Plasmid Cloning: Steps, Mechanisms, and Applications

By Dr. Zubair Khalid, DVM, MS, PhD · 2026-08-11

Introduction to Plasmid Cloning

Plasmid cloning is the process of inserting a foreign DNA fragment into a plasmid vector, propagating that recombinant molecule within a host organism, and isolating multiple identical copies of the inserted sequence. This technique forms the backbone of recombinant DNA technology and underpins nearly every branch of modern molecular biology, from gene function studies to the commercial production of therapeutic proteins.

What is a Plasmid?

A plasmid is a small, circular, double-stranded DNA molecule that exists independently of the chromosomal DNA within a cell. Plasmids are naturally found in bacteria and some eukaryotes, where they carry genes that confer advantageous traits such as antibiotic resistance, metabolic capabilities, or virulence factors. For a detailed treatment of plasmid biology, see the Plasmid Definition entry.

In the laboratory, naturally occurring plasmids have been engineered into cloning vectors—vehicles designed to accept, replicate, and express foreign DNA. These engineered plasmids typically range from 2 to 10 kilobases (kb) in size and are maintained at copy numbers ranging from a few to several hundred per cell, depending on the origin of replication used.

Why Clone DNA into Plasmids?

Cloning DNA into plasmids serves several fundamental purposes:

1. Amplification: A single DNA fragment can be replicated millions of times within overnight bacterial cultures, yielding sufficient material for downstream applications such as sequencing, probing, or further subcloning.
2. Storage: Plasmids provide a stable, long-term repository for DNA fragments. Bacterial stocks containing recombinant plasmids can be frozen indefinitely and revived when needed.
3. Manipulation: Cloned DNA can be mutagenized, tagged, or recombined to study gene function, protein localization, or regulatory elements.
4. Expression: Plasmids can be engineered to drive high-level production of recombinant proteins in bacterial, yeast, insect, or mammalian hosts.
5. Delivery: Recombinant plasmids serve as delivery vehicles for gene therapy, vaccine development, and the generation of transgenic organisms.

Key Components of a Cloning Vector

A functional plasmid cloning vector must contain several essential elements. Understanding each component is critical for designing successful cloning experiments.

Origin of Replication (ori)

The origin of replication is a specific DNA sequence that recruits the replication machinery of the host cell. This sequence determines two critical properties of the plasmid: copy number and host range.

The pMB1 origin (derived from the naturally occurring ColE1 plasmid) is used in the common pUC and pBluescript vectors. These plasmids replicate at 500–700 copies per cell because the pUC series carries a point mutation in the RNA II primer that enhances replication initiation. High copy number is advantageous for maximizing DNA yield but can be problematic when expressing toxic proteins, as the high gene dosage can overwhelm the host.

The p15A origin, found in vectors such as pACYC184, maintains a copy number of approximately 10–15 per cell. This is useful for complementation studies where a second plasmid must coexist with a high-copy vector, provided the two origins are compatible (i.e., they do not interfere with each other's replication).

The pSC101 origin maintains only 5 copies per cell and is useful for cloning genes whose overexpression is lethal.

Selectable Markers (Antibiotic Resistance)

A selectable marker is a gene that allows only cells harboring the plasmid to survive under specific growth conditions. The most common selectable markers confer resistance to antibiotics. The marker is essential because plasmid uptake during transformation is inefficient—typically only 1 in 10,000 to 1 in 1,000,000 cells acquires a plasmid. Without selection, plasmid-free cells would rapidly outgrow transformants.

Common antibiotic resistance markers include:

Ampicillin is widely used but has a practical limitation: β-lactamase is secreted into the culture medium, where it degrades the antibiotic. Satellite colonies—cells that do not contain the plasmid—can therefore grow in the zone surrounding true transformants. Carbenicillin, a more stable ampicillin derivative, is often preferred. For a deeper discussion of marker selection, see Selectable Marker in Plasmid.

Multiple Cloning Site (MCS)

The multiple cloning site, also called a polylinker, is a short DNA sequence (typically 50–100 bp) containing multiple unique restriction enzyme recognition sites arranged in tandem. The MCS is the insertion point for foreign DNA. Because each restriction site appears only once in the vector, digestion with a single enzyme linearizes the plasmid at a defined position without fragmenting it.

