niu et al., 2019 ). The successful application of this technique has been demonstrated in the isolation of neutralizing mAbs against various viral infections, such as human immunodeficiency virus (HIV) and SARS-CoV-2 ( Corti et al., 2015 ). In recent years, the combination of single-cell reverse transcription polymerase chain reaction (RT-PCR) and single B cell sorting has significantly improved the success rate of antibody gene cloning. The acquisition of single antigen-binding memory B cells through fluorescence-activated cell sorting (FACS) or optofluidic platforms has been a major technical advancement ( Zhou et al., 2021 ). These techniques enable subsequent nested RT-PCR using primers designed to amplify naturally paired antibody heavy- and light-chain gene sequences from individual memory B cells ( Liao et al., 2009 ). Furthermore, high-throughput single-cell RNA and VDJ deep sequencing of BCR repertoires, coupled with bioinformatics analysis, have surpassed single-cell RT-PCR in terms of efficiently screening large pools of diverse memory B cells ( Cao et al., 2020 ). Notably, immunization using RBD/S proteins or direct infection of mice with genetically humanized immune systems has been demonstrated to generate complete HuNAbs against SARS-CoV-2 ( Hansen et al., 2020 ). This platform has successfully facilitated the cloning and screening of certain SARS-CoV-2-specific HuNAbs. Deep sequencing of variable regions and BCR repertoires has provided insights into the characteristics of human antibody heavy and light chain pairing ( Zhou et al., 2021 ). In addition, the combination of microfluidic-based techniques and bioinformatics analysis has the potential to further enhance the efficiency of identifying highly potent HuNAbs against specific viral antigens ( Hansen et al., 2020 ). These advancements hold implications for combating COVID-19 and other emerging infectious diseases.

Phage display has emerged as a widely utilized approach for cloning human antibodies. This technique involves two main stages: constructing an antibody gene library and screening for antibodies specific to a particular antigen ( Zhou et al., 2021 ). To generate a human Fab library, researchers can extract peripheral blood mononuclear cells (PBMCs) from individuals who have recovered from COVID-19. By amplifying the antibody gene pool using specific primers targeting the variable regions of the heavy and light chains, a phage library can be constructed ( Zhou et al., 2021 ). The library can also be synthesized using selected human germline immunoglobulin variable segments, with diversity introduced in the complementarity-determining regions (CDRs) through mutagenesis. The library can be screened using SARS-CoV-2 receptor-binding domain (RBD) as a bait to isolate RBD-specific human Fabs ( Zhou et al., 2021 ). These Fabs can then be further developed into full-length IgG1 antibodies for subsequent testing of their biochemical and functional properties. Although the phage display technique has some limitations, such as the unnatural pairing of heavy and light chains and a time-consuming panning procedure, it remains a valuable tool for obtaining antigen-specific antibodies, including potent SARS-CoV-2-specific human neutralizing antibodies (HuNAbs) ( Zhou et al., 2021 ). The SARS-CoV-2 spike protein (S protein) plays a crucial role in viral entry by binding to the ACE2 receptor on host cells ( Lv et al., 2020 ). The receptor-binding domain (RBD) within the S1 subunit of the S protein undergoes conformational changes, transitioning between “up” and “down” states. The “up” conformation is particularly targeted by HuNAbs since it represents the active state for ACE2 binding ( Zhou et al., 2021 ). Multiple neutralizing domains within the S1 region have been identified, including RBD and the N-terminal domain (NTD). HuNAbs can be classified into three main groups based on their binding sites ( Yuan et al., 2021 ). The first group, known as receptor-binding site (RBS) antibodies, targets epitopes within the RBD that overlap with the ACE2 binding site ( Brouwer et al., 2020 ). RBS antibodies can further be divided into subclasses based on their interaction angles with the viral RBD ( Yuan et al., 2021 ). The second group includes cross-reactive antibodies, such as CR3022, which recognize cryptic sites in the RBD and exhibit varying neutralizing activities against both SARS-CoV and SARS-CoV-2 ( Yuan et al., 2021 ). The third group is represented by antibodies like S309, which engage the RBD through an epitope containing the N343 glycan and can neutralize both SARS-CoV-2 and SARS-CoV ( Yuan et al., 2021 ).

Several anti-SARS-CoV-2 HuNAbs have been developed and undergone clinical trials. Paired HuNAbs have been formulated to enhance the breadth of neutralization against emerging SARS-CoV-2 variants and minimize the occurrence of HuNAb escape variants. HuNAb combinations under Emergency Use Authorization (EUA) exhibit live virus neutralization IC50 values below 0.1 µg/ml ( Jones et al., 2021 ). Typically, these paired antibodies consist of one RBS-A HuNAb along with either one RBS-B or one RBS-C HuNAb. For example, the combination of LY-CoV555 (bamlanivimab, BAM) and LY-CoV016 (etesevimab, ETE) from Eli Lilly has been administered to COVID-19 patients ( Zhou et al., 2021 ). The randomized phase 2/3 trial demonstrated a statistically significant reduction in viral load at day 11 among non-hospitalized patients with mild to moderate illness who received the combination therapy ( Gottlieb et al., 2021 ). Similarly, the REGN-CoV2 antibody cocktail (casirivimab, CAS, and imdevimab, IMD) showed a substantial reduction in viral loads in patients with baseline viral load higher than 107 copies/mL compared to the placebo group ( Hansen et al., 2020 ). Due to concerns regarding reduced neutralizing abilities against SARS-CoV-2 variants of concern (VOCs), ongoing clinical trials are investigating the efficacy of more broadly reactive HuNAbs, as well as next-generation antibodies such as bispecific antibodies and engineered antibodies. In addition, the development of BRII-196 and BRII-198, an antibody combination approved for clinical use in China, has shown promising results with 78% efficacy ( Zhou et al., 2021 ). SARS-CoV-2 virus has the ability to undergo mutations during its replication in humans. Unlike the SARS virus in 2003, which was eliminated, COVID-19 has persisted, leading to concerns about viral mutations that could evade immunity from natural infection or vaccination ( Romano et al., 2020 ). Several VOCs have emerged during the pandemic, including the Alpha, Beta, Gamma, Delta, and Omicron variants ( Chakraborty et al., 2022 ). These VOCs have mutations in the spike (S) glycoprotein of the virus, which can confer resistance to human neutralizing antibodies (HuNAbs) and increase viral transmissibility. The D614G mutation, found in all VOCs, enhances viral transmissibility ( Plante et al., 2021 ). Mutations such as E484K/A and amino acid deletions in the S protein reduce the potency of HuNAbs targeting specific regions ( Jangra et al., 2021 ). The emergence of these VOCs poses challenges to public health and vaccine responses.

Efforts have been focused on developing vaccines to combat COVID-19, and several vaccines have been approved for emergency use. These include mRNA-based vaccines, adenovirus-vectored vaccines, and inactivated vaccines ( Zhou et al., 2021 ). The efficacy rates reported for these vaccines mainly reflect their ability to prevent severe diseases, while their effectiveness against SARS-CoV-2 nasal infection and VOCs requires further investigation ( Zhou et al., 2021 ). Studies have shown reduced neutralization of VOCs by antibodies from vaccinated individuals, indicating the potential for decreased effectiveness against certain variants ( Wang et al., 2021 ). However, widespread vaccination campaigns have contributed to reduced hospitalizations, deaths, and infections in various countries. Future strategies for vaccination and immunotherapy should aim to provide broadly reactive immune protection against multiple variants ( Zhou et al., 2021 ). The role of mucosal immunity and tissue-resident memory T cells in the upper respiratory tract needs to be explored for long-term protection ( Baum et al., 2020 ). In addition, careful monitoring of antibody-mediated enhancement of SARS-CoV-2 infection and immunopathogenesis is crucial in the context of human infections and clinical management.

Synergy of Molecular Cloning and the CRISPR-Cas System in SARS-CoV-2 Treatment

COVID-19, caused by the SARS-CoV-2 virus, has triggered a global pandemic, with the emergence of highly transmissible and more fatal mutant strains. Effective management of this infectious disease requires a number of measures, including social distancing and isolation of confirmed cases. However, there is a pressing need for a highly sensitive diagnostic kit that enables rapid early detection ( Lou et al., 2022 ). Diagnostic approaches for COVID-19 involve immunological and molecular techniques. Immunological methods detect viral antigens or antibodies in the lungs or blood, aiding in disease understanding and transmission dynamics ( Mahase, 2020 ). Molecular methods primarily focus on nucleic acid detection, with RT-PCR being the gold standard due to its high sensitivity and accuracy ( Broughton et al., 2020 ). However, the limited availability of RT-PCR equipment and materials can lead to delays and false negative results, making it unsuitable for mass screening ( Broughton et al., 2020 , Li and Ren, 2020 ). Hence, there is a need for simplified and time-efficient diagnostic methods with high accuracy ( Xiang et al., 2020 ). The CRISPR-Cas system, with nucleic acid detection technologies like SHERLOCK (Cas13a), DETECTR (Cas12a), CDetection (Cas12b), and CAS14-DETECTR, offers a new avenue for pathogen screening ( Ding et al., 2021 ). In recent times, diverse detection methods combined with isothermal amplification and the CRISPR-Cas system have emerged as rapid diagnostic tools for detecting SARS-CoV-2 viral RNA. For instance, a visual analysis method called CLAP combining AUNP and Cas12a-assisted RT-LAMP demonstrated the ability to detect low levels of SARS-CoV-2 RNA rapidly ( Lou et al., 2022 ). In addition, engineered Cas12a enzyme-based LAMP showed promising results in detecting wild-type and mutant SARS-CoV-2 within a short time frame ( Lou et al., 2022 ). Cas13a crRNAs can specifically target SARS-CoV-2 and related coronaviruses, and when combined with a generic autonomous enzyme-free hybridization chain reaction (HCR), they offer a detection method for monitoring virus transmission via objects ( Yang et al., 2021 ). The CRISPR-Cas13 amplification strategy holds potential for efficient surveillance of SARS-CoV-2 transmission ( Lou et al., 2022 ). Viral control over host cells has long intrigued virologists due to the limited number of viral genes. The explanation lies, in part, in the utilization of host factors by viruses during their life cycle. Identifying these host factors that either facilitate or impede novel virus replication can uncover potential targets for antiviral therapies ( Deol et al., 2022 ). A number of techniques, including forward genetic screens, are employed to study virus-host interactions. Although the CRISPR/Cas9 system is not the first tool for genetic screens, it has emerged as a highly robust method. Genome-wide CRISPR screens are being conducted to identify host genes involved in SARS-CoV-2 replication. For instance, Wang et al. identified ACE-2 as the entry receptor and highlighted TMEM106B, VAC14, cholesterol regulators, and exocyst subunits as additional host factors supporting SARS-CoV-2 infection, thus offering potential targets for antiviral strategies ( Wang et al., 2021 ). Other researchers have also employed CRISPR-based screens to identify candidate host genes for SARS-CoV-2, paving the way for the development of CRISPR/Cas9-mediated antiviral therapeutics ( Daniloski et al., 2021 , Zhu et al., 2021 ). Huang et al. devised a CRISPR-based diagnostic approach that utilizes customized CRISPR Cas12a/gRNA complex and fluorescent probes to detect target amplifiers generated by standard RT-PCR or isothermal recombinase polymerase amplification (RPA). This method enables sensitive detection at locations where the RT-PCR system necessary for qPCR diagnosis is unavailable ( Huang et al., 2021 ). The technique exhibited high sensitivity, with a reaction time of approximately 50 minutes and a detection limit of two samples per sample.

The diagnostic results obtained using the CRISPR analysis from nasal swab samples of suspected COVID-19 patients were comparable to quantitative RT-PCR tests and outperformed standard clinical laboratory tests. With no specific treatment available for emerging infectious diseases like COVID-19, it is crucial to explore effective diagnostic and therapeutic approaches ( Lou et al., 2022 ). A number of antiviral approaches focus on inhibiting different stages of the viral life cycle by targeting structural or non-structural components. CRISPR/Cas systems have gained significant attention as a means to limit viral replication by specifically targeting the viral genome. CRISPR/Cas9, in particular, has demonstrated its potential as an antiviral strategy against DNA viruses, both in vitro and in vivo ( Lee, 2019 ). Similarly, the CRISPR/Cas12a system has shown promise in inactivating integrated HIV DNA genomes, surpassing Cas9 in HIV inhibition. On the contrary, the CRISPR/Cas13 system has emerged as a novel RNA-guided RNA targeting system, capable of recognizing and degrading the genomic RNA of SARS-CoV-2, thereby impeding virus replication. Recent research has demonstrated the efficacy of CRISPR-Cas13 in safeguarding host bacterial cells against phage infection through specific cRNA ( Yan et al., 2019 ). This strategy can potentially be leveraged to design therapeutic drugs targeting the single-stranded RNA genomes of emerging infectious diseases. The Cas13 enzymes utilize a short hairpin crRNA to recognize specific sequences on the target RNA, and unlike Cas9, they do not require a protospacer adjacent motif (PAM) sequence ( Burmistrz et al., 2020 ). Some subtypes of Cas13 also exhibit collateral cleavage activity, resulting in non-specific cleavage of target and non-target RNAs. This property has been utilized in diagnostic applications such as SHERLOCK. Different subtypes of Cas13 have been explored for their antiviral potential. Abbott et al. introduced a CRISPR-Cas13-based strategy known as PAC-MAN (Prophylactic Antiviral CRISPR in Human Cells), which effectively degrades SARS-CoV-2 RNA in cells by identifying functional crRNA specifically targeting SARS-CoV-2 ( Lou et al., 2022 ). This method reduces viral load within cells, and a small set of six crRNAs can target over 90% of coronaviruses, making PAC-MAN a promising pan-coronavirus suppression strategy ( Abbott et al., 2020 ). The development of CRISPR-based tools for the treatment of emerging infectious diseases holds great research prospects.

The primary drawback hindering the extensive use of the CRISPR-Cas system lies in its inability to accurately identify specific nucleic acids for diagnostic and therapeutic purposes ( Doudna, 2020 ). To address this issue, one approach is to enhance the specificity of target nucleic acids by modifying the CRISPR protein, thus minimizing off-target effects ( Naeem et al., 2020 ). Bioinformatics methods are commonly employed to detect off-target effects, and advancements in Cas9 have shown promise in reducing such effects ( Lou et al., 2022 , Coelho et al., 2020 ). Future research should focus on investigating off-target effects and refining detection techniques. In utilizing the CRISPR-Cas system for the prevention and control of infectious diseases, delivery tools play a crucial role in targeting cells within the body ( Lou et al., 2022 ). However, the carrier function of CRISPR is limited by the size of viral genes, and most Cas proteins are relatively large in molecular weight ( Liu et al., 2020 ). Overcoming this challenge necessitates the search for low molecular weight Cas proteins ( Luther et al., 2018 ). Moreover, when Cas proteins derived from prokaryotes are administered to the human body, they can elicit an immune response and trigger the production of specific antibodies, which interfere with CRISPR’s immune response ( Lou et al., 2022 ). Enhancing the efficiency of the CRISPR-Cas system is vital to mitigate the immune response in future CRISPR tool development. Cas9 and Cas12 proteins exhibit PAM-dependent recognition and cleavage of dsDNA, allowing them to target specific gene sequences within target genes ( Karvelis et al., 2020 ). However, some highly specific sequences may not be accessible, emphasizing the need for further enhancement of the CRISPR tool for target selection ( Lou et al., 2022 ). In addition, the presence of RNA enzymes widely in existence leads to RNA instability, thereby affecting the diagnostic efficacy of CRISPR.

There has been considerable progress in molecular cloning techniques in research laboratories as well as widespread operation in medical microbiology. While DNA-based procedures have improved the diagnosis of diseases, conventional cell factories have drained their competence ( Khan et al., 2016 ). It is obligatory to discover and integrate new production systems into the production process. In addition, a deeper understanding of the physiology of host cells and their responses to stress would enable enhanced tools by gene manipulation either at the genetic or at the metabolic levels for growing yield and eminence ( Khan et al., 2016 ). The progressions in recombinant DNA technology, microbiology, genomics, bioinformatics, and related fields have, however, contributed significantly to the understanding of the pathogenic mechanisms underlying the microbial infectious diseases and how pathogens interrelate with hosts. Several new vaccine strategies have been established, as outcomes of these advances have produced promising results. Masses of children, globally, still die from infectious diseases notwithstanding the existence of currently available vaccines. Therefore, it is essential to understand the trials involved in developing recombinant vaccines and estimate the stability between charge, remunerations, and hazard before bringing a vaccine applicant to the hospital. As a result of this approach, patients with unrecognized or hard-to-diagnose infections will be identified and treated punctually, resulting in reduced stays and a reduction in iatrogenic events. Considering the relative costs of novel diagnostics compared to existing standards, public reimbursements will need to be carefully explored. Hence, molecular cloning can be delineated as a scientific effort to treat a lengthy list of diseases that have claimed a lot of human lives in the course of long period.