The MCS is usually embedded within a reporter gene (most commonly lacZ, encoding β-galactosidase) to enable blue-white screening, described in detail below.

Optional Elements (e.g., reporter genes, tags)

Beyond the minimal requirements, many cloning vectors include additional features that expand their utility:

• Promoters: Sequences such as T7, T5, or lac drive transcription of the cloned gene. The T7 promoter requires a host strain (e.g., BL21(DE3)) that carries the T7 RNA polymerase gene under an inducible promoter.
• Fusion tags: Sequences encoding peptides such as polyhistidine (His₆), glutathione S-transferase (GST), maltose-binding protein (MBP), or green fluorescent protein (GFP) can be fused to the cloned gene to facilitate protein purification or detection.
• Reporter genes: lacZ, gfp, or luc (luciferase) allow visual or quantitative assessment of cloning success or gene expression.
• Phage origins: The f1 origin of replication allows production of single-stranded DNA when the host is superinfected with helper phage, useful for site-directed mutagenesis or phage display.

The Plasmid Cloning Workflow

The standard plasmid cloning procedure follows a defined sequence of steps. While variations exist—such as Golden Gate Cloning or Topo Ta Cloning Kit methods—the classical restriction-ligation approach remains the conceptual foundation.

Step 1: Isolate Vector and Insert DNA

The vector plasmid is purified from an overnight bacterial culture using alkaline lysis followed by column-based purification. The insert DNA can be obtained by:

• PCR amplification from genomic DNA, cDNA, or another plasmid, using primers that incorporate restriction sites at their 5′ ends.
• Restriction digestion of a larger DNA molecule to release the fragment of interest.
• Chemical synthesis of the desired sequence (gene synthesis).

Both vector and insert must be pure and free of nucleases. Quantify DNA by spectrophotometry (A₂₆₀) or fluorometry; a typical cloning reaction uses 50–100 ng of vector and a 3:1 to 5:1 molar ratio of insert to vector.

Step 2: Restriction Enzyme Digestion

The vector and insert are digested with restriction enzymes that generate compatible ends. The choice of enzymes depends on the restriction sites present in the MCS and the ends of the insert. Digestion is performed in the manufacturer's recommended buffer at the optimal temperature (usually 37°C) for 1–2 hours.

After digestion, the enzymes are heat-inactivated (where possible) or removed by column purification. The digested vector is often treated with calf intestinal alkaline phosphatase (CIP) or shrimp alkaline phosphatase (SAP) to remove 5′ phosphate groups, preventing self-ligation of the vector (see Section 4).

Step 3: Ligation of Insert into Vector

The purified, digested vector and insert are combined in a ligation reaction containing T4 DNA ligase and ATP. The enzyme catalyzes the formation of a phosphodiester bond between the 3′ hydroxyl of one DNA strand and the 5′ phosphate of an adjacent strand.

A typical ligation reaction (20 µL total volume) contains:

• 50–100 ng vector DNA
• 3:1 to 5:1 molar excess of insert
• 1× T4 DNA ligase buffer (containing 1 mM ATP and 10 mM MgCl₂)
• 1–2 units T4 DNA ligase (for sticky ends) or 10–20 units (for blunt ends)
• Incubation at 16°C for 1–4 hours, or overnight at 4°C

Step 4: Transformation into Host Cells

The ligation mixture is introduced into competent Escherichia coli cells. Competent cells are bacteria that have been treated to increase their permeability to foreign DNA. Two common methods are chemical transformation (heat shock) and electroporation, both described in Section 5.

After transformation, cells are plated on selective medium containing the appropriate antibiotic. Only cells that have acquired a plasmid carrying the resistance gene will form colonies.

Step 5: Selection and Screening of Recombinant Clones

Colonies that grow on selective medium contain plasmid DNA, but not necessarily the desired insert. The vector may have self-ligated (if dephosphorylation was incomplete), or the insert may have ligated in the wrong orientation. Screening methods—colony PCR, restriction digest analysis, and sequencing—are used to identify correct clones (Section 6).

Restriction Enzymes and Ligation

Restriction Enzyme Specificity

Restriction enzymes (restriction endonucleases) recognize specific, usually palindromic, DNA sequences of 4–8 base pairs and cleave the phosphodiester backbone within or near that sequence. Type II restriction enzymes, the workhorses of molecular cloning, cleave at defined positions within their recognition site.

For example, *Eco*RI recognizes the sequence 5′-GAATTC-3′ and cleaves between G and A on both strands, producing a 4-base 5′ overhang. *Bam*HI recognizes 5′-GGATCC-3′ and similarly produces a 5′ overhang. *Sma*I recognizes 5′-CCCGGG-3′ and cleaves blunt-ended.

The frequency of restriction sites in random DNA depends on recognition site length: a 6-base cutter will occur approximately once every 4⁶ = 4,096 bp, while an 8-base cutter occurs once every 65,536 bp.

Sticky vs. Blunt End Ligation

Sticky (cohesive) ends are generated when a restriction enzyme makes staggered cuts, leaving short single-stranded overhangs. These overhangs hydrogen-bond with complementary overhangs on another DNA molecule, holding the fragments together while ligase seals the nicks. Sticky-end ligation is efficient because the complementary overhangs stabilize the interaction.

Blunt ends are generated when the enzyme cuts straight across both strands. Ligation of blunt ends requires higher concentrations of ligase and insert DNA because there are no overhangs to guide annealing. The efficiency of blunt-end ligation is roughly 10- to 100-fold lower than sticky-end ligation.

Dephosphorylation and Directional Cloning

To prevent the vector from self-ligating (recircularizing without an insert), the 5′ phosphate groups are removed from the digested vector using alkaline phosphatase. DNA ligase requires a 5′ phosphate on one of the two DNA molecules being joined; without it, the vector cannot circularize. The insert retains its 5′ phosphates, so it can ligate to the dephosphorylated vector. This reduces the background of non-recombinant colonies by 90–95%.

Directional cloning uses two different restriction enzymes to generate incompatible ends on the vector and insert. Because the two ends cannot ligate to each other, the insert can only be placed in one orientation. For example, digesting the vector with *Bam*HI and *Hind*III and the insert with the same enzymes ensures that the insert's *Bam*HI end ligates only to the vector's *Bam*HI end, and the *Hind*III end only to the *Hind*III end.

Transformation and Selection

Chemical Transformation (Heat Shock)

Chemical transformation uses calcium chloride to make bacterial cells competent for DNA uptake. The divalent calcium ions neutralize the negative charge of the lipopolysaccharide layer and the DNA phosphate backbone, allowing DNA to associate with the cell surface. A brief heat shock (42°C for 30–90 seconds) creates a thermal gradient that drives DNA uptake.

The standard protocol:

1. Thaw competent cells (stored at −80°C) on ice for 5–10 minutes.
2. Add 1–10 µL of ligation mixture (containing 10–100 ng DNA) to 50–100 µL of cells.
3. Incubate on ice for 30 minutes.
4. Heat shock at 42°C for exactly 45 seconds.
5. Return to ice for 2 minutes.
6. Add 900 µL of pre-warmed SOC or LB broth (without antibiotic).
7. Incubate at 37°C with shaking (225 rpm) for 1 hour to allow expression of the antibiotic resistance gene.
8. Plate 50–200 µL on selective agar.

Transformation efficiency for chemically competent cells is typically 10⁶–10⁸ colony-forming units (CFU) per microgram of supercoiled plasmid DNA.

Electroporation

Electroporation uses a brief, high-voltage electrical pulse (typically 1.8 kV, 25 µF, 200 Ω for E. coli) to create transient pores in the bacterial membrane through which DNA enters. Electrocompetent cells are prepared by washing log-phase cultures extensively in cold 10% glycerol to remove salts that would conduct electricity.

Electroporation is 10- to 100-fold more efficient than chemical transformation (10⁸–10¹⁰ CFU/µg), making it the method of choice when DNA is limiting or when transforming large plasmids (>10 kb). The cells must be kept ice-cold and the electroporation cuvette (0.1 cm or 0.2 cm gap) must be chilled to prevent arcing.