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Datta, N. (2024) 'Unlocking the Power of Molecular Cloning: Revolutionizing Medical Microbiology Procedures', University of Michigan Undergraduate Research Journal . 17(0) doi: 10.3998/umurj.5509

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• Methodology
• Open access
• Published: 19 January 2015

Molecular cloning using polymerase chain reaction, an educational guide for cellular engineering

• Sayed Shahabuddin Hoseini 1 , 2 &
• Martin G Sauer 1 , 2 , 3  

Journal of Biological Engineering volume  9 , Article number:  2 ( 2015 ) Cite this article

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Over the last decades, molecular cloning has transformed biological sciences. Having profoundly impacted various areas such as basic science, clinical, pharmaceutical, and environmental fields, the use of recombinant DNA has successfully started to enter the field of cellular engineering. Here, the polymerase chain reaction (PCR) represents one of the most essential tools. Due to the emergence of novel and efficient PCR reagents, cloning kits, and software, there is a need for a concise and comprehensive protocol that explains all steps of PCR cloning starting from the primer design, performing PCR, sequencing PCR products, analysis of the sequencing data, and finally the assessment of gene expression. It is the aim of this methodology paper to provide a comprehensive protocol with a viable example for applying PCR in gene cloning.

Exemplarily the sequence of the tdTomato fluorescent gene was amplified with PCR primers wherein proper restriction enzyme sites were embedded. Practical criteria for the selection of restriction enzymes and the design of PCR primers are explained. Efficient cloning of PCR products into a plasmid for sequencing and free web-based software for the consecutive analysis of sequencing data is introduced. Finally, confirmation of successful cloning is explained using a fluorescent gene of interest and murine target cells.

Conclusions

Using a practical example, comprehensive PCR-based protocol with important tips was introduced. This methodology paper can serve as a roadmap for researchers who want to quickly exploit the power of PCR-cloning but have their main focus on functional in vitro and in vivo aspects of cellular engineering.

Various techniques were introduced for assembling new DNA sequences [ 1 – 3 ], yet the use of restriction endonuclease enzymes is the most widely used technique in molecular cloning. Whenever compatible restriction enzyme sites are available on both, insert and vector DNA sequences, cloning is straightforward; however, if restriction sites are incompatible or if there is even no restriction site available in the vicinity of the insert cassette, cloning might become more complex. The use of PCR primers, in which compatible restriction enzyme sites are embedded, can effectively solve this problem and facilitate multistep cloning procedures.

Although PCR cloning has been vastly used in biological engineering [ 4 – 8 ], practical guides explaining all necessary steps and tips in a consecutive order are scarce. Furthermore, the emergence of new high-fidelity DNA polymerases, kits, and powerful software makes the process of PCR cloning extremely fast and efficient. Here we sequentially explain PCR cloning from the analysis of the respective gene sequence, the design of PCR primers, performing the PCR procedure itself, sequencing the resulting PCR products, analysis of sequencing data, and finally the cloning of the PCR product into the final vector.

Results and discussion

Choosing proper restriction enzymes based on defined criteria.

In order to proceed with a concise example, tdTomato fluorescent protein was cloned into an alpharetroviral vector. Consecutively, a murine leukemia cell line expressing tdTomato was generated. This cell line will be used to track tumor cells upon injection into mice in preclinical immunotherapy studies. However, this cloning method is applicable to any other gene. To begin the cloning project, the gene of interest (GOI) should be analyzed. First, we check whether our annotated sequence has a start codon (ATG, the most common start codon) and one of the three stop codons (TAA, TAG, TGA). In case the gene was previously manipulated or fused to another gene (e.g. via a 2A sequence), it happens that a gene of interest might not have a stop codon [ 9 ]. In such cases, a stop codon needs to be added to the end of your annotated sequence. It is also beneficial to investigate whether your GOI contains an open reading frame (ORF). This is important since frequent manipulation of sequences either by software or via cloning might erroneously add or delete nucleotides. We use Clone Manager software (SciEd) to find ORFs in our plasmid sequences; however, there are several free websites you can use to find ORFs including the NCBI open reading frame finder ( http://www.ncbi.nlm.nih.gov/gorf/gorf.html ).

The tdTomato gene contains ATG start codon and TAA stop codon (Figure  1 ). The size of the tdTomato gene is 716 bp.

Overview of the start and the end of the gene of interest. (A) The nucleotide sequences at the start and the end of the tdTomato gene are shown. The coding strand nucleotides are specified in bold (B) The nucleotide sequences of the forward and reverse primers containing proper restriction enzyme sites and the Kozak sequence are shown.

In a next step, PCR primers that include proper restriction enzyme sites need to be designed for the amplification of the GOI. Several criteria should be considered in order to choose the optimal restriction enzymes. First, binding sites for restriction enzymes should be ideally available at a multiple cloning site within the vector. Alternatively they can be located downstream of the promoter in your vector sequence. Restriction enzymes should be single cutters (single cutters target one restriction site only within a DNA sequence) (Figure  2 A). If they are double or multiple cutters, they should cut within a sequence that is not necessary for proper functioning of the vector plasmid and will finally be removed (Figure  2 B). It is also possible to choose one double cutter or multiple cutter enzymes cutting the vector downstream of the promoter and also not within a vital sequence of the plasmid (Figure  2 C). Double cutter or multiple cutter enzymes have two or more restriction sites on a DNA sequence, respectively. Cutting the vector with double or multiple cutters would give rise to two identical ends. In such a case, the insert cassette should also contain the same restriction enzyme sites on both of its ends. Therefore, when the insert and vector fragments are mixed in a ligation experiment, the insert can fuse to the vector in either the right orientation (from start codon to stop codon) or reversely (from stop codon to start codon). A third scenario can occur, if the vector fragment forms a self-ligating circle omitting the insert at all. Once the DNA has been incubated with restriction enzymes, dephosphorylation of the 5′ and 3′ ends of the vector plasmid using an alkaline phosphatase enzyme will greatly reduce the risk of self-ligation [ 10 ]. It is therefore important to screen a cloning product for those three products (right orientation, reverse orientation, self-ligation) after fragment ligation.

Choosing proper restriction enzymes based on defined criteria for PCR cloning. (A) Two single-cutter restriction enzymes (E1 and E2) are located downstream of the promoter. (B) E1 and E2 restriction enzymes cut the plasmid downstream of the promoter several (here two times for each enzyme) times. (C) The E1 restriction enzyme cuts the plasmid downstream of the promoter more than once. (D) The PCR product, which contains the tdTomato gene and the restriction enzyme sites, was run on a gel before being extracted for downstream applications.

Second, due to higher cloning efficiency using sticky-end DNA fragments, it is desirable that at least one (better both) of the restriction enzymes is a so-called sticky-end cutter. Sticky end cutters cleave DNA asymmetrically generating complementary cohesive ends. In contrast, blunt end cutters cut the sequence symmetrically leaving no overhangs. Cloning blunt-end fragments is more difficult. Nevertheless, choosing a higher insert/vector molar ratio (5 or more) and the use 10% polyethylene glycol (PEG) can improve ligation of blunt-end fragments [ 11 ].

Third, some restriction enzymes do not cut methylated DNA. Most of the strains of E. coli contain Dam or Dcm methylases that methylate DNA sequences. This makes them resistant to methylation-sensitive restriction enzymes [ 12 ]. Since vector DNA is mostly prepared in E. coli , it will be methylated. Therefore avoiding methylation-sensitive restriction enzymes is desirable; however, sometimes the isoschizomer of a methylation-sensitive restriction enzyme is resistant to methylation. For example, the Acc 65I enzyme is sensitive while its isoschizomer kpn I is resistant to methylation [ 13 ]. Isoschizomers are restriction enzymes that recognize the same nucleotide sequences. If there remains no other option than using methylation-sensitive restriction enzymes, the vector DNA needs to be prepared in dam − dcm − E. coli strains. A list of these strains and also common E. coli host strains for molecular cloning is summarized in Table  1 . Information regarding the methylation sensitivity of restriction enzymes is usually provided by the manufacturer.

Fourth, it makes cloning easier if the buffer necessary for the full functionality of restriction enzymes is the same because one can perform double restriction digest. This saves time and reduces the DNA loss during purification. It may happen that one of the restriction enzymes is active in one buffer and the second enzyme is active in twice the concentration of the same buffer. For example the Nhe I enzyme from Thermo Scientific is active in Tango 1X buffer (Thermo Scientific) and Eco R1 enzyme is active in Tango 2X buffer (Thermo Scientific). In such cases, the plasmid DNA needs to be first digested by the enzyme requiring the higher buffer concentration (here Eco R1). This will be followed by diluting the buffer for the next enzyme (requiring a lower concentration (here Nhe I)) in the same buffer. However, the emergence of universal buffers has simplified the double digest of DNA sequences [ 15 ]. In our example the vector contains the Age I and Sal I restriction sites. These enzyme sites were used for designing PCR primers (Figure  1 ). It is essential for proper restriction enzyme digestion that the plasmid purity is high. DNA absorbance as measured by a spectrophotometer can be used to determine the purity after purification. DNA, proteins, and solvents absorb at 260 nm, 280 nm, and 230 nm, respectively. An OD 260/280 ratio of >1.8 and an OD 260/230 ratio of 2 to 2.2 is considered to be pure for DNA samples [ 16 ]. The OD 260/280 and 260/230 ratios of our exemplary plasmid preparations were 1.89 and 2.22, respectively. We observed that the purity of the gel-extracted vector and insert DNA fragments were lower after restriction digest; ligation works even in such cases, however, better results can be expected using high-purity fragments.

The following plasmid repository website can be useful for the selection of different vectors (viral expression and packaging, empty backbones, fluorescent proteins, inducible vectors, epitope tags, fusion proteins, reporter genes, species-specific expression systems, selection markers, promoters, shRNA expression and genome engineering): http://www.addgene.org/browse/ .

A collection of cloning vectors of E. coli is available under the following website: http://www.shigen.nig.ac.jp/ecoli/strain/cvector/cvectorExplanation.jsp .

Designing cloning primers based on defined criteria

For PCR primer design, check the start and stop codons of your GOI. Find the sequence of the desired restriction enzymes (available on the manufacturers’ websites) for the forward primer (Figure  3 A). It needs to be located before the GOI (Figure  1 B). The so-called Kozak sequence is found in eukaryotic mRNAs and improves the initiation of translation [ 17 ]. It is beneficial to add the Kozak sequence (GCCACC) before the ATG start codon since it increases translation and expression of the protein of interest in eukaryotes [ 18 ]. Therefore, we inserted GCCACC immediately after the restriction enzyme sequence Age I and before the ATG start codon. Then, the first 18 to 30 nucleotides of the GOI starting from the ATG start codon are added to the forward primer sequence. These overlapping nucleotides binding to the template DNA determine the annealing temperature (Tm). The latter is usually higher than 60°C. Here, we use Phusion high-fidelity DNA polymerase (Thermo Scientific). You can use the following websites for determination of the optimal Tm: http://www.thermoscientificbio.com/webtools/tmc/ .

Designing primers based on defined criteria for PCR cloning. (A-B) Sequences of the forward and the reverse primer are depicted. The end of the coding strand is to be converted into the reverse complement format for the reverse primer design. For more information, please see the text.

https://www.neb.com/tools-and-resources/interactive-tools/tm-calculator .

The Tm of our forward primer is 66°C.

Choose the last 18 to 30 nucleotides including the stop codon of your GOI for designing the reverse primer (Figure  3 B). Then calculate the Tm for this sequence which should be above 60°C and close to the Tm of the forward primer. Tm of the overlapping sequence of our reverse primer was 68°C. Then, add the target sequence of the second restriction enzyme site (in this case Sal I) immediately after the stop codon. Finally, convert this assembled sequence to a reverse-complement sequence. The following websites can be used to determine the sequence of the reverse primer:

http://reverse-complement.com/

http://www.bioinformatics.org/sms/rev_comp.html This is important since the reverse primer binds the coding strand and therefore its sequence (5′ → 3′) must be reverse-complementary to the sequence of the coding strand (Figure  1 A).

Performing PCR using proofreading polymerases

Since the PCR reaction follows logarithmic amplification of the target sequence, any replication error during this process will be amplified. The error rate of non-proofreading DNA polymerases, such as the Taq polymerase, is about 8 × 10 −6 errors/bp/PCR cycle [ 19 ]; however, proofreading enzymes such as Phusion polymerase have a reported error rate of 4.4 × 10 −7 errors/bp/PCR cycle. Due to its superior fidelity and processivity [ 20 – 22 ], the Phusion DNA polymerase was used in this example. It should be noted that Phusion has different temperature requirements than other DNA polymerases. The primer Tm for Phusion is calculated based on the Breslauer method [ 23 ] and is higher than the Tm using Taq or pfu polymerases. To have optimal results, the Tm should be calculated based on information found on the website of the enzyme providers. Furthermore, due to the higher speed of Phusion, 15 to 30 seconds are usually enough for the amplification of each kb of the sequence of interest.

After the PCR, the product needs to be loaded on a gel (Figure  2 D). The corresponding band needs to be cut and the DNA extracted. It is essential to sequence the PCR product since the PCR product might include mutations. There are several PCR cloning kits available some of which are shown in Table  2 . We used the pJET1.2/blunt cloning vector (Thermo Scientific, patent publication: US 2009/0042249 A1, Genbank accession number EF694056.1) and cloned the PCR product into the linearized vector. This vector contains a lethal gene ( eco47IR ) that is activated in case the vector becomes circularized. However, if the PCR product is cloned into the cloning site within the lethal gene, the latter is disrupted allowing bacteria to grow colonies upon transformation. Circularized vectors not containing the PCR product express the toxic gene, which therefore kills bacteria precluding the formation of colonies. Bacterial clones are then to be cultured, plasmid DNA consecutively isolated and sequenced. The quality of isolated plasmid is essential for optimal sequencing results. We isolated the plasmid DNA from a total of 1.5 ml cultured bacteria (yield 6 μg DNA; OD 260/280 = 1.86; OD 260/230 = 2.17) using a plasmid mini-preparation kit (QIAGEN). The whole process of PCR, including cloning of the PCR product into the sequencing vector and transfection of bacteria with the sequencing vector can be done in one day. The next day, bacterial clones will be cultured overnight before being sent for sequencing.

Analysis of sequencing data

Sequencing companies normally report sequencing data as a FASTA file and also as ready nucleotide sequences via email. For sequence analysis, the following websites can be used:

http://blast.ncbi.nlm.nih.gov/Blast.cgi

http://xylian.igh.cnrs.fr/bin/align-guess.cgi

Here we will focus on the first website. On this website page, click on the “nucleotide blast” option (Figure  4 A). A new window opens. By default, the “blastn” (blast nucleotide sequences) option is marked (Figure  4 B). Then check the box behind “Align two or more sequences”. Now two boxes will appear. In the “Enter Query Sequence” box (the upper box), insert the desired sequence of your gene of interest, which is flanked by the restriction sites you have already designed for your PCR primers. In the “Enter Subject Sequence” box (the lower box), enter the sequence or upload the FASTA file you have received from the sequencing company. Then click the “BLAST” button at the bottom of the page. After a couple of seconds, the results will be shown on another page. A part of the alignment data is shown in Figure  4 C. For interpretation, the following points should be considered: 1) the number of identical nucleotides (shown under the “Identities” item) must be equal to the nucleotide number of your gene of interest. In our example, the number of nucleotides of the tdTomato gene together with those of the restriction enzyme sites and the Kozak sequence was 735. This equals the reported number (Figure  4 C). 2) The sequence identity (under the “Identities” item) should be 100%. Occasionally, the sequence identity is 100% but the number of identical nucleotides is lower than expected. This can happen if one or more of the initial nucleotides are absent. Remember, all sequencing technologies have an error rate. For Sanger sequencing, this error rate is reported to range from 0.001% to 1% [ 30 – 33 ]. Nucleotide substitution, deletion or insertion can be identified by analyzing the sequencing results [ 34 ]. Therefore, if the sequence identity does not reach 100%, the plasmid should be resequenced in order to differentiate errors of the PCR from simple sequencing errors. 3) Gaps (under the “Gaps” item) should not be present. If gaps occur, the plasmid should be resequenced.

Sequence analysis of the PCR product using the NCBI BLAST platform. (A) On the NCBI BLAST webpage, the “nucleotide blast” option is chosen (marked by the oval line). (B) The “blastn” option appears by default (marked by the circle). The sequence of the gene of interest (flanked by the restriction sites as previously designed for the PCR primers) and the PCR product are to be inserted to the “Enter Query Sequence” and “Enter Subject Sequence” boxes. Sequences can also be uploaded as FASTA files. (C) Nucleotide alignment of the first 60 nucleotides is shown. Two important items for sequence analysis are marked by oval lines.

The average length of a read, or read length, is at least 800 to 900 nucleotides for Sanger sequencing [ 35 ]. For the pJET vector one forward and one reverse primer need to be used for sequencing the complete gene. These primers can normally cover a gene size ranging up to 1800 bp. If the size of a gene is larger than 1800, an extra primer should be designed for each 800 extra nucleotides. Since reliable base calling does not start immediately after the primer, but about 45 to 55 nucleotides downstream of the primer [ 36 ], the next forward primer should be designed to start after about 700 nucleotides from the beginning of the gene. Different websites, including the following, can be used to design these primers:

http://www.ncbi.nlm.nih.gov/tools/primer-blast/

http://www.yeastgenome.org/cgi-bin/web-primer

http://www.genscript.com/cgi-bin/tools/sequencing_primer_design

Being 735 bp in length, the size of the PCR product in this example was well within the range of the pJET sequencing primers.