Antibiotic Selection

After transformation, cells are plated on agar containing the antibiotic corresponding to the plasmid's resistance marker. Only cells that have acquired the plasmid can form colonies. The antibiotic concentration must be sufficient to kill plasmid-free cells but not so high as to inhibit the growth of transformants.

Standard concentrations for E. coli:

• Ampicillin: 50–100 µg/mL
• Kanamycin: 25–50 µg/mL
• Chloramphenicol: 25–34 µg/mL
• Tetracycline: 10–15 µg/mL

Blue-White Screening (lacZ)

Blue-white screening distinguishes recombinant clones (containing an insert) from non-recombinants (vector only) based on the disruption of the lacZ gene. The MCS is located within the coding sequence of lacZα, a short fragment of β-galactosidase.

When the vector is intact, lacZα is expressed, and the α-fragment complements a truncated β-galactosidase encoded on the host chromosome (in strains such as DH5α or JM109). The active enzyme converts the chromogenic substrate X-gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside, 40 µg/mL) into a blue precipitate.

When an insert is cloned into the MCS, lacZα is disrupted, no functional β-galactosidase is produced, and the colonies remain white. IPTG (isopropyl β-D-1-thiogalactopyranoside, 0.5 mM) is added to induce lacZα expression.

White colonies are candidate recombinants, but not all white colonies contain the correct insert—some may contain self-ligated vector with a small deletion in lacZα. Therefore, white colonies must be verified by additional screening.

Verifying Successful Cloning

Colony PCR

Colony PCR is a rapid method to determine whether a bacterial colony contains the desired insert. A small portion of a colony is transferred directly into a PCR reaction containing primers that flank the MCS (e.g., M13 forward and reverse primers) or primers specific to the insert. The initial denaturation step (95°C for 5 minutes) lyses the cells and releases the plasmid DNA.

The PCR product is analyzed by agarose gel electrophoresis. A band of the expected size indicates the presence of the insert. This method is fast (2–3 hours) and can screen dozens of colonies simultaneously.

Restriction Digest Analysis

Restriction digest analysis confirms both the presence and the orientation of the insert. Plasmid DNA is purified from a small overnight culture (miniprep) and digested with restriction enzymes that cut asymmetrically within the vector or insert. The resulting fragments are separated by agarose gel electrophoresis.

For example, if the insert contains an internal *Eco*RI site and the vector has a single *Eco*RI site in the MCS, digestion of the recombinant plasmid will produce two fragments whose sizes confirm the insert's presence. To determine orientation, digest with an enzyme that cuts once in the vector and once in the insert; the fragment sizes will differ depending on whether the insert is in the forward or reverse orientation.

DNA Sequencing

DNA sequencing is the definitive method for verifying a clone. Sanger sequencing using primers that anneal to the vector backbone (e.g., T7, SP6, M13 forward/reverse) reads across the MCS and into the insert. This confirms:

• The insert is present.
• The sequence is correct (no mutations introduced by PCR).
• The insert is in the correct reading frame (if expression is intended).
• The insert is in the correct orientation.

Modern sequencing services provide results within 24–48 hours and are inexpensive enough to be used routinely.

Applications of Plasmid Cloning

Protein Expression

Plasmid cloning enables the production of recombinant proteins in heterologous hosts. The gene of interest is cloned downstream of a strong, inducible promoter. In E. coli, the T7 promoter system is most common: the gene is cloned into a vector such as pET series, and expression is induced with IPTG in a host strain (BL21(DE3)) that carries T7 RNA polymerase. The protein can be purified using affinity tags such as His₆.

Gene Knockout and Knock-in

Plasmids are used to construct targeting vectors for gene disruption in bacteria, yeast, and mammalian cells. In bacteria, a plasmid carrying a modified gene with a selectable marker flanked by homology arms can replace the chromosomal gene via homologous recombination. In mammalian cells, similar targeting constructs are used to create knockout mice or cell lines.

Vaccine Development

Plasmid DNA vaccines deliver genes encoding antigens directly into host cells, where they are expressed and trigger an immune response. Plasmid cloning is used to construct these vaccines, which offer advantages over traditional vaccines including rapid production, stability, and the ability to stimulate both humoral and cellular immunity.