After choosing the sequence-verified clone, vector and insert plasmids were digested by the Age I and Sal I restriction enzymes (Figure  5 ). This was followed by gel purification and ligation of the fragments. Transformation of competent E. coli with the ligation mixture yielded several clones that were screened by restriction enzymes. We assessed eight clones, all of which contained the tdTomato insert (Figure  6 ). It is important to pick clones that are large. Satellite clones might not have the right construct. We used a fast plasmid mini-preparation kit (Zymo Research) to extract the plasmid from 0.6 ml bacterial suspension. The yield and purity were satisfying for restriction enzyme-based screening (2.3 μg DNA; OD 260/280 = 1.82; OD 260/230 = 1.41). For large-scale plasmid purification, a maxi-preparation kit (QIAGEN) was used to extract the plasmid from 450 ml of bacterial culture (yield 787 μg DNA; OD 260/280 = 1.89; OD 260/230 = 2.22). The expected yield of a pBR322-derived plasmid isolation from 1.5 ml and 500 ml bacterial culture is about 2-5 μg and 500-4000 μg of DNA, respectively [ 37 ].

Vector and insert plasmid maps A) Illustration of the CloneJET plasmid containing the PCR product. Insertion of the PCR product in the cloning site of the plasmid disrupts the integrity of the toxic gene eco47IR and allows the growth of transgene positive clones. The plasmid was cut with the Age I and Sal I enzymes generating two fragments of 3 kb and 0.7 kb in size. The 0.7 kb fragment (tdTomato gene) was used as the insert for cloning. (B) Illustration of the vector plasmid. The plasmid was cut with the Age I and Sal I enzymes generating two fragments of 4.9 kb and 0.7 kb in size. The 4.9 kb fragment was used as the vector for cloning. AMP: Ampicillin resistance gene; PRE: posttranscriptional regulatory element; MPSV: myeloproliferative sarcoma virus promoter.

Screening of the final plasmid with restriction enzymes. Illustration of the final plasmid is shown. For screening, the plasmid was cut with the Bsiw I enzyme generating two fragments of 4.8 kb and 0.8 kb in size. AMP: Ampicillin resistance gene; PRE: posttranscriptional regulatory element; MPSV: myeloproliferative sarcoma virus promoter.

Some plasmids tend to recombine inside the bacterial host creating insertions, deletions and recombinations [ 38 ]. In these cases, using a recA-deficient E. coli can be useful (Table  1 ). Furthermore, if the GOI is toxic, incubation of bacteria at lower temperatures (25-30°C) and using ABLE C or ABLE K strains might circumvent the problem.

Viral production and transduction of target cells

To investigate the in vitro expression of the cloned gene, HEK293T cells were transfected with plasmids encoding the tdTomato gene, alpharetroviral Gag/Pol, and the vesicular stomatitis virus glycoprotein (VSVG) envelope. These cells, which are derived from human embryonic kidney, are easily cultured and readily transfected [ 39 ]. Therefore they are extensively used in biotechnology and gene therapy to generate viral particles. HEK293T cells require splitting every other day using warm medium. They should not reach 100% confluency for optimal results. To have good transfection efficiency, these cells need to be cultured for at least one week to have them in log phase. Transfection efficiency was 22%, as determined based on the expression of tdTomato by fluorescence microscopy 24 hours later (Figure  7 A-B). To generate a murine leukemia cell line expressing the tdTomato gene for immunotherapy studies, C1498 leukemic cells were transduced with freshly harvested virus (36 hours of transfection). Imaging studies (Figure  7 C) and flow cytometric analysis (Figure  7 D) four days after transduction confirmed the expression of tdTomato in the majority of the cells.

Assessing in vitro expression of the cloned gene. (A, B) HEK293T cells were transfected with Gag/Pol, VSVG, and tdTomato plasmids. The expression of the tdTomato gene was assessed using a fluorescence microscope. Fluorescent images were superimposed on a bright-field image for the differentiation of positively transduced cells. Transfection efficiency was determined based on the expression of tdTomato after 24 hours. Non-transfected HEK293T cells were used as controls (blue histogram). (C, D) The murine leukemia cell line C1498 was transduced with fresh virus. Four days later, transgene expression was assessed by fluorescence microscopy (C) and flow cytometry (D) . Non-transduced C1498 cells were used as controls (blue histogram). Scale bars represent 30 μm.

In this manuscript, we describe a simple and step-by-step protocol explaining how to exploit the power of PCR to clone a GOI into a vector for genetic engineering. Several PCR-based creative methods have been developed being extremely helpful for the generation of new nucleotide sequences. This includes equimolar expression of several proteins by linking their genes via a self-cleaving 2A sequence [ 40 , 41 ], engineering fusion proteins, as well as the use of linkers for the design of chimeric proteins [ 42 – 44 ]. Furthermore, protein tags [ 45 , 46 ] and mutagenesis (site-directed, deletions, insertions) [ 47 ] have widened the applications of biological engineering. The protocol explained in this manuscript covers for most situations of PCR-assisted cloning; however, alternative PCR-based methods are available being restriction enzyme and ligation independent [ 6 , 48 – 51 ]. They are of special interest in applications where restriction enzyme sites are lacking; nevertheless, these methods might need several rounds of PCR or occasionally a whole plasmid needs to be amplified. In such cases, the chance of PCR errors increases and necessitates sequencing of multiple clones. In conclusion, this guideline assembles a simple and straightforward protocol using resources that are tedious to collect on an individual basis thereby trying to minimize errors and pitfalls from the beginning.

Cell lines and media

The E. coli HB101 was used for the preparation of plasmid DNA. The bacteria were cultured in Luria-Bertani (LB) media. Human embryonic kidney (HEK) 293 T cells were cultured in Dulbecco’s Modified Eagle medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 1 mM sodium pyruvate, 0.1 mM nonessential amino acids, 2 mM L-glutamine, 100 mg/ml streptomycin, and 100 units/ml penicillin. A myeloid leukemia cell line C1498 [ 52 ], was cultured in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with the same reagents used for DMEM. Cells were split every other day to keep them on log phase.

Plasmids, primers, PCR and sequencing

A plasmid containing the coding sequence of the tdTomato gene, plasmid containing an alpha-retroviral vector, and plasmids containing codon-optimized alpharetroviral gag/pol [ 53 ] were kindly provided by Axel Schambach (MHH Hannover, Germany). A forward (5′- ACCGGTGCCACCATGGCCACAACCATGGTG-3′) and a reverse (5′-GTCGACTTACTTGTACAGCTCGTCCATGCC-3′) primer used for the amplification of the tdTomato gene were synthesized by Eurofins Genomics (Ebersberg, Germany).

The optimal buffers for enzymes or other reagents were provided by the manufacturers along with the corresponding enzymes or inside the kits. If available by the manufacturers, the pH and ingredients of buffers are mentioned. Primers were dissolved in ultrapure water at a stock concentration of 20 pmol/μl. The template plasmid was diluted in water at a stock concentration of 50 ng/μl. For PCR, the following reagents were mixed and filled up with water to a total volume of 50 μl: 1 μl plasmid DNA (1 ng/μl final concentration), 1.25 μl of each primer (0.5 pmol/μl final concentration for each primer), 1 μL dNTP (10 mM each), 10 μl of 5X Phusion HF buffer (1X buffer provides 1.5 mM MgCl2), and 0.5 μl Phusion DNA polymerase (2U/μl, Thermo Scientific).

PCR was performed using a peqSTAR thermocycler (PEQLAB Biotechnologie) at: 98°C for 3 minutes; 25 cycles at 98°C for 10 seconds, 66°C for 30 seconds, 72°C for 30 seconds; and 72°C for 10 minutes. To prepare a 0.8% agarose gel, 0.96 g agarose (CARL ROTH) was dissolved in 120 ml 1X TAE buffer (40 mM Tris, 20 mM acetic acid, 1 mM EDTA, pH of 50X TAE: 8.4) and boiled for 4 minutes. Then 3 μl SafeView nucleic acid stain (NBS Biologicals) was added to the solution and the mixture was poured into a gel-casting tray.

DNA was mixed with 10 μl loading dye (6X) (Thermo Scientific) and loaded on the agarose gel (CARL ROTH) using 80 V for one hour in TAE buffer. The separated DNA fragments were visualized using an UV transilluminator (365 nm) and quickly cut to minimize the UV exposure. DNA was extracted from the gel slice using Zymoclean™ Gel DNA Recovery Kit (Zymo Research). The concentration of DNA was determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific).

For sequence validation, the PCR product was subcloned using CloneJET PCR cloning kit (Thermo Scientific). 1 μl of blunt vector (50 ng/μl), 50 ng/μl of the PCR product, and 10 μl of 2X reaction buffer (provided in the kit) were mixed and filled with water to a total volume of 20 μl. 1 μl of T4 DNA ligase (5 U/μl) was added to the mixture, mixed and incubated at room temperature for 30 minutes. For bacterial transfection, 10 μl of the mixture was mixed with 100 μl of HB101 E. coli competent cells and incubated on ice for 45 minutes. Then the mixture was heat-shocked (42°C/2 minutes), put on ice again (5 minutes), filled up with 1 ml LB medium and incubated in a thermomixer (Eppendorf) for 45 minutes/37°C/450RPM. Then the bacteria were spun down for 4 minutes. The pellet was cultured overnight at 37°C on an agarose Petri dish containing 100 μg/mL of Ampicillin. The day after, colonies were picked and cultured overnight in 3 ml LB containing 100 μg/mL of ampicillin.

After 16 hours (overnight), the plasmid was isolated from the cultured bacteria using the QIAprep spin miniprep kit (QIAGEN) according to the manufacturer’s instructions. 720 to 1200 ng of plasmid DNA in a total of 12 μl water were sent for sequencing (Seqlab) in Eppendorf tubes. The sequencing primers pJET1.2-forward (5′-CGACTCACTATAGGGAG-3′), and pJET1.2-reverse (5′-ATCGATTTTCCATGGCAG-3′), were generated by the Seqlab Company (Göttingen, Germany). An ABI 3730XL DNA analyzer was used by the Seqlab Company to sequence the plasmids applying the Sanger method. Sequence results were analyzed using NCBI Blast as explained in the Results and discussion section.

Manipulation of DNA fragments

For viewing plasmid maps, Clone Manager suite 6 software (SciEd) was used. Restriction endonuclease enzymes (Thermo Scientific) were used to cut plasmid DNA. 5 μg plasmid DNA, 2 μl buffer O (50 mM Tris–HCl (pH 7.5 at 37°C), 10 mM MgCl2, 100 mM NaCl, 0.1 mg/mL BSA, Thermo Scientific), 1 μl Sal I (10 U), and 1 μl AgeI (10 U) were mixed in a total of 20 μl water and incubated (37°C) overnight in an incubator to prevent evaporation and condensation of water under the tube lid. The next day, DNA was mixed with 4 μl loading dye (6X) (Thermo Scientific) and run on a 0.8% agarose gel at 80 V for one hour in TAE buffer. The agarose gel (120 ml) contained 3 μl SafeView nucleic acid stain (NBS Biologicals). The bands were visualized on a UV transilluminator (PEQLAB), using a wavelength of 365 nm, and quickly cut to minimize the UV damage. DNA was extracted from the gel slices using the Zymoclean™ gel DNA recovery kit (Zymo Research). The concentration of DNA was determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific).

For the ligation of vector and insert fragments, a ligation calculator was designed (the Excel file available in the Additional file 1 ) for easy calculation of the required insert and vector volumes. The mathematical basis of the calculator is inserted into the excel spreadsheet. The size and concentration of the vector and insert fragments and the molar ratio of vector/insert (normally 1:3) must be provided for the calculation. Calculated amounts of insert (tdTomato) and vector (alpha-retroviral backbone) were mixed with 2 μl of 10X T4 ligase buffer (400 mM Tris–HCl, 100 mM MgCl2, 100 mM DTT, 5 mM ATP (pH 7.8 at 25°C), Thermo Scientific), 1 μl of T4 ligase (5 U/μl, Thermo Scientific), filled up to 20 μl using ultrapure water and incubated overnight at 16°C. The day after, HB101 E. coli was transfected with the ligation mixture as mentioned above. The clones were picked and consecutively cultured for one day in LB medium containing ampicillin. Plasmid DNA was isolated using Zyppy™ plasmid miniprep kit (Zymo Research) and digested with proper restriction enzymes for screening. Digested plasmids were mixed with the loading dye and run on an agarose gel as mentioned above. The separated DNA fragments were visualized using a Gel Doc™ XR+ System (BIO-RAD) and analyzed by the Image Lab™ software (BIO-RAD). The positive clone was cultured overnight in 450 ml LB medium containing ampicillin. Plasmid DNA was isolated using QIAGEN plasmid maxi kit (QIAGEN), diluted in ultrapure water and stored at −20°C for later use.

Production of viral supernatant and transduction of cells

HEK293T cells were thawed, split every other day for one week and grown in log phase. The day before transfection, 3.5 × 10 6 cells were seeded into tissue culture dishes (60.1 cm 2 growth surface, TPP). The day after, the cells use to reach about 80% confluence. If over confluent, transfection efficiency decreases. The following plasmids were mixed in a total volume of 450 μl ultrapure water: codon-optimized alpharetroviral gag/pol (2.5 μg), VSVG envelope (1.5 μg), and the alpharetroviral vector containing the tdTomato gene (5 μg). Transfection was performed using calcium phosphate transfection kit (Sigma-Aldrich). 50 μl of 2.5 M CaCl 2 was added to the plasmid DNA and the mixture was briefly vortexed. Then, 0.5 ml of 2X HEPES buffered saline (provided in the kit) was added to a 15 ml conical tube and the calcium-DNA mixture was added dropwise via air bubbling and incubated for 20 minutes at room temperature. The medium of the HEK293T cells was first replaced with 8 ml fresh medium (DMEM containing FCS and supplement as mentioned above) containing 25 μM chloroquine. Consecutively the transfection mixture was added. Plates were gently swirled and incubated at 37°C. After 12 hours, the medium was replaced with 6 ml of fresh RPMI containing 10% FCS and supplements. Virus was harvested 36 hours after transfection, passed through a Millex-GP filter with 0.22 μm pore size (Millipore), and used freshly to transduce C1498 cells. Before transduction, 24 well plates were coated with retronectin (Takara, 280 μl/well) for 2 hours at room temperature. Then, retronectin was removed and frozen for later use (it can be re-used at least five times) and 300 μl of PBS containing 2.5% bovine serum albumin (BSA) was added to the wells for 30 minutes at room temperature. To transduce C1498 cells, 5 × 10 4 of cells were spun down and resuspended with 1 ml of fresh virus supernatant containing 4 μg/ml protamine sulfate. The BSA solution was removed from the prepared plates and plates were washed two times with 0.5 ml PBS. Then cells were added to the wells. Plates were centrifuged at 2000RPM/32°C/90 minutes. Fresh medium was added to the cells the day after.

Flow cytometry and fluorescence microscope

For flow cytometry assessment, cells were resuspended in PBS containing 0.5% BSA and 2 mM EDTA and were acquired by a BD FACSCanto™ (BD Biosciences) flow cytometer. Flow cytometry data were analyzed using FlowJo software (Tree Star). Imaging was performed with an Olympus IX71 fluorescent microscope equipped with a DP71 camera (Olympus). Images were analyzed with AxioVision software (Zeiss). Fluorescent images were superimposed on bright-field images using adobe Photoshop CS4 software (Adobe).

Abbreviations

Polymerase chain reaction

Gene of interest

Open reading frame

Melting temperature

Basic local alignment search tool

Vesicular stomatitis virus G glycoprotein

Luria-Bertani

Dulbecco’s Modified Eagle medium

Roswell Park Memorial Institute

Bovine serum albumin

Ethylenediaminetetraacetic acid

Fluorescence-activated cell sorting

Human embryonic kidney

Phosphate buffered saline

Fetal calf serum

Hydroxyethyl-piperazineethane-sulfonic acid

Ampicillin resistance gene

Posttranscriptional regulatory element

Myeloproliferative sarcoma virus promoter.

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Acknowledgments

The authors would like to thank Jessica Herbst, Abbas Behpajooh, Christian Kardinal and Juwita hübner for their fruitful discussions. We also thank Gang Xu for helping to design the cover page. This work was supported by the Deutsche Forschungsgemeinschaft, the Bundesministerium für Bildung und Forschung, the Deutsche Jose-Carreras Leukämiestiftung (grants SFB-738, IFB-TX CBT_6, DJCLS R 14/10 to M.G.S.) and the Ph.D. program Molecular Medicine of the Hannover Medical School.

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Sayed Shahabuddin Hoseini & Martin G Sauer

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Martin G Sauer

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SSH conceived the study subject, carried out experiments and drafted the initial manuscript. MGS participated in study design and coordination and edited the manuscript. Both authors have read and approved the final manuscript.

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13036_2014_161_moesm1_esm.xlsx.

Additional file 1: Ligation calculator. To calculate the amounts of the vector and insert fragments for a ligation reaction, you need to provide the size of the vector and insert (in base pairs), the molar ration of insert/vector (normally 3 to 5), vector amount (normally 50 to 100 ng), and vector and insert fragment concentrations (ng/μl). The computational basis of this ligation calculator is mentioned in the lower box. (XLSX 50 KB)

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Hoseini, S.S., Sauer, M.G. Molecular cloning using polymerase chain reaction, an educational guide for cellular engineering. J Biol Eng 9 , 2 (2015). https://doi.org/10.1186/1754-1611-9-2

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GreenGate - A Novel, Versatile, and Efficient Cloning System for Plant Transgenesis

Affiliation Centre for Organismal Studies, Heidelberg University, Heidelberg, Baden-Württemberg, Germany

* E-mail: [email protected]

• Athanasios Lampropoulos, 
• Zoran Sutikovic, 
• Christian Wenzl, 
• Ira Maegele, 
• Jan U. Lohmann, 
• Joachim Forner

• Published: December 20, 2013
• https://doi.org/10.1371/journal.pone.0083043
• Reader Comments

Building expression constructs for transgenesis is one of the fundamental day-to-day tasks in modern biology. Traditionally it is based on a multitude of type II restriction endonucleases and T4 DNA ligase. Especially in case of long inserts and applications requiring high-throughput, this approach is limited by the number of available unique restriction sites and the need for designing individual cloning strategies for each project. Several alternative cloning systems have been developed in recent years to overcome these issues, including the type IIS enzyme based Golden Gate technique. Here we introduce our GreenGate system for rapidly assembling plant transformation constructs, which is based on the Golden Gate method. GreenGate cloning is simple and efficient since it uses only one type IIS restriction endonuclease, depends on only six types of insert modules (plant promoter, N-terminal tag, coding sequence, C-terminal tag, plant terminator and plant resistance cassette), but at the same time allows assembling several expression cassettes in one binary destination vector from a collection of pre-cloned building blocks. The system is cheap and reliable and when combined with a library of modules considerably speeds up cloning and transgene stacking for plant transformation.

Citation: Lampropoulos A, Sutikovic Z, Wenzl C, Maegele I, Lohmann JU, Forner J (2013) GreenGate - A Novel, Versatile, and Efficient Cloning System for Plant Transgenesis. PLoS ONE 8(12): e83043. https://doi.org/10.1371/journal.pone.0083043

Editor: Paul Jaak Janssen, Belgian Nuclear Research Centre SCK/CEN, Belgium

Received: August 10, 2013; Accepted: November 8, 2013; Published: December 20, 2013

Copyright: © 2013 Lampropoulos et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License , which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: The work was funded by the Collaborative Research Centre SFB873 of the DFG ( www.dfg.de ) and the ERC (erc.europa.eu) grant 282139 “StemCellAdapt”. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Ever since the first construction of a recombinant plasmid [1] , [2] , genetic engineering and molecular cloning mostly rely on the use of type II restriction endonucleases and DNA ligases. The DNA fragments to be combined are first excised from their precursor molecules via the endonucleases and then in a separate reaction re-assembled by the ligase, usually after spontaneous annealing of complementary single-stranded overhangs created during the endonuclease cut. While this approach is generally successful, there are certain limitations, especially when it comes to the assembly of complex plasmids from multiple elements, since with increasing numbers of DNA fragments the ligation reaction becomes less and less efficient. Thus, in case more than four DNA elements have to be assembled, success-rates drop significantly and additional rounds of cloning may be necessary. However, since most recognition sites used in a digestion-ligation cycle remain in the construct, the corresponding enzymes cannot be used for adding further DNA fragments in subsequent steps. Furthermore, many of these recognition sites occur fairly frequently in a given piece of DNA, making the assembly of long constructs even more difficult because of the lack of unique restriction targets. To overcome these limitations, researchers had to devise complicated cloning strategies to assemble plasmids, which involved many steps and required a large number of diverse restriction enzymes. With the advent of high throughput approaches and widespread use of transgenic models to test gene function in vivo , classical restriction based cloning rapidly became a major limitation and alternative technologies began to emerge.

One of the first ligation independent cloning methods was the univector plasmid fusion system, which is based on Cre/ loxP -mediated recombination. Here, a gene of interest was cloned in the so-called pUNI vector and then transferred into a pHOST vector providing the regulatory sequences for expression [3] , which allowed standardized and quick shuffling between a collection of entry and destination vectors. However, the 34 basepairs (bp) of the loxP sites lead to long cloning scars and the number of elements that can be combined is limited to two.

Another very popular recombination based cloning method is Invitrogen’s Gateway® technology [4] , which exploits the lambda phage’s integration and excision mechanism. Briefly, the sequences to be joined need to be flanked by mutually exclusive variants of the attachment sites B and P or L and R. Two different enzyme mixes catalyze either the recombination between B and P sites (creating L and R sites) or between L and R sites (creating B and P sites). Often coding sequences are cloned into so-called entry vectors flanked by attL1 and attL2 sites and then transferred into destination vectors. These carry the regulatory sequences for expression and attR1 and attR2 sites between which the reading frame is inserted. After the recombination reaction, the insert is flanked by attB1 and attB2 sites of 25 bp each. A further development of this technique [5] – MultiSite Gateway® – allows the combination of up to five different fragments with more variants of the attachment sites. Gateway® cloning has been extensively used in many experimental systems, including plants, and large collections of Gateway® compatible vectors are available [6] . Despite this success, Gateway® cloning suffers from three main disadvantages: Firstly, the recombination sites leave 25 bp of unwanted junk sequence - so-called scars - and their inverted repeat sequence poses a problem for expression, sequencing, and RNA probe generation. Secondly, even in the Multisite flavour, the number of fragments that can be combined is limited and the reaction is fairly inefficient. And thirdly, the enzyme mixes required are rather expensive posing a limitation to many labs.

Another elegant way to avoid restriction enzymes and recombinases is ligation independent cloning (LIC) [7] , [8] . This method makes use of the dual function of T4 DNA polymerase, as both 3′-5′ exonuclease and 5′-3′ DNA polymerase. The latter activity predominates in the presence of dNTPs. If only one kind of dNTP is added, the enzyme will digest the 3′-5′ strands of both ends of the DNA fragment until it reaches the first nucleotide for which dNTP is available. Thus single strand overhangs of defined length and sequence are created on both sides, usually 10–15 bp in length for cloning purposes. When fragments with compatible ends are mixed, they spontaneously anneal. Since mostly PCR products are used for LIC cloning, issues with sequence errors due to PCR or primers remain and in case modules are subcloned prior to the LIC reaction, suitable restriction enzymes are necessary to linearize the plasmids. Another issue are the long cloning scars dictated by 10–15 bp overhangs and severe limitations in combining multiple fragments at once.

A major breakthrough in cloning technology was the recent development of the Golden Gate method [9] . It allows fast and efficient assembly of a large number of building blocks requiring only two or three relatively cheap enzymes. The key components of this system are type IIS restriction endonucleases. These enzymes have specific recognition sites but cut arbitrary sequences at a defined distance from the recognition site. This offers several main advantages for cloning: One single enzyme is sufficient to release fragments from diverse preassembled modules and the recognition sites can be placed such that after ligation they cannot be cut again. This enables a one-way reaction of cutting and ligation in one tube, minimizing the experimental steps during the cloning procedure and increasing the efficiency of ligation even for large numbers of fragments. Since the overhangs created by these enzymes can be chosen freely because they are outside of the recognition site, the number of different fragments that can be combined in one reaction is very high. The overhangs generated by these enzymes are small, usually 4 bp, leaving only very short scars. Building blocks can be sub-cloned and sequenced prior to assembly, allowing labs to set up a library of ready-to-use components without the need to verify the final construct by sequencing.

Based on these Golden Gate mechanisms, we designed a simple and versatile cloning system for the generation of plant transformation vectors, which we named GreenGate. The GreenGate system allows rapid and efficient assembly of six modules typically representing promoter, N-terminal tag, coding sequence (CDS) (i.e. the gene of interest), C-terminal tag, plant terminator and plant resistance cassette into a T-DNA transformation plasmid. Furthermore, it offers the option to stack several of these expression cassettes onto a single T-DNA in subsequent rounds, dramatically reducing the time required to generate multi-construct transgenic plants.

Golden Gate Principle

The Golden Gate cloning method allows the rapid and efficient assembly of constructs from pre-cloned building blocks in a single-tube reaction. The method is based on the use of type IIS restriction endonucleases to release DNA fragments from entry vectors and guide them to their specific position in the target plasmid. Type IIS enzymes bind to a defined recognition site, but cut the DNA strand at a fixed distance outside of the recognition motif regardless of the local sequence. This feature can be exploited for a cloning system in which several components are arranged into a single, pre-defined construct by simple incubation with a suitable destination vector, restriction endonuclease and ligase ( Fig. 1 ).

• PPT PowerPoint slide
• PNG larger image
• TIFF original image

A) Type IIS restriction endonucleases, such as Bsa I, have a distinct, non-palindromic recognition site (red) and asymmetrically cut at a precisely defined distance regardless of the local sequence (green). Bsa I for instance creates a four base 5′-overhang starting from the second nucleotide downstream of the recognition site. B) A Golden Gate style cloning system requires two types of components, a destination vector and entry vectors containing the modules to be assembled. Each vector carries two recognition sites for the type IIS endonuclease (red) flanking the counter-selective marker on the destination vector and the modules on the entry vectors, respectively. Destination and entry vectors confer different markers for bacterial selection. The sequences in purple, blue and green represent the cutting sites. C) The orientation and position of the recognition sites is such that after digestion they remain with the backbone of the entry vectors, but are excised from the destination vector along with the counter-selectable marker ( ccdB ). D) The single stranded overhangs generated by the endonuclease can anneal to complementary sequences and be covalently linked by T4 DNA ligase. During the Golden Gate reaction in the presence of endonuclease and ligase the desired final product, but also the original vectors or a plethora of side-products (one of them shown at the bottom) can be created. However, only the desired final product is resistant to further endonucleolytic cleavage, whereas all other molecules will be cut again and again and thus will disappear from the reaction over time.

https://doi.org/10.1371/journal.pone.0083043.g001

Such a cloning system requires all inserts to be flanked by type IIS recognition sites in an orientation that fragments do not carry the sites after type IIS mediated release. This can be achieved either by cloning the fragments into entry vectors that harbour type IIS sites, or by incorporating them by PCR. Specific overhang sequences for each class of insert not only define the orientation of each fragment, but also the order in which they will be assembled in the final construct: Dependent on the intended position in the construct, each module is flanked by a different overhang at 5′- and 3′-end, while the overhangs of adjacent fragments are complementary. Thus, modules will only anneal to each other and the destination vector in the desired orientation and order, upon which ligase activity will complete the assembly. Importantly, no restriction enzyme recognition sites will be present in the final product after ligation, whereas re-ligated entry vectors retain restriction sites and thus can be cut over and over again. For GreenGate we designed the overhangs to be non-palindromic to avoid tandem ligation of inverted fragments and to differ by at least two out of the four bases to ensure specificity of annealing [10] . In contrast to the entry plasmids, the destination vector carries recognition sites in an orientation such that they are removed from the backbone after type IIS digest and that overhangs compatible to those of the outermost insert modules are exposed. In addition, the destination vector encodes an antibiotic resistance different from the one used for the entry vectors, as well as a negative selection marker on the insert. Thus, a reaction including entry vectors, destination vector, type IIS restriction enzyme and ligase will only produce a single product that is able to support bacterial growth after transformation in the presence of positive and negative selection specific to the destination vector. A limitation of Golden Gate cloning is that DNA fragments used as substrates for the reaction must be devoid of recognition sites for the type IIS restriction endonuclease used. This can be overcome by choosing an enzyme that rarely cuts the genome of the targeted model species and/or PCR based mutation of recognition sites prior to cloning.

In essence, Golden Gate cloning is suitable for assembling arrays of any kind of DNA fragments, including PCR products, but it is especially useful when creating multiple permutations of a finite number of building blocks. Constructing expression cassettes for transgenesis is a prime example for such a case, since a limited number of modules (promoters, coding sequences, tags, vector backbones etc.) need to be assembled in an invariant order. Leveraging combinatorial power and re-use of validated building blocks, Golden Gate cloning allows rapid and cost-efficient generation of complex constructs, which is easily scalable to meet high-throughput demands. To make these advantages accessible for the plant science community, we adapted the Golden Gate methodology and created GreenGate, a system for flexible and efficient assembly of plant transformation vectors.

General GreenGate Design Principles

When designing the GreenGate system we aimed at creating a simple and efficient toolbox, which would allow the assembly of routine plant expression constructs, while at the same time facilitate the creation of more complex T-DNAs. Another important aim was to provide full flexibility in the choice of selectable markers for plant transformation, since this has proven to be one of the major bottlenecks in transgenic analyses.

The first important component for a Golden Gate cloning system is the type IIS restriction endonuclease, since it dictates the recognition site sequence. For GreenGate we chose the well characterized Bsa I ( Eco 31I), which recognizes a GC-rich six bp sequence (GGTCTCN ∧ NNNN) underrepresented in the genome of Arabidopsis thaliana ( A. thaliana ), and is commercially available through a number of suppliers. Consequently, DNA fragments containing a Bsa I site need to be modified by mutagenic PCR before cloning into the entry vector.

The second important choice for GreenGate was the number of different modules that can be assembled into a single construct. Since the final plasmid can only be ligated when DNA fragments representing all modules are present in the reaction, dummy sequences need to be introduced if certain modules, e.g. tags, are not needed in a given construct. Thus, we wanted to limit the number of modules for rarely used functions to reduce junk sequences in our expression cassettes. Having a system in mind that would support day-to-day cloning, the number of modules should reflect the essential molecular functions of a gene. Consequently, in the GreenGate system six modules represent the plant promoter (1), an N-terminal tag (2), the coding sequence of the gene of interest (3), a C-terminal tag (4), the plant terminator (5) and the plant resistance cassette (6) ( Fig. 2 ). However, with the exception of the resistance cassette, the modules could also take on other functions, if desired. In those cases where six modules are insufficient to assemble a construct, multiple functions can be bundled in a single module, e.g. by fusion PCR or Golden Gate style fragment joining before cloning into the corresponding entry vector. In addition, we incorporated the option to stack multiple expression cassettes onto a single T-DNA, allowing the generation of highly complex plasmids with a simple workflow that relies exclusively on Bsa I mediated fragment release.

A) The GreenGate cloning system uses six different types of pUC19 based entry vectors into which the individual elements are inserted and a pGreen-IIS based destination vector. Magenta scissors represent Bsa I recognition sites. In each GreenGate reaction, six modules are ligated between the left border (LB) and the right border (RB) sequences of the destination vector yielding a ready-to-use plant transformation vector with expression unit and resistance cassette. These six modules encompass a plant promoter, an N-terminal tag, a coding sequence (i.e. the gene of interest), a C-terminal tag, a plant terminator and a plant resistance cassette for selection of transgenic plants. The modules can only be ligated in the pre-defined order. B) The orderly assembly is enabled by a set of seven different overhangs. Each module is flanked at its 5′-end by the same overhang as the 3′-end of its preceding neighbor. The individual overhangs all differ from each other by at least two out of the four nucleotides. The underlined nucleotides define coding triplets to which all other coding elements have to be in frame. C) Empty entry vector. The multiple cloning site of pUC19 has been replaced by two Bsa I recognition sites (magenta scissors), the respective overhangs for each module type and a counter-selectable ccdB gene. DNA fragments can be cloned via the specific overhangs, via the Bam HI and Kpn I sites or via A-overhangs after Xcm I digestion. Plac  =  lac promoter, SP6 = SP6 promoter, caR  =  chloramphenicol acetyltransferase gene, T7 = T7 promoter, lacZ  =  lacZα coding sequence, ampR  =  beta-lactamase gene, ori  = origin of replication. D) Empty destination vector. A counter-selectable ccdB -cassette has been inserted between the LB and RB sequences of pGreen-IIS, flanked by Bsa I sites, with overhangs A and G. promoter  = bacterial promoter. The pSa origin of replication ( ori A. tum. ) requires the presence of the helper plasmid pSOUP in agrobacteria.

https://doi.org/10.1371/journal.pone.0083043.g002

GreenGate Entry Vectors

We first established the six basal entry vectors into which DNA fragments need to be cloned to create the GreenGate modules ( Fig. 2 , A to C). As basis for the entry vectors, we chose pUC19, because it is a small high-copy plasmid that allows for blue-and-white selection [11] which is maintained in our design. pUC19 has been successfully used in molecular cloning for decades and confers resistance to ampicillin, which is compatible with the most commonly used Escherichia coli ( E. coli ) strains. To create the entry vectors, we replaced the multiple cloning site (MCS) by two Bsa I recognition sites and the respective module-specific overhangs, flanking a chloramphenicol acetyltransferase ( caR ) -ccdB cassette. This cassette allows counter-selection against the original vector in ccdB sensitive E. coli strains during the initial cloning of the DNA fragment of interest [12] . In addition, blue-and-white selection is possible, since the Shine-Dalgarno sequence and the start codon for lacZα are removed along with the ccdB cassette if the GreenGate reaction is successful. To create ready-to-use GreenGate modules, DNA fragments can be inserted into our entry vectors in three ways: 1) Addition of Bsa I sites and respective overhangs by PCR and cloning using these sites. 2) Fragments can be cloned via Bam HI and Kpn I sites present in the vectors. 3) A-tailed DNA fragments can be ligated after Xcm I-digest of the plasmid. Once the desired piece of DNA has been cloned into any of the entry vectors, it can be sequenced from both ends using standard T7 and SP6 primers. These promoters also allow for direct RNA probe generation using the corresponding polymerases. Since our vectors do not contain inverted repeat structures, sequencing as well as probe generation can be carried out with high efficiency. The entry vectors serve as a universal hub for the DNA fragments assigned to the diverse functions in the final construct, such as promoter, tag, or coding sequence. This identity is encoded in the specific overhangs, which are exposed after Bsa I digest. We designed the overhangs of the individual modules according to the following criteria: 1) All overhangs are non-palindromic to prevent inverted tandem ligation of the same insert. 2) At least two out of four nucleotides differ between overhangs to avoid mis-ligation. 3) We optimized the sequence of the overhangs for functionality, if applicable: The B overhang between promoter and N-tag was designed to be the plant Kozak consensus sequence (AACA) [13] ; C and D overhangs between N-tag and CDS and CDS and C-tag, respectively, encode for glycine or serine codons, commonly used in linker sequences. Furthermore, we designed all N- and C-terminal tags to carry start and stop codons, respectively, and consequently have removed stop codons from all CDS modules to make them fully compatible with our tags. The start codon of the CDS module, however, needs to be retained, since not all B-modules carry an ATG codon. For cases where no tags are desired, we designed dummy inserts for modules B and D (B-dummy and D-dummy), which can replace the tags by short sequences. Since in our design the CDS (module C) is devoid of a stop codon, the D-dummy carries a stop codon to provide this function.

Destination Vectors

For the destination vectors, we chose the plant transformation vector pGreen-IIS as backbone [14] [15] . This vector worked well in our hands both for A. thaliana floral dip transformation and Nicotiana benthamiana (N. benthamiana ) leaf infiltration. A major advantage of pGreen-IIS is its reduced backbone sequence, which only contains the spectinomycin resistance and origins of replication for E. coli and Agrobacterium tumefaciens ( A. tumefaciens ). The replicase required for growth in A. tumefaciens needs to be provided in trans , e.g. on the pSOUP plasmid [14] . The small backbone allows for a larger T-DNA which will be transferred into the plant genome. To create a GreenGate compatible destination vector, we replaced the sequence between the left border (LB) and the right border (RB) of the T-DNA by a caR - ccdB - lacZα cassette flanked by two Bsa I recognition sites in opposite orientation and A and G overhangs ( Fig. 2D ). We made two versions of the destination vector, one where the plant resistance cassette is located at the RB of the T-DNA (pGGZ001) and one where it is located at the LB (pGGZ003). With such a design both ccdB counterselection in a ccdB sensitive strain and blue-and-white screening are possible to discriminate against the original vector after the GreenGate reaction, which reduces the background of undesired colonies. All available GreenGate vectors are listed in Table 1 .

https://doi.org/10.1371/journal.pone.0083043.t001

Plant Resistance Cassettes

One of the main issues we wanted to address with the GreenGate system was to facilitate transgene stacking. Since most plant transformation vectors confer resistance to either Basta™ or kanamycin, selecting transgenic events in secondary or tertiary transformations (e.g. into lines from the Salk [16] or SAIL [17] T-DNA insertion mutant collections) can be challenging. To overcome these limitations we designed a collection of different plant resistance cassettes as GreenGate modules, which can now be included into the assembly of any transformation vector. We chose five selectable markers that work well in our hands, namely phosphinotricine acetyltransferase ( BastaR ) conferring resistance against the herbicide Basta™, neomycin phosphotransferase II ( kanR ) against kanamycin, hygromycinB phosphotransferase ( hygR ) to hygromycin, dihydropteroate synthase ( SulfR ) allowing growth on sulfadiazine [18] , [19] and D-amino acid oxidase ( D-alaR ) detoxifying D-alanine [20] . Since multiple copies of identical sequences in transgenes can trigger gene silencing [21] , we designed the cassettes in a way to maximize regulatory sequence divergence for these markers. In addition to the nopaline synthase ( NOS ) promoter and terminator, we used the mannopine synthase ( MAS ) and cauliflower mosaic virus 35S ( 35S ) promoters and terminators, the UBIQUITIN10 ( UBQ10 ) promoter and the octopine synthase ( OCS ) terminator in different combinations. We also removed the chi sequence [22] that potentially enhances recombination in bacteria from the phosphinotricine acetyltransferase coding sequence in one of the cassettes. An overview over available resistance cassettes is given in Table 1 .

To test if all eight resistance cassettes are fully functional when selecting T1 plants, we created a construct harboring the promoter ( p ) of APETALA3 ( AP3 ) fused to mCherry, which was connected to the WUSCHEL ( WUS ) cDNA by a linker coding for 34 amino acids followed by the same linker and GREEN FLUORESCENT PROTEIN ( GFP ). As transcriptional terminator ( t ) we used 650 bp of the 3′-downstream region of the RIBULOSE-1,5-BISPHOSPHATE CARBOXYLASE OXYGENASE SMALL CHAIN ( RBCS ) gene. This pAP3:mCherry-linker-WUS-linker-GFP:tRBCS expression cassette was combined with all eight resistance modules. We recovered resistant seedlings for all of the modules and nearly all transgenic plants displayed the expected phenotype (see below). The availability of a wide range of selectable markers for plant transgenesis will support simple generation of double and triple transgenic plants.

Proof of Principle

After establishing all basic components for GreenGate, we tested them in a number of applications. To this end, we generated functional modules for all classes and assembled them into plant transformation constructs via GreenGate reactions. The pTL003 plasmid coded for an mCherry-linker-WUSCHEL-linker-GFP fusion driven by the flower specific AP3 promoter and flanked on the 3′-end by the RBCS terminator [23] . We chose the resistance to the popular herbicide Basta™ as plant selectable marker, which was flanked by MAS regulatory sequences. For pTL004 we used the same coding modules combined with the WUS promoter and terminator [24] , as well as the pUBQ10:hygR:tOCS resistance module. pTL005 was assembled using the 35S promoter, mCherry-linker [25] , the WUS ORF, the D-dummy module and the RBCS terminator combined with the pMAS:D-alaR:tMAS resistance module. We obtained more than a hundred colonies after transformation into chemically competent E. coli for each GreenGate reaction. We randomly picked eight colonies for each construct and checked for the presence of the desired insert by colony PCR. Since all colonies tested were positive, we isolated plasmid DNA from four clones each and analyzed it by several test digestions. In all but one instance digestion patterns matched the expected sequence, which was finally confirmed by Sanger sequencing of a single clone for each construct.

All three GreenGate constructs were used for A. thaliana transformation. From 500 µL of T1 seeds sown out for each construct on selective medium, we recovered 62 transgenic seedlings for pTL003, eight for pTL004 and none for pTL005. 61 of the 62 pTL003 plants exhibited the strong pAP3:WUS phenotype ( Fig. 3 , D and E). Instead of 4 petals and 6 stamens, their flowers showed a large number of carpelloid organs and were mostly infertile [26] . As expected from the addition of the N-terminal mCherry and the C-terminal GFP tags, pTL003 positive plants also showed red and green fluorescence in the nuclei of cells in whorls 2 and 3 in floral buds ( Fig. 3 , C and F). We were able to recover a small number of seeds from two independent T1 plants and could confirm the AP3:WUS floral phenotype in T2. Transgenic plants recovered from the transformation with pTL004 did not show a phenotype, as expected, since here the endogenous WUS promoter drives a fluorescently labelled version of WUS. Confocal microscopy revealed that all plants showed mCherry and GFP activity in nuclei of the organizing centre, in line with the expected WUS expression pattern (not shown). The low number of T1 transgenics recovered in our experiment is consistent with our experience with transgenes containing the WUS promoter in other vector systems. Similarly, we were not surprised to be unable to recover transgenic plants after transformation with pTL005, since ubiquitous expression of WUS is lethal. However, when we infiltrated N. benthamiana leaves with agrobacteria carrying this construct, we detected mCherry activity in a large number of nuclei (not shown), showing that the expression cassette on the T-DNA is functional.

pTL003 (a pAP3:mCherry-linker-WUS-linker-GFP:tRBCS ; pMAS:BastaR:tMAS construct) served as positive control for the GreenGate concept. When compared to wild-type (A and B), 61 of the 62 T1 transformants displayed a severe pAP3:WUS -flower phenotype (D and E), where instead of four petals and six stamen a large number of carpelloid floral organs is formed. Confocal microscopy of inflorescence apices revealed GFP (C) and mCherry (F) expression in whorls two and three of floral buds. AP3  =  APETALA3 , WUS  =  WUSCHEL , RBCS  =  RuBisCO small subunit . Two identical constructs were created with either GreenGate or a 2-component Gateway®-based cloning method. pTL013 ( pUBQ10:B-dummy-GFP-NLS-D-dummy:tRBCS ; pMAS:BastaR:tMAS ) for GreenGate, pJF343 ( pUBQ10:attB1-GFP-NLS-attB2:tRBCS ; pNOS:BastaR:tNOS ) in case of Gateway®. UBQ10  =  UBIQUITIN10 , NLS  =  nuclear localization signal . Pictures of Nicotiana benthamiana leaves were taken three days after infiltration with pTL013 (G) and pJF343 (H); the signal from leaves infiltrated with the GreenGate derived construct is visibly brighter. Quantification of the fluorescence intensity of single nuclei from both approaches was done by confocal laser scanning (I) and epifluorescence microscopy (J) in two independent experiments. The signal from the GreenGate derived construct is stronger, and intensity ranges between both approaches hardly overlap.

https://doi.org/10.1371/journal.pone.0083043.g003

Furthermore, we created two test constructs to compare the expression strength between a GreenGate-derived plasmid (pTL013) and one generated by Gateway®-cloning (pJF343). Both constructs contain the same elements and use the UBQ10 promoter to drive expression of nuclear localized GFP. Thus, the main difference between the two transcriptional units are the attB1 and attB2 sites flanking the coding sequence in the Gateway® assembled plasmids. When N. benthamiana leaves were infiltrated with these constructs, the fluorescence from the GreenGate version was visibly brighter when compared to the Gateway® derived clones ( Fig. 3 , G and H) and quantification of fluorescence intensities supported this impression ( Fig. 3 , I and J).

Since in many experiments N- and C-terminal tags are not necessary, we also compared the influence of our dummy sequences, the B-dummy (pGGB002), the Ω-element (pGGB003), a translational enhancer from tobacco mosaic virus (U1) [27] , and the D-dummy (pGGD002), by using the pAP3:WUS phenotypes as a quantitative readout. In constructs analogous to pTL003, the N-terminal mCherry-linker tag was replaced by either the B-dummy (pZS165) or the Ω-element (pZS163). When scoring for the strength of the pAP3:WUS phenotype, we found 106 plants out of 134 ( = 79%) showing the strong phenotype for pZS165 and 54 out of 98 ( = 55%) for pZS163, suggesting that the inclusion of the Ω-element did not enhance protein expression in this scenario. When we exchanged the C-terminal linker-GFP tag for the D-dummy to create pZS164, nearly all T1 plants (78 out of 79) showed the strong phenotype as was the case for pTL003, which carried the doubly tagged form of WUS.

Two Expression Cassettes on One T-DNA

To speed up transgene stacking we devised a system to place multiple expression cassettes on a single T-DNA. We aimed to achieve this using our pre-cloned modules without the need to set up a parallel system with different overhangs or recognition sites for a second type IIS enzyme.

In a first attempt we limited ourselves to two expression cassettes on one T-DNA plasmid ( Fig. 4A ). To this end, we created two intermediate vectors, pGGM000 and pGGN000, and two adapter molecules. The final construct is created in two steps: First, the two transcriptional units are assembled separately in two parallel reactions into the intermediate vectors. The first of these two supermodules consists of a plant promoter, an N-terminal tag, a coding sequence, a C-terminal tag and a plant terminator. Furthermore, a pre-cloned adapter module with F and H overhangs is added to the reaction. The intermediate vector has Bsa I recognition sites remaining in the vector backbone with matching A and H overhangs and confers resistance to kanamycin. The elements are combined in a GreenGate reaction but because two Bsa I sites remain in the final construct, an additional ligation reaction is performed before transformation. The second supermodule is built similarly, but the adapter molecule has H and A overhangs and at the 3′-end a plant resistance cassette is added. Thus, the resulting plasmid will have H and G overhangs. The two supermodules are then combined with the destination vector in a second step in a standard GreenGate reaction yielding the final construct.

A) The first strategy uses one additional overhang (“H” = TAGG), two adapter modules and two intermediate vectors. In a first step, two expression cassettes (“supermodules”) are assembled in parallel in two different intermediate vectors (pGGM000 and pGGN000). The Bsa I sites in the intermediate vectors are retained in the supermodule. In the second step, these two supermodules are then transferred into a destination vector via a normal GreenGate reaction. The overhang types are given in capital letters. p1/2 = promoter, n1/2 = N-terminal tag, cds1/2 = coding sequence, c1/2 = C-terminal tag, t1/2 = terminator, r1 = plant resistance, ad.1 = FH-adapter module, ad.2 = HA-adapter module. B) Fluorescence microscopy images show Nicotiana benthamiana leaves infiltrated with a construct harboring two expression cassettes on one T-DNA created via this method. The images were taken 72 hours after infiltration and 24 hours after ethanol induction (picture on the right). The first transcriptional unit drives constitutive expression of the ALCR transcription factor ( pUBQ10:B-dummy-ALCR-D-dummy:tRBCS ; pMAS:sulfR:t35S ), the second one ( pALCA:Ω-element-GFP-NLS-D-dummy:tRBCS ) of nuclear localized GFP in presence of ethanol-bound ALCR protein. C) Only one additional element is required for the second strategy. Instead of a plant resistance cassette module, the FH-adapter module from strategy #1 and an oligo duplex (orange) with unpaired H and G overhangs are used in the GreenGate reaction. The oligo duplex contains internal Bsa I sites that would result in A and G overhangs after digestion. However, digestion is blocked by methylation of the cytosine residues in the Bsa I recognition sites, since Bsa I is sensitive to methylation. After transformation of the resulting construct into bacteria, the methylation is lost during replication because no dcm site is present. Thus, after re-isolation from bacteria, the plasmid, already containing one expression cassette, can function as an empty GreenGate destination vector, releasing A and G overhangs after digestion by Bsa I and removal of the Bsa I recognition sites from the vector backbone. This process can in principle be re-iterated infinitely. The construct is finalized by using a standard plant resistance module in the last step. D) N. benthamiana leaves infiltrated with a destination vector (pTL019) carrying three transcriptional units assembled by this method. The fluorescence signal from all three individual expression cassettes, i.e. nuclear localized BFP (left), ER-localized GFP (second from left) and nuclear localized mCherry (third from left), is visible in all transformed cells. Merge shown on the right.

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As a proof of concept we made an ethanol-inducible GFP-nuclear localization signal ( NLS ) construct, using the ALCA promoter and the ALCR transcription factor from the Aspergillus nidulans alc regulon [28] and the Ω-element . First supermodules pGGM002 ( pALCA:Ω-element-GFP-NLS-D-dummy:tRBCS ; FH-adapter) and pGGN002 (HG-adapter; pUBQ10:B-dummy-ALCR-D-dummy:tRBCS ; pMAS:sulfR:t35S ) were created. These two were combined with the destination vector pGGZ001 to give rise to pTL016 ( pALCA:Ω-element-GFP-NLS-D-dummy:tRBCS ; FH-adapter; HG-adapter; pUBQ10:B-dummy-ALCR-D-dummy:tRBCS ; pMAS:sulfR:t35S ).

Cloning the two intermediate supermodules was not as efficient as the standard GreenGate reactions. We observed a reduced absolute number of transformants and among these in one case more than 50% had aberrant plasmid digestion patterns. Combining the two supermodules with the destination vector, however, was again extremely efficient.

N. benthamiana leaves from four plants were infiltrated with agrobacteria carrying this construct. 48 hours after infiltration half of the plants were watered with 1% ethanol to induce GFP expression, the other half served as control. GFP fluorescence was clearly visible 24 hours later in the nuclei of the induced leaves, but rarely in the control. One control plant was then watered with 1% ethanol and checked for fluorescence again 24 hours later. This time the nuclei were brightly green, showing that the double expression construct works as designed ( Fig. 4B ).

More than Two Expression Cassettes on One T-DNA

To combine even more transcriptional units on one plasmid the way described for the double expression constructs, the system would have required a substantial number of additional overhangs and supermodule vectors, while at the same time limiting ourselves to a fixed maximum of expression cassettes. To avoid these complications and to retain full flexibility with regard to transgene complexity, we came up with a much simpler procedure, which required only a single additional element ( Fig. 4C ). This module is a synthetic DNA duplex with unpaired ends that allows ligation to H and G overhangs during a GreenGate reaction. Furthermore, it contains two internal Bsa I recognition sites, which cannot be cut by Bsa I due to overlapping cytosine methylation. Thus, the element will behave as an ordinary H to G module in the first GreenGate reaction and replace the resistance cassette. However, after isolation of the resulting plasmid from bacteria the methylation will be lost because no dcm sites are present in the oligo duplex. Consequently the two internal Bsa I recognition sites are now targets for Bsa I in a second round of GreenGate, giving rise to A and G overhangs, simulating an empty destination vector. Thus, in each subsequent GreenGate round either the uncleavable oligo duplex can be used to prepare the plasmid for the insertion of another expression cassette or the plasmid can be finalized by adding a plant resistance cassette to the reaction. In this approach, the transcriptional units are assembled serially, so it requires one additional step for each expression cassette to be added. The FH-adapter used along with the oligo duplex separates the individual expression units in the final construct and thus can be freely designed to act as a spacer of arbitrary length and sequence minimizing the risk of mutual interference of the expression behavior.

We tested this approach with a triple expression construct, namely pTL019 ( p35S:ER signal sequence-GFP-ER retrieval signal:tRBCS ; FH-adapter; pRPS5A:SV40 NLS-E. coli biotin ligase-linker-BFP:TAt1g04880 ; FH-adapter; pUBQ10:B-dummy-mCherry-linker-NLS:tAt1g76110 ; pMAS:BastaR:tMAS in pGGZ003). The expression cassettes were assembled in the given order using the serial strategy described above. Similar to our experience with using supermodules, cloning was not as efficient as for single unit plasmids. Again, we observed a severely reduced number of transformants, and many clones gave rise to an aberrant digestion pattern. In steps two and three we also observed a high percentage of the original destination vector, even after including an additional Bsa I digestion step after the GreenGate reaction. Once we had identified a correct clone, the final construct was used for infiltration of N. benthamiana leaves. Fluorescence was analyzed 64 hours after infiltration ( Fig. 4D ) and all three fluorescent proteins were clearly detectable in the subcellular compartments predicted by their primary sequence. Thus with these two approaches it is possible to create ready-to-use constructs containing multiple expression cassettes within 2–3 weeks.

We established GreenGate, a novel cloning system for plant transgenesis that supports easy, quick, modular and reliable assembly of plant expression cassettes directly in plant transformation vectors. Several independent and complex constructs have been successfully created and were functional in stable transformation of A. thaliana and transient essays in N. benthamiana leaves, respectively.

GreenGate is designed to match the requirements of routine and advanced cloning for plant transgenesis and, therefore, we adapted the Golden Gate [10] layout to encompass the six most frequently used elements in plant expression cassettes, namely plant promoters, N-terminal tags, coding sequences of the gene of interest, C-terminal tags, plant terminators and resistance cassettes for selection of transgenic plants. Furthermore, unlike in the MoClo [29] or GoldenBraid systems [30] , where two or three different enzymes are used, GreenGate makes use of only one type IIS restriction endonuclease minimizing the likelihood of naturally occurring restriction sites in inserts, which would interfere with the reaction and thus need to be removed prior to cloning. Nevertheless, like the MoClo [29] or GoldenBraid systems [30] , GreenGate allows stacking of several transcription units onto one construct and generation of libraries of re-usable higher-order supermodules. Since agrobacteria in our hands only tolerate pGreen-IIs based constructs of up to 20 kb, the number of possible expression cassettes on a single construct is, however, limited irrespective of the cloning system.

When comparing GreenGate to the most frequently used cloning systems for plant transgenesis so far, namely classical cloning and 2-component Gateway®, the main advantages are the full modularity, including the plant selectable marker, the fact that unwanted “scar” sequences are kept to a minimum and that entry clones can serve as templates for high quality sequencing and probe generation. While classical cloning allows sequencing and RNA probe generation, individual cloning strategies have to be worked out for every construct and the limited availability of unique restriction sites may severely reduce options for assembly of more complex transgenes. In contrast, 2-component Gateway®, is independent of restriction site limitations when it comes to cloning an insert into a given destination vector, but still promoter, terminator and resistance cassette need to be cloned traditionally during the design of this element, severely limiting the modularity of the system. In addition, long inverted-repeat recombination sites hamper sequencing and probe generation from entry clones and since they remain in the final construct also affect transgene function. This notion was supported by our finding that GreenGate assembled constructs not only lead to higher fluorescence intensities in transiently expressing leaf mesophyll cells of N. benthamiana ( Fig. 3 , G–J), but also produced a higher frequency of transgenic lines with strong AP3:WUS phenotypes in A. thaliana . Similar to Gateway®, ligation independent cloning leaves large scars in the constructs and either requires restriction enzyme sites to be removed when inserts are pre-cloned or to rely on PCR products without prior sequencing. Both systems in addition do not offer full modularity for assembly of constructs and thus require a large number of destination vectors to be designed and produced. In contrast, the MultiSite Gateway® cloning system allows five elements to be combined, but is rather expensive, in our hands rather inefficient, and still suffers from the cloning scars discussed above. GreenGate overcomes most of these limitations and thus should greatly facilitate the assembly of plant transformation constructs.

While being able to quickly assemble a large number of constructs expands our experimental scope, functional studies in plants often require two or more expression units to be tested together, for example regulator-reporter combinations, making plant transgenesis the next bottleneck. Co-transformation is a way to avoid time-consuming crossing of plants transformed with individual constructs, however, this requires the use of multiple plant resistance cassettes and co-transformation efficiency only lies in the range of 20–30% [31] . Combination of several transgenes in general can cause problems with gene silencing caused by multiple copies of identical sequences, for example when resistances are driven by the same promoter [21] . Thus, minimizing the number of resistance cassettes is an important aspect of experimental design. Therefore, we expanded our system to enable cloning of multiple transcriptional units on the same T-DNA from our established module library. The advantages in the downstream analysis of the transgenic plants in our opinion vastly outweighs the disadvantages, which lie in the substantially reduced efficiency of assembling supermodules or working with the methylated synthetic oligo duplex element. On the upside, once the supermodules are generated, they can be re-used just like ordinary modules at the same high efficiency.

After having routinely used GreenGate in the laboratory for more than eight months, we find it working well as everyday technique. However, especially when several people share components, care has to be taken to ensure a certain minimal standard for all elements. We have identified plasmid DNA quality and quantity to be an important determinant of GreenGate efficiency with a DNA concentration of at least 100 ng/µl after column preparation being a minimum requirement. Even more importantly, the activity of the restriction endonuclease is a limiting factor for GreenGate cloning. We found both Bsa I-HF from NewEngland Biolabs, as well as the isoschizomer FastDigest Eco 31I from Fermentas to be highly sensitive to temperature fluctuations in a batch dependent manner. Thus, we highly recommend preparing small aliquots of all components, including the enzymes to avoid repeated freeze thaw cycles to ensure efficient and reliable cloning using GreenGate.

Materials and Methods

Construction of empty entry plasmids.

pUC19 [11] was chosen as basis for the entry constructs. In the first step, the Bsa I site in the beta lactamase ( ampR ) coding sequence was removed and converted into an Xho I site by two silent mutations via PCR. Apart from that no changes were made to the vector backbone. In the same step, the N-terminal part of the lacZα open reading frame around the multiple cloning site was replaced by Hind III and Eco RI sites flanked by SP6 and T7 promoter sequences. The lacZα Shine-Dalgarno sequence and translation start codon were deleted, the rest of the lacZα expression cassette was left unchanged. Next, the Bsa I recognition sites in opposite orientation with the respective overhangs for the six basal entry vectors were inserted, separated by Bam HI and Kpn I sites. In the third step, the Xcm I sites, the Shine-Dalgarno sequence for lacZα and Xba I and Pst I sites were added. The vectors were finalized integrating the open reading frames for chloramphenicol acetyltransferase ( caR ) and ccdB . The internal Bsa I site in ccdB was destroyed by a silent nucleotide exchange.

Vector elements were either generated via PCR amplification or via artificially synthesized oligo duplices; all constructs were verified by sequencing and test digestion. PCRs were done with Phusion High-Fidelity DNA Polymerase (Fisher Scientific - Germany GmbH, Schwerte, Germany). PCR products and intermediate plasmids were cut by conventional restriction endonucleases (obtained from Fisher Scientific - Germany GmbH, Schwerte, Germany) and ligated with T4 DNA ligase (Fisher Scientific - Germany GmbH, Schwerte, Germany). For complex PCR products, the single elements were amplified separately. Sometimes overlap extension PCR was used to combine them to larger constructs, but normally they were assembled by ligation assisted PCR. Classical restriction enzyme sites were added to the primers, the single PCR products were digested with the respective enzymes, ligated and the reaction products used as templates for subsequent PCRs.

For the constructs expressing the ccdB gene, E. coli strain DB3.1 was used, Mach1™-T1 R and SURE otherwise.

pGGE000 was converted into pGGH000 by cutting out the Bsa I recognition site and the E overhang via Hind III and Bam HI digestions and replacing it by an oligonucleotide pair providing the Bsa I site and the B overhang. pGGI000 was constructed similarly from pGGD000.

Construction of Empty Destination Vectors

To create the GreenGate destination vectors, pGreen-IIS [14] was used as PCR template. The vector backbone was left unaltered, but the sequence between left border (LB) and right border (RB) was replaced by a short multiple cloning site. For the final constructs with the plant resistance cassette at the RB, the MCS had the orientation LB- Xho I… Bam HI-RB, and the orientation LB- Bam HI… Xho I-RB for the version with the plant resistance cassette at the LB. In the next step, the Not I and Pae I sites as well as the short spacer sequences in front of the borders were introduced via an oligo duplex (destroying the Bam HI/ Xho I sites), again for both versions of the destination vector. The rest of the multiple cloning site was added in the third step, again using oligo duplices. Finally, the caR-ccdB-lacZα cassette flanked by two Bsa I sites in opposite orientation with the A and G overhangs was created via PCR and ligated into the Kpn I and Xba I sites of the pre-vector. For lack of transformants, only the final destination vector for the plant resistance cassette at the RB (pGGZ001) could be created that way, the one for the plant resistance cassette at the LB (pGGZ002) required an additional step. An intermediate Bsa I- Xho I- Eco RI- Bsa I oligo duplex was cloned into the Kpn I and Xba I sites of the prevectors and the PCR product then added via Xho I and Eco RI sites. Due to the observed instability of pGGZ002 in bacteria, we created pGGZ003 by PCR amplifying the ccdB cassette with exchanged Xho I and Eco RI sites thus inverting the ccdB cassette orientation but still retaining the plant resistance cassette at the LB design.

Bacterial host strains were DH5α (first vector), Mach1™-T1 R (all other intermediate vectors) and DB3.1 (final vectors).

To change the bacterial resistance in the destination vector from spectinomycin to gentamicin, the respective plasmids (pGGZ001, pGGZ002, pGGZ003) were amplified from the nucleotide directly downstream of the spectinomycin adenyltransferase ( specR ) gene stop codon to the nucleotide directly upstream of the start codon. External Bsa I sites were added to the primers. The gentamicin acetyltransferase ( gentR ) reading frame from A. tumefaciens strain GV3101 (pMP90RK) was amplified also with external Bsa I sites and compatible overhangs. Because this did not yield any transformants, a second PCR product was created additionally containing 69 nucleotides of the gentR 3′-UTR. E. coli cells transformed with the resulting plasmids (pGGY001– pGGY003) were grown in medium with 5 µg/mL gentamicin, for A. tumefaciens the respective concentrations ranged from 10 to 20 µg/mL.

Ampicillin and spectinomycin were used at 100 µg/mL, kanamycin at 50 µg/mL, chloramphenicol at 25 µg/mL and tetracycline (for pSOUP) at 5 µg/mL.

Construction of the Intermediate Vectors for Two Constructs on One T-DNA

pGGM000 and pGGN000 were created from two PCR products each. pENTR1A was amplified without the attL sites and the ccdB cassette flanked by Eco RI and Xho I sites, the primers also providing the Bsa I recognition sites and the suitable overhangs. The caR-ccdB-lacZα cassette was also amplified with the same restriction sites at the end. The PCR products were digested with these enzymes and ligated.

Methylated Oligonucleotide Duplex

The oligonucleotide duplex used in the more than two constructs on one T-DNA approach was created by annealing oligonucleotides A02827 (5′-taggaccttgagacCgaaaaggtggtctCa-3′) and A02828 (5′-atactgagacCaccttttcggtctCaaggt-3′). The nucleotides in capital letters were methylated.

Creation of the GreenGate Entry Modules

In most cases the respective inserts were PCR amplified to create the GreenGate entry modules. The nucleotides 5′-AACA-GGTCTC-A- NNN N (nn)-3′ were added to the forward primer in front of the gene specific sequence. GGTCTC is the Bsa I recognition site, AACA was added because the enzyme does not cut if the restriction site is at the extreme ends of PCR products. NNNN represents the module specific overhang and 2 nucleotides (nn) are needed in case of the coding sequence and C-tag modules to bring the modules into frame ( NNN represents an in-frame coding triplet in the overhangs). The sequence 5′-AACA-GGTCTC-A-N NNN -3′ was added to the reverse primers, followed by the reverse complement of the sequence of interest. N NNN stands for the reverse complement of the module specific overhang, the coding triplet being underlined.

After amplification, the PCR reactions were separated on agarose gels, the product bands excised, purified with innuPREP DOUBLEpure Kit (Analytik Jena AG, Jena, Germany) and digested with Bsa I. The respective empty entry modules (∼ 100 ng) were also cut with this enzyme, usually in the same tube (1 h, 37°C). The digestion was purified with the above mentioned kit and ligated with T4 DNA ligase (1 h room temperature, overnight 4°C). After heat-inactivation (10 min, 70°C) the reaction was transformed via heat shock into ccdB sensitive E. coli strains (Mach1™-T1 R , DH5α, XL1-Blue MR). Transformants were checked by colony PCR, plasmid DNA was isolated from positive clones and checked by sequencing and test digestion.

If internal Bsa I recognition sites were present in the module sequence, they were removed by nucleotide substitution. For protein-coding sequences, silent mutations were chosen. In promoter and terminator sequences, the nucleotides to be changed were selected at random, but for later constructs we switched to always replace the first guanine by a cytosine. For simplicity, we used scar-free Bsa I-cloning to create the substitutions. Primers were designed on both sides of the internal Bsa I recognition sites introducing the mismatch and flanked on their 5′-ends by external Bsa I recognition sites. The overhangs generated by the external Bsa I cut were designed to be part of the gene-specific sequence and being different from the module specific overhangs.

Shorter modules were assembled as oligonucleotide duplices created from overlapping primers with unpaired 5′-overhangs complementary to the module specific overhangs. The oligonucleotides (10 µM or 100 µM) were mixed in equimolar ratios with each other, soused with boiling water, allowed to cool slowly down to room temperature and then ligated into Bsa I digested and purified entry vector.

GreenGate Reaction

Plasmids were isolated from the bacterial hosts using the innuPREP Plasmid Mini Kit (Analytik Jena AG, Jena, Germany) according to the recommendations of the manufacturer. DNA was eluted with 100–125 µL elution buffer. The DNA concentration was usually not determined exactly but after gel electrophoresis estimated to be around 100 ng/µL judging from earlier preparations done with the same kit.

For the GreenGate reaction itself 1.5 µL plasmid of each of the six modules were mixed with 1 µL of the destination vector, 1.5 µL CutSmart Buffer (alternatively: FastDigest buffer), 1.5 µL ATP (10 mM), 1 µL T4 DNA ligase (30 u/µL) and 1 µL Bsa I-HF (alternatively FastDigest Eco 31I) in a total volume of 15 µL. Bsa I-HF and CutSmart buffer were purchased from New England Biolabs GmbH, Frankfurt am Main, Germany, FastDigest Eco 31I, FastDigest buffer, T4 DNA ligase and ATP from Fisher Scientific - Germany GmbH, Schwerte, Germany. Initially, we performed 50 cycles of 37°C for 5 minutes and 16°C for 5 minutes each, followed by 50°C for 5 minutes and 80°C for 5 minutes. Later we reduced the incubation time to 2 minutes each and the cycle number to 30. 6 µL of the reaction were used for heat-shock transformation of ccdB -sensitive E. coli (strains Mach1™-T1 R , DH5α or XL1-Blue MR). 25–100 µL of competent cells (1.0–4.0*10 8 cfu/1 µg pUC19 DNA/100 µL) were used and usually hundreds of transformants were recovered after plating (on two separate plates - 1/15 and the rest).

The products of the first GreenGate reactions were analyzed by both restriction endonuclease test digestions and sequencing of the ligation sites, later on by test digestions only.

When creating the intermediate supermodules for the two constructs on one T-DNA approach, ligase and ATP were added again after the GreenGate reaction and the mixture incubated for one more hour at room temperature before heat-inactivation and heat-shock transformation.

When using the methylated oligonucleotide pair, Bsa I was added after the GreenGate reaction and digestions carried out for one hour at 37°C prior to heat-inactivation and transformation.

Troubleshooting

Usually the GreenGate system works quite reliably. In instances where the total number of colonies and/or the ratio of correct clones dropped, we could mostly attribute it to either bad enzyme quality or low DNA concentration of the modules and destination vectors. Since the activity of the restriction endonucleases suffers substantially from prolonged exposure to room temperature, we recommend aliquoting all enzymes and buffers to avoid repeated freeze-thaw cycles. Plasmid DNA minipreparations should be column purified, plasmid integrity checked by gel electrophoresis and their DNA concentration measured (100 ng/µl minimum). In case of frequently used elements, such as the destination vector or standard tags, we found midi preparations to be preferable and overall more reliable.

In case final constructs are obtained in which modules are ligated in arbitrary order, which suggests that 5′-overhangs were lost during the reaction, we suggest using a fresh enzyme batch. For some enzyme batches we found a reduction of enzyme amount by 50% to be very beneficial. Alternatively, using the long reaction program (50 cycles of 5 min at 37°C and 16°C each) helped to increase efficiency. When restriction enzyme activity is insufficient, plasmids with an integration of a complete entry vector with intact Bsa I sites might be recovered. This can be remedied by incubating with fresh Bsa I after the GreenGate reaction.

Nicotiana Benthamiana Infiltration

A. tumefaciens strain ASE (pSOUP + ) was transformed with the respective constructs. Transgenic clones were cultured in liquid selective LB medium for 2 nights at 28°C, resuspended in infiltration medium (10 mM MgCl 2 , 10 mM MES, 150 µM acetosyringone, pH 5.7) and pressed with a blunt syringe through the stomata at the abaxial site of leaves from approximately four weeks old plants [32] . The plants were put back to the incubators for 48 or 72 hours before analyzing the fluorescence levels.

For ethanol induction, plants were watered with a 1% v/v solution of ethanol.

Confocal laser scanning microscopy was done on a Nikon A1 Confocal Microscope with a 25× apochromatic lens, epifluorescence microscopy at a Zeiss Axio Imager.M1. Fluorescence intensities were measured with the ImageJ software.

Arabidopsis thaliana Transformation and Plant Selection

A. thaliana plants were transformed by a modified version of the floral dip protocol [33] , where only the tips of the inflorescences were dipped into the agrobacteria solution.

For selection on soil, Basta™ (glufosinate-ammonium; Bayer CropScience Deutschland GmbH, Langenfeld, Germany) was used at a concentration of 20 mg/L, both for spraying and watering. The concentrations of sulfadiazine, D-alanine, Basta™, kanamycin and hygromycin for selection on ½ MS plates were 0.75–7.5 µg/mL, 12 mM, 10 µg/mL, 50 µg/ml and 25 µg/mL, respectively.

Plasmid Availability

All plasmids listed in Table 1 are available from Addgene (Massachussetts, USA) at www.addgene.org/cloning/greengate/lohmann . The sequence information has been deposited in GenBank under accession numbers KF718964 - KF719019.

Acknowledgments

The authors would like to thank Stephan Kirchmaier for the introduction into Golden Gate cloning and advice on optimizing the parameters of the GreenGate reaction.

Author Contributions

Conceived and designed the experiments: AL ZS JF IM JL. Performed the experiments: AL ZS CW JF IM. Analyzed the data: AL ZS CW JF JL. Wrote the paper: JF JL.

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Advances in molecular cloning

• Published: 30 March 2016
• Volume 50 , pages 1–6, ( 2016 )

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• Malla Ashwini 1 ,
• Shanmugaraj Bala Murugan 1 , 2 ,
• Srinivasan Balamurugan 1 , 3 &
• Ramalingam Sathishkumar 1  

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“Molecular cloning” meaning creation of recombinant DNA molecules has impelled advancement throughout life sciences. DNA manipulation has become easy due to powerful tools showing exponential growth in applications and sophistication of recombinant DNA technology. Cloning genes has become simple what led to an explosion in the understanding of gene function by seamlessly stitching together multiple DNA fragments or by the use of swappable gene cassettes, maximizing swiftness and litheness. A novel archetype might materialize in the near future with synthetic biology techniques that will facilitate quicker assembly and iteration of DNA clones, accelerating the progress of gene therapy vectors, recombinant protein production processes and new vaccines by in vitro chemical synthesis of any in silico-specified DNA construct. The advent of innovative cloning techniques has opened the door to more refined applications such as identification and mapping of epigenetic modifications and high-throughput assembly of combinatorial libraries. In this review, we will examine the major breakthroughs in cloning techniques and their applications in various areas of biological research that have evolved mainly due to easy construction of novel expression systems.

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Plant Genetic Engineering Laboratory, Department of Biotechnology, Bharathiar University, Coimbatore, Tamil Nadu, India

Malla Ashwini, Shanmugaraj Bala Murugan, Srinivasan Balamurugan & Ramalingam Sathishkumar

Department of Biotechnology, PSR College of Engineering, Sivakasi, Tamil Nadu, India

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Department of Genetic Engineering, SRM University, Chennai, Tamil Nadu, India

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Published in Russian in Molekulyarnaya Biologiya, 2016, Vol. 50, No. 1, pp. 3–9.

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The Biotechnology Revolution: PCR and the Use of Reverse Transcriptase to Clone Expressed Genes

The cloning of expressed genes and the polymerase chain reaction (PCR), two biotechnological breakthroughs of the 1970s and 1980s, continue to play significant roles in science today. Both technologies give researchers the means to make more DNA , but they do so in different ways. In particular, cloning involves the synthesis of DNA from mRNA using an enzyme called reverse transcriptase . Although this method reverses the flow of genetic information as described by the central dogma , it effectively mimics the process by which RNA viruses "flip" the direction of transcription in their host cells, thereby causing these cells to manufacture viral DNA even though the viruses themselves contain only RNA. In contrast, the polymerase chain reaction does not involve the use of an initial mRNA template to manufacture DNA. Rather, PCR involves the synthesis of multiple copies of specific DNA fragments using an enzyme known as DNA polymerase . This method allows for the creation of literally billions of DNA molecules within a matter of hours, making it much more efficient than the cloning of expressed genes. However, cloning remains the go-to method for researchers when only the mRNA template (and not the DNA template) of a sequence of interest is available.

Making DNA from RNA: Reversal of the Central Dogma

--"Central Dogma Reversed," Nature , June 27, 1970

The so-called central dogma of molecular biology states that all genetic information flows in one direction: from DNA to RNA through the process of transcription , and then from RNA to protein through the process of translation (Crick, 1958). For over a decade, the central dogma was thought to be a universal truth--in other words, researchers believed that genetic information always flowed in this order, otherwise it could not be passed along. In 1970, however, the two experiments mentioned in the Nature quote--one conducted by David Baltimore, then of the California Institute of Technology in Pasadena, and the other by Howard Temin and Satoshi Mizutani, then of the University of Wisconsin in Madison--called this belief into question. Specifically, these researchers independently published scientific papers demonstrating that RNA tumor viruses contain enzymes that use viral RNA as a template for the synthesis of DNA, thereby reversing the direction of transcription (Baltimore, 1970; Temin & Mizutani, 1970). Not only did these two experiments challenge the validity of the central dogma, but they also laid the foundation for a series of technological developments that eventually earned reverse transcription and the synthesis of complementary DNA, or cDNA , central places in the molecular biologist's toolbox.

Discovering Reverse Transcription

During the late 1960s, Baltimore, Temin, and Mizutani were each driven by unanswered questions about how RNA viruses transformed healthy cells into tumor cells. They knew that transformation ensued when healthy cells incorporated DNA from the external environment (in this case, RNA tumor virus DNA) into their genomes. But how could a eukaryotic cell incorporate DNA from a virus that didn't have any DNA?

Howard Temin had hypothesized the existence of an enzyme capable of making DNA from RNA as early as 1964 ("Central Dogma Reversed," 1970). But, as is the case with all scientific hypotheses, the research community remained skeptical of this proposal until the 1970 publications wiped that skepticism away. At that point, the race was on to identify the enzyme responsible for the creation of DNA from RNA. Today, that enzyme is known as reverse transcriptase.

Interestingly, in their groundbreaking papers, the two sets of scientists didn't actually identify reverse transcriptase, but they did provide clear and conclusive evidence of the existence of an enzyme that utilized viral RNA as a template for DNA synthesis. The experiments supporting the existence of this DNA polymerase produced data that revealed the following:

• The DNA polymerase only incorporated deoxyribonucleotides, not ribonucleotides, into its product.
• The product itself "behaved" like DNA--in other words, it was sensitive to treatment by deoxyribonucleases but not ribonucleases.
• The RNA itself was the template, as shown by the fact that treatment of virions with ribonucleases destroyed the ability of the polymerase to incorporate radioactively labeled nucleotides.

Although the motivation for both studies was to better understand the role of viruses in some cancers, there is also some suggestion in the papers that the scientists were aware, at least on an intuitive level, that there were far greater implications to their findings. As Temin and Mizutani (1970) wrote, "This result would have strong implications for theories of viral carcinogenesis and, possibly, for theories of information transfer in other biological systems."

It did not take long for scientists to isolate the reverse transcriptase responsible for Baltimore's findings (Verma et al ., 1972). Another team (Bank et al. , 1972) then used the enzyme to synthesize DNA from mRNA in a test tube for the first time. (The so-called complementary DNA that results is referred to as cDNA.) Both teams used globin mRNAs, or mRNAs that encode blood hemoglobin polypeptides, to demonstrate that reverse transcriptase does in fact synthesize DNA from mRNA templates. Moreover, the teams also found that the reaction works best in the abundance of short sequences composed entirely of thymine nucleotides known as oligo(dT) primers. Knowing that most eukaryotic mRNAs have a string of adenine nucleotides--also known as a poly(A) tail--at their 3′ end, the scientists had predicted that cDNA synthesis would require oligo(dT) primers, or that it would at least be made more efficient by the presence of these primers.

How Reverse Transcriptase Works

The first step relies on the fact that most eukaryotic mRNAs have poly(A) tails at their 3′ ends. The poly(A) tails serve as "hooks" during the separation process. This process involves pouring all of the cellular RNA ( tRNA , rRNA , snRNA, mRNA, etc.) into what is known as an elution column of short DNA pieces, mostly thymine nucleotides. As the RNA mixture moves downward through the column, the poly(A) tails of the mRNA molecules bind to the thymine nucleotides. The rest of the RNA molecules--those without poly(A) tails, in other words--run right through. Afterward, the column is washed with a solution that breaks the hydrogen A-T bonds, and the released mRNA molecules are collected.

The second step, the copying of the now-isolated mRNA molecules into cDNA, involves adding oligo(dT) primers to the mRNA collection. These primers pair with the poly(A) tails of the mRNA molecules, again at the 3′ end, providing the exposed 3′-OH group required for the initiation of DNA synthesis . At this point, the reverse transcriptase enzyme is added, and this enzyme proceeds to utilize the mRNA strand as a template for the synthesis of a complementary DNA strand. The enzyme adds nucleotides, one by one, to the 3′ end of the new strand, with each newly added nucleotide a complement to its template pair just as in DNA replication , with the exception that RNA contains Us in place of Ts. Thus, Cs are paired with Gs and vice versa, Ts are paired with As, and As are paired with Us. Scientists then use several different methods to convert the RNA-DNA hybrids into double-stranded cDNA molecules, such as enzymatic digestion of the RNA strand followed by DNA synthesis utilizing, this time, the cDNA strand as the template.

Using Reverse Transcriptase to Clone Expressed Genes

Following the development of this method, the use of reverse transcriptase to clone expressed genes grew for several decades. However, there were limits to this practice. For example, most cDNA molecules that were synthesized in a single reaction were incomplete, with the 5′ end of the mRNA not represented in the final cDNA. Therefore, the cDNA did not contain a complete copy of the amino acid-encoding region of the gene.

Eventually, in the late 1990s, Piero Carninci and his colleagues at the Genome Science Laboratory in Ibaraki, Japan, devised a series of methods to get around this and other problems. In particular, these researchers developed a new technique for selecting full-length cDNA molecules. This process is known as biotin capping, and it involves capping the 5′ end of the mRNA with a biotin group and then washing the cDNAs with an RNA digestion enzyme, like RNAse I. When washed with a solution containing a cap-binding protein, all of the cDNA-mRNA hybrids with only partial cDNA copies are carried away, thus only leaving behind the hybrids with full-length cDNA molecules. In fact, the researchers demonstrated that biotin capping yielded about 95% full-length cDNA clones (Carninci et al. , 1996).

Today, scientists continue to build and utilize what are known as cDNA libraries, or collections of cDNAs from particular tissues gathered at particular times during an organism 's life cycle. The synthesis of cDNA molecules is referred to as cloning, because the cDNA molecules are matching copies of the DNA responsible for encoding the mRNA template.

Scientists often generate cDNA libraries as a way to find genes of interest. They screen these libraries using what are known as probes--complementary pieces of DNA that hybridize to the cDNA molecules. They also use cDNA libraries to identify genes that are expressed differently in different types of tissues or at different developmental stages. Libraries of cDNA molecules provide snapshots of gene activity, because only those genes that are actually expressed and transcribed into mRNA molecules can be cloned. For example, one would expect a cDNA library compiled from mRNA isolated during a stage of prenatal development to be very different from a cDNA library generated from sequences transcribed during adulthood.

Making Copies via Polymerase Chain Reaction

How pcr works.

Basically, PCR is DNA replication on a grander scale . The polymerase chain reaction relies on the use of several essential chemical ingredients, including the following:

• A DNA polymerase
• A small amount of DNA to serve as the initial template
• The four deoxyribonucleotides to serve as the substrates for the DNA polymerase and the raw ingredients of the new DNA molecules
• A few necessary ions and salts
• A pair of primers with exposed 3′-OH groups that will bind to the particular sequence of interest in the DNA template

As previously mentioned, the DNA polymerases can only add new nucleotides to the 3′-OH end of a growing strand. They therefore require the presence of a primer to get started, because they cannot begin synthesis de novo . In fact, two primers are required--one to initiate replication of each of the two DNA strands.

A single PCR reaction involves three temperature-dependent steps, described as follows:

• The starting solution is heated, usually to between 90° and 100°C. The high temperatures break the hydrogen bonds between the two strands of the original DNA double helix , providing the necessary single-stranded templates.
• After just a couple of minutes at that temperature, the reaction mixture is quickly cooled, usually to somewhere between 30° and 65°C. It is then held for less than a minute at this lower temperature--which is enough time for the primers to bind to their complementary sequences on the single-stranded templates.
• The sample is next heated to 60° to 75°C for less than a minute, during which time the DNA polymerase adds nucleotides to the primer, synthesizing a new DNA strand using only the template sequences that bind the primers (Figure 2).

The many changes in temperature required during multiple PCR cycles are carried out in a thermocycler, also known as a PCR machine. After PCR cycling is complete, the amplification products can be subjected to cloning, sequencing, or analysis via gel electrophoresis .

Advances in PCR Technology

As with all genetic technologies, of course, scientists have improved and refined the original PCR process described by Mullis and Faloona in 1987. For example, one of the major limitations of early PCR methods was that fresh DNA polymerase had to be added during every cycle. This repetitive step was not just tedious, but it also greatly increased the likelihood of error. Mullis and colleagues addressed this deficiency just a year later when they demonstrated how a particular type of DNA polymerase, a heat-resistant enzyme isolated from the bacterium Thermus aquaticus , eliminated the need to add fresh polymerase during every cycle. Thermus aquaticus --often referred to by its popular nickname "Taq polymerase" -- is a thermophilic bacterium that can survive temperatures up to 95°C. In fact, its natural habitat is the hot spring ecosystem of Yellowstone National Park. This innovation greatly improved the quantity and quality of PCR products (Saiki et al. , 1988).

More recently, another major PCR innovation was the development of real-time PCR. This refinement involves the use of dyes or fluorescent probes that eliminate the need for post-PCR electrophoresis . In real-time PCR, the fluorescence that is associated with the accumulation of newly amplified DNA is measured through the use of an optical sensing system.

Thus, both cloning of expressed genes and PCR continue to serve as essential tools for genetic researchers. Cloning--which involves the creation of DNA from mRNA and thus represents a reversal of the central dogma--is particularly useful when scientists aren't able to isolate the DNA template of a sequence of interest. In addition, because this method relies on mRNA rather than DNA, it provides an excellent means for studying the differences in gene expression in different cells at different points in development. PCR, on the other hand, is more akin to "traditional" DNA synthesis in that it requires the presence of an initial DNA (rather than RNA) template. The primary advantage of PCR is its speed--even if researchers begin with only a single segment of DNA, they can produce literally billions of molecules within a matter of hours. As with all technologies, scientists continue to improve both PCR and the cloning process, thereby ensuring that these methods will play a role in genetic breakthroughs for years to come.

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National Academy of Sciences (US), National Academy of Engineering (US), Institute of Medicine (US) and National Research Council (US) Committee on Science, Engineering, and Public Policy. Scientific and Medical Aspects of Human Reproductive Cloning. Washington (DC): National Academies Press (US); 2002.

Scientific and Medical Aspects of Human Reproductive Cloning.

• Hardcopy Version at National Academies Press

2 Cloning: Definitions And Applications

In this chapter, we address the following questions in our task statement:

What does cloning of animals including humans mean? What are its purposes? How does it differ from stem cell research?

To organize its response to those questions, the panel developed a series of subquestions, which appear as the section headings in the following text.

• WHAT IS MEANT BY REPRODUCTIVE CLONING OF ANIMALS INCLUDING HUMANS?

Reproductive cloning is defined as the deliberate production of genetically identical individuals. Each newly produced individual is a clone of the original. Monozygotic (identical) twins are natural clones. Clones contain identical sets of genetic material in the nucleus—the compartment that contains the chromosomes—of every cell in their bodies. Thus, cells from two clones have the same DNA and the same genes in their nuclei.

All cells, including eggs, also contain some DNA in the energy-generating “factories” called mitochondria. These structures are in the cytoplasm, the region of a cell outside the nucleus. Mitochondria contain their own DNA and reproduce independently. True clones have identical DNA in both the nuclei and mitochondria, although the term clones is also used to refer to individuals that have identical nuclear DNA but different mitochondrial DNA.

• HOW IS REPRODUCTIVE CLONING DONE?

Two methods are used to make live-born mammalian clones. Both require implantation of an embryo in a uterus and then a normal period of gestation and birth. However, reproductive human or animal cloning is not defined by the method used to derive the genetically identical embryos suitable for implantation. Techniques not yet developed or described here would nonetheless constitute cloning if they resulted in genetically identical individuals of which at least one were an embryo destined for implantation and birth.

The two methods used for reproductive cloning thus far are as follows:

• Cloning using somatic cell nuclear transfer (SCNT) [ 1 ]. This procedure starts with the removal of the chromosomes from an egg to create an enucleated egg. The chromosomes are replaced with a nucleus taken from a somatic (body) cell of the individual or embryo to be cloned. This cell could be obtained directly from the individual, from cells grown in culture, or from frozen tissue. The egg is then stimulated, and in some cases it starts to divide. If that happens, a series of sequential cell divisions leads to the formation of a blastocyst, or preimplantation embryo. The blastocyst is then transferred to the uterus of an animal. The successful implantation of the blastocyst in a uterus can result in its further development, culminating sometimes in the birth of an animal. This animal will be a clone of the individual that was the donor of the nucleus. Its nuclear DNA has been inherited from only one genetic parent.

The number of times that a given individual can be cloned is limited theoretically only by the number of eggs that can be obtained to accept the somatic cell nuclei and the number of females available to receive developing embryos. If the egg used in this procedure is derived from the same individual that donates the transferred somatic nucleus, the result will be an embryo that receives all its genetic material—nuclear and mitochondrial—from a single individual. That will also be true if the egg comes from the nucleus donor's mother, because mitochondria are inherited maternally. Multiple clones might also be produced by transferring identical nuclei to eggs from a single donor. If the somatic cell nucleus and the egg come from different individuals, they will not be identical to the nuclear donor because the clones will have somewhat different mitochondrial genes [ 2 ; 3 ]

• Cloning by embryo splitting. This procedure begins with in vitro fertilization (IVF): the union outside the woman's body of a sperm and an egg to generate a zygote. The zygote (from here onwards also called an embryo) divides into two and then four identical cells. At this stage, the cells can be separated and allowed to develop into separate but identical blastocysts, which can then be implanted in a uterus. The limited developmental potential of the cells means that the procedure cannot be repeated, so embryo splitting can yield only two identical mice and probably no more than four identical humans.

The DNA in embryo splitting is contributed by germ cells from two individuals—the mother who contributed the egg and the father who contributed the sperm. Thus, the embryos, like those formed naturally or by standard IVF, have two parents. Their mitochondrial DNA is identical. Because this method of cloning is identical with the natural formation of monozygotic twins and, in rare cases, even quadruplets, it is not discussed in detail in this report.

• WILL CLONES LOOK AND BEHAVE EXACTLY THE SAME?

Even if clones are genetically identical with one another, they will not be identical in physical or behavioral characteristics, because DNA is not the only determinant of these characteristics. A pair of clones will experience different environments and nutritional inputs while in the uterus, and they would be expected to be subject to different inputs from their parents, society, and life experience as they grow up. If clones derived from identical nuclear donors and identical mitocondrial donors are born at different times, as is the case when an adult is the donor of the somatic cell nucleus, the environmental and nutritional differences would be expected to be more pronounced than for monozygotic (identical) twins. And even monozygotic twins are not fully identical genetically or epigenetically because mutations, stochastic developmental variations, and varied imprinting effects (parent-specific chemical marks on the DNA) make different contributions to each twin [ 3 ; 4 ].

Additional differences may occur in clones that do not have identical mitochondria. Such clones arise if one individual contributes the nucleus and another the egg—or if nuclei from a single individual are transferred to eggs from multiple donors. The differences might be expected to show up in parts of the body that have high demands for energy—such as muscle, heart, eye, and brain—or in body systems that use mitochondrial control over cell death to determine cell numbers [ 5 ; 6 ].

• WHAT ARE THE PURPOSES OF REPRODUCTIVE CLONING?

Cloning of livestock [ 1 ] is a means of replicating an existing favorable combination of traits, such as efficient growth and high milk production, without the genetic “lottery” and mixing that occur in sexual reproduction. It allows an animal with a particular genetic modification, such as the ability to produce a pharmaceutical in milk, to be replicated more rapidly than does natural mating [ 7 ; 8 ]. Moreover, a genetic modification can be made more easily in cultured cells than in an intact animal, and the modified cell nucleus can be transferred to an enucleated egg to make a clone of the required type. Mammals used in scientific experiments, such as mice, are cloned as part of research aimed at increasing our understanding of fundamental biological mechanisms.

In principle, those people who might wish to produce children through human reproductive cloning [ 9 ] include:

• Infertile couples who wish to have a child that is genetically identical with one of them, or with another nucleus donor
• Other individuals who wish to have a child that is genetically identical with them, or with another nucleus donor
• Parents who have lost a child and wish to have another, genetically identical child
• People who need a transplant (for example, of cord blood) to treat their own or their child's disease and who therefore wish to collect genetically identical tissue from a cloned fetus or newborn.

Possible reasons for undertaking human reproductive cloning have been analyzed according to their degree of justification. For example, in reference 10 it is proposed that human reproductive cloning aimed at establishing a genetic link to a gametically infertile parent would be more justifiable than an attempt by a sexually fertile person aimed at choosing a specific genome.

Transplantable tissue may be available without the need for the birth of a child produced by cloning. For example, embryos produced by in vitro fertilization (IVF) can be typed for transplant suitability, and in the future stem cells produced by nuclear transplantation may allow the production of transplantable tissue.

The alternatives open to infertile individuals are discussed in Chapter 4 .

• HOW DOES REPRODUCTIVE CLONING DIFFER FROM STEM CELL RESEARCH?

The recent and current work on stem cells that is briefly summarized below and discussed more fully in a recent report from the National Academies entitled Stem Cells and the Future of Regenerative Medicine [ 11 ] is not directly related to human reproductive cloning. However, the use of a common initial step—called either nuclear transplantation or somatic cell nuclear transfer (SCNT)—has led Congress to consider bills that ban not only human reproductive cloning but also certain areas of stem cell research. Stem cells are cells that have the ability to divide repeatedly and give rise to both specialized cells and more stem cells. Some, such as some blood and brain stem cells, can be derived directly from adults [ 12 - 19 ] and others can be obtained from preimplantation embryos. Stem cells derived from embryos are called embryonic stem cells (ES cells). The above-mentioned report from the National Academies provides a detailed account of the current state of stem cell research [ 11 ].

ES cells are also called pluripotent stem cells because their progeny include all cell types that can be found in a postimplantation embryo, a fetus, and a fully developed organism. They are derived from the inner cell mass of early embryos (blastocysts) [ 20 - 23 ]. The cells in the inner cell mass of a given blastocyst are genetically identical, and each blastocyst yields only a single ES cell line. Stem cells are rarer [ 24 ] and more difficult to find in adults than in preimplantation embryos, and it has proved harder to grow some kinds of adult stem cells into cell lines after isolation [ 25 ; 26 ].

Production of different cells and tissues from ES cells or other stem cells is a subject of current research [ 11 ; 27 - 31 ]. Production of whole organs other than bone marrow (to be used in bone marrow transplantation) from such cells has not yet been achieved, and its eventual success is uncertain.

Current interest in stem cells arises from their potential for the therapeutic transplantation of particular healthy cells, tissues, and organs into people suffering from a variety of diseases and debilitating disorders. Research with adult stem cells indicates that they may be useful for such purposes, including for tissues other than those from which the cells were derived [ 12 ; 14 ; 17 ; 18 ; 25 - 27 ; 32 - 43 ]. On the basis of current knowledge, it appears unlikely that adults will prove to be a sufficient source of stem cells for all kinds of tissues [ 11 ; 44 - 47 ]. ES cell lines are of potential interest for transplantation because one cell line can multiply indefinitely and can generate not just one type of specialized cell, but many different types of specialized cells (brain, muscle, and so on) that might be needed for transplants [ 20 ; 28 ; 45 ; 48 ; 49 ]. However, much more research will be needed before the magnitude of the therapeutic potential of either adult stem cells or ES cells will be well understood.

One of the most important questions concerning the therapeutic potential of stem cells is whether the cells, tissues, and perhaps organs derived from them can be transplanted with minimal risk of transplant rejection. Ideally, adult stem cells advantageous for transplantation might be derived from patients themselves. Such cells, or tissues derived from them, would be genetically identical with the patient's own and not be rejected by the immune system. However, as previously described, the availability of sufficient adult stem cells and their potential to give rise to a full range of cell and tissue types are uncertain. Moreover, in the case of a disorder that has a genetic origin, a patient's own adult stem cells would carry the same defect and would have to be grown and genetically modified before they could be used for therapeutic transplantation.

The application of somatic cell nuclear transfer or nuclear transplantation offers an alternative route to obtaining stem cells that could be used for transplantation therapies with a minimal risk of transplant rejection. This procedure—sometimes called therapeutic cloning, research cloning, or nonreproductive cloning, and referred to here as nuclear transplantation to produce stem cells —would be used to generate pluripotent ES cells that are genetically identical with the cells of a transplant recipient [ 50 ]. Thus, like adult stem cells, such ES cells should ameliorate the rejection seen with unmatched transplants.

Two types of adult stem cells—stem cells in the blood forming bone marrow and skin stem cells—are the only two stem cell therapies currently in use. But, as noted in the National Academies' report entitled Stem Cells and the Future of Regenerative Medicine , many questions remain before the potential of other adult stem cells can be accurately assessed [ 11 ]. Few studies on adult stem cells have sufficiently defined the stem cell's potential by starting from a single, isolated cell, or defined the necessary cellular environment for correct differentiation or the factors controlling the efficiency with which the cells repopulate an organ. There is a need to show that the cells derived from introduced adult stem cells are contributing directly to tissue function, and to improve the ability to maintain adult stem cells in culture without the cells differentiating. Finally, most of the studies that have garnered so much attention have used mouse rather than human adult stem cells.

ES cells are not without their own potential problems as a source of cells for transplantation. The growth of human ES cells in culture requires a “feeder” layer of mouse cells that may contain viruses, and when allowed to differentiate the ES cells can form a mixture of cell types at once. Human ES cells can form benign tumors when introduced into mice [ 20 ], although this potential seems to disappear if the cells are allowed to differentiate before introduction into a recipient [ 51 ]. Studies with mouse ES cells have shown promise for treating diabetes [ 30 ], Parkinson's disease [ 52 ], and spinal cord injury [ 53 ].

The ES cells made with nuclear transplantation would have the advantage over adult stem cells of being able to provide virtually all cell types and of being able to be maintained in culture for long periods of time. Current knowledge is, however, uncertain, and research on both adult stem cells and stem cells made with nuclear transplantation is required to understand their therapeutic potentials. (This point is stated clearly in Finding and Recommendation 2 of Stem Cells and the Future of Regenerative Medicine [ 11 ] which states, in part, that “studies of both embryonic and adult human stem cells will be required to most efficiently advance the scientific and therapeutic potential of regenerative medicine.”) It is likely that the ES cells will initially be used to generate single cell types for transplantation, such as nerve cells or muscle cells. In the future, because of their ability to give rise to many cell types, they might be used to generate tissues and, theoretically, complex organs for transplantation. But this will require the perfection of techniques for directing their specialization into each of the component cell types and then the assembly of these cells in the correct proportion and spatial organization for an organ. That might be reasonably straightforward for a simple structure, such as a pancreatic islet that produces insulin, but it is more challenging for tissues as complex as that from lung, kidney, or liver [ 54 ; 55 ].

The experimental procedures required to produce stem cells through nuclear transplantation would consist of the transfer of a somatic cell nucleus from a patient into an enucleated egg, the in vitro culture of the embryo to the blastocyst stage, and the derivation of a pluripotent ES cell line from the inner cell mass of this blastocyst. Such stem cell lines would then be used to derive specialized cells (and, if possible, tissues and organs) in laboratory culture for therapeutic transplantation. Such a procedure, if successful, can avoid a major cause of transplant rejection. However, there are several possible drawbacks to this proposal. Experiments with animal models suggest that the presence of divergent mitochondrial proteins in cells may create “minor” transplantation antigens [ 56 ; 57 ] that can cause rejection [ 58 - 63 ]; this would not be a problem if the egg were donated by the mother of the transplant recipient or the recipient herself. For some autoimmune diseases, transplantation of cells cloned from the patient's own cells may be inappropriate, in that these cells can be targets for the ongoing destructive process. And, as with the use of adult stem cells, in the case of a disorder that has a genetic origin, ES cells derived by nuclear transplantation from the patient's own cells would carry the same defect and would have to be grown and genetically modified before they could be used for therapeutic transplantation. Using another source of stem cells is more likely to be feasible (although immunosuppression would be required) than the challenging task of correcting the one or more genes that are involved in the disease in adult stem cells or in a nuclear transplantation-derived stem cell line initiated with a nucleus from the patient.

In addition to nuclear transplantation, there are two other methods by which researchers might be able to derive ES cells with reduced likeli hood for rejection. A bank of ES cell lines covering many possible genetic makeups is one possibility, although the National Academies report entitled Stem Cells and the Future of Regenerative Medicine rated this as “difficult to conceive” [ 11 ]. Alternatively, embryonic stem cells might be engineered to eliminate or introduce certain cell-surface proteins, thus making the cells invisible to the recipient's immune system. As with the proposed use of many types of adult stem cells in transplantation, neither of these approaches carries anything close to a promise of success at the moment.

The preparation of embryonic stem cells by nuclear transplantation differs from reproductive cloning in that nothing is implanted in a uterus. The issue of whether ES cells alone can give rise to a complete embryo can easily be misinterpreted. The titles of some reports suggest that mouse embryos can be derived from ES cells alone [ 64 - 72 ]. In all cases, however, the ES cells need to be surrounded by cells derived from a host embryo, in particular trophoblast and primitive endoderm. In addition to forming part of the placenta, trophoblast cells of the blastocyst provide essential patterning cues or signals to the embryo that are required to determine the orientation of its future head and rump (anterior-posterior) axis. This positional information is not genetically determined but is acquired by the trophoblast cells from events initiated soon after fertilization or egg activation. Moreover, it is critical that the positional cues be imparted to the inner cells of the blastocyst during a specific time window of development [ 73 - 76 ]. Isolated inner cell masses of mouse blastocysts do not implant by themselves, but will do so if combined with trophoblast vesicles from another embryo [ 77 ]. By contrast, isolated clumps of mouse ES cells introduced into trophoblast vesicles never give rise to anything remotely resembling a postimplantation embryo, as opposed to a disorganized mass of trophoblast. In other words, the only way to get mouse ES cells to participate in normal development is to provide them with host embryonic cells, even if these cells do not remain viable throughout gestation (Richard Gardner, personal communication). It has been reported that human [ 20 ] and primate [ 78 - 79 ] ES cells can give rise to trophoblast cells in culture. However, these trophoblast cells would presumably lack the positional cues normally acquired during the development of a blastocyst from an egg. In the light of the experimental results with mouse ES cells described above, it is very unlikely that clumps of human ES cells placed in a uterus would implant and develop into a fetus. It has been reported that clumps of human ES cells in culture, like clumps of mouse ES cells, give rise to disorganized aggregates known as embryoid bodies [ 80 ].

Besides their uses for therapeutic transplantation, ES cells obtained by nuclear transplantation could be used in laboratories for several types of studies that are important for clinical medicine and for fundamental research in human developmental biology. Such studies could not be carried out with mouse or monkey ES cells and are not likely to be feasible with ES cells prepared from normally fertilized blastocysts. For example, ES cells derived from humans with genetic diseases could be prepared through nuclear transplantation and would permit analysis of the role of the mutated genes in both cell and tissue development and in adult cells difficult to study otherwise, such as nerve cells of the brain. This work has the disadvantage that it would require the use of donor eggs. But for the study of many cell types there may be no alternative to the use of ES cells; for these cell types the derivation of primary cell lines from human tissues is not yet possible.

If the differentiation of ES cells into specialized cell types can be understood and controlled, the use of nuclear transplantation to obtain genetically defined human ES cell lines would allow the generation of genetically diverse cell lines that are not readily obtainable from embryos that have been frozen or that are in excess of clinical need in IVF clinics. The latter do not reflect the diversity of the general population and are skewed toward genomes from couples in which the female is older than the period of maximal fertility or one partner is infertile. In addition, it might be important to produce stem cells by nuclear transplantation from individuals who have diseases associated with both simple [81] and complex (multiple-gene) heritable genetic predilections. For example, some people have mutations that predispose them to “Lou Gehrig's disease” (amyotrophic lateral sclerosis, or ALS); however, only some of these individuals become ill, presumably because of the influence of additional genes. Many common genetic predilections to diseases have similarly complex etiologies; it is likely that more such diseases will become apparent as the information generated by the Human Genome Project is applied. It would be possible, by using ES cells prepared with nuclear transplantation from patients and healthy people, to compare the development of such cells and to study the fundamental processes that modulate predilections to diseases.

Neither the work with ES cells, nor the work leading to the formation of cells and tissues for transplantation, involves the placement of blastocysts in a uterus. Thus, there is no embryonic development beyond the 64 to 200 cell stage, and no fetal development.

2-1. Reproductive cloning involves the creation of individuals that contain identical sets of nuclear genetic material (DNA). To have complete genetic identity, clones must have not only the same nuclear genes, but also the same mitochondrial genes.

2-2. Cloned mammalian animals can be made by replacing the chromosomes of an egg cell with a nucleus from the individual to be cloned, followed by stimulation of cell division and implantation of the resulting embryo.

2-3. Cloned individuals, whether born at the same or different times, will not be physically or behaviorally identical with each other at comparable ages.

2-4. Stem cells are cells that have an extensive ability to self-renew and differentiate, and they are therefore important as a potential source of cells for therapeutic transplantation. Embryonic stem cells derived through nuclear transplantation into eggs are a potential source of pluripotent (embryonic) stem cell lines that are immunologically similar to a patient's cells. Research with such cells has the goal of producing cells and tissues for therapeutic transplantation with minimal chance of rejection.

2-5. Embryonic stem cells and cell lines derived through nuclear transplantation could be valuable for uses other than organ transplantation. Such cell lines could be used to study the heritable genetic components associated with predilections to a variety of complex genetic diseases and test therapies for such diseases when they affect cells that are hard to study in isolation in adults.

2-6. The process of obtaining embryonic stem cells through nuclear transplantation does not involve the placement of an embryo in a uterus, and it cannot produce a new individual.

• Cite this Page National Academy of Sciences (US), National Academy of Engineering (US), Institute of Medicine (US) and National Research Council (US) Committee on Science, Engineering, and Public Policy. Scientific and Medical Aspects of Human Reproductive Cloning. Washington (DC): National Academies Press (US); 2002. 2, Cloning: Definitions And Applications.
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