Biopharmaceutical Production

Many therapeutic proteins are produced by cloning the corresponding gene into expression plasmids and introducing them into production hosts. Insulin (recombinant human insulin, produced in E. coli or Saccharomyces cerevisiae), human growth hormone, erythropoietin, and monoclonal antibodies are all manufactured using plasmid-based expression systems. The Ti Plasmid has been adapted for plant transformation, enabling the production of recombinant proteins in transgenic plants.

Common Pitfalls and Troubleshooting

Low Ligation Efficiency

Symptoms: Few or no colonies after transformation.

Causes and solutions:

• Insufficient insert DNA: Increase the insert:vector molar ratio to 5:1 or 7:1. Calculate the required amount using the formula: ng insert = (ng vector × insert size in kb / vector size in kb) × molar ratio.
• Damaged or missing 5′ phosphates on insert: If the insert was generated by PCR, the primers must be phosphorylated at the 5′ end, or the PCR product must be treated with polynucleotide kinase (PNK) before ligation.
• Inactive ligase or degraded ATP: Use fresh ligase and buffer. Avoid multiple freeze-thaw cycles of the buffer.
• Incompatible ends: Verify that the restriction enzymes used generate compatible overhangs. Some enzymes produce ends that are not compatible with each other even if they recognize similar sequences.
• Ligation temperature too high: Sticky-end ligations should be performed at 16°C, not 37°C, to maintain the stability of the annealed overhangs.

High Background of Non-recombinants

Symptoms: Many colonies, but most lack the insert (blue colonies in blue-white screening, or white colonies with no insert).

Causes and solutions:

• Incomplete vector dephosphorylation: Increase the amount of alkaline phosphatase or extend the incubation time. Ensure the enzyme is heat-inactivated before ligation.
• Incomplete restriction digestion: Verify complete digestion by running an aliquot of the digested vector on a gel. A single band at the linear size confirms complete digestion.
• Religation of vector without insert: Always include a no-insert control ligation to assess background. If background is high, re-purify the digested vector by gel extraction to remove undigested circular plasmid.

Incorrect Insert Orientation

Symptoms: Clones contain the insert, but it is in the reverse orientation relative to the promoter.

Causes and solutions:

• Using a single restriction enzyme for both ends: This produces identical ends, allowing the insert to ligate in either orientation. Use two different enzymes (directional cloning) to force the correct orientation.
• Blunt-end cloning: Blunt ends have no orientation. Use sticky-end directional cloning or screen multiple clones by colony PCR with one primer annealing to the vector and one to the insert.

Poor Transformation Efficiency

Symptoms: Few or no colonies even with a positive control plasmid.

Causes and solutions:

• Cells not properly competent: Check the transformation efficiency of the competent cells using a known supercoiled plasmid (e.g., pUC19). Efficiency should be at least 10⁶ CFU/µg for chemically competent cells.
• Too much DNA in the transformation: Excess DNA can inhibit transformation. Use 10–100 ng of plasmid DNA per 50 µL of competent cells.
• Antibiotic concentration too high: Verify the correct concentration for the antibiotic and the host strain.
• Recovery time too short: Allow at least 45–60 minutes of outgrowth in non-selective medium to permit expression of the resistance gene.
• Heat shock temperature or duration incorrect: Use a calibrated water bath or heat block. 42°C for 45 seconds is optimal for most E. coli strains.

Summary and Best Practices

Quick Reference Checklist

1. Design: Choose a vector with the appropriate origin, marker, and promoter for your application. Verify that the MCS contains unique restriction sites compatible with your insert.
2. Prepare insert: Design primers with restriction sites and additional 3–6 flanking bases to ensure efficient enzyme digestion. Purify the PCR product before digestion.
3. Digest: Use 1–2 µg of vector and a 3–5-fold molar excess of insert. Include a vector-only control. Heat-inactivate or purify after digestion.
4. Dephosphorylate: Treat the vector with alkaline phosphatase to prevent self-ligation.
5. Ligate: Use a 3:1 (sticky) or 5:1 (blunt) insert:vector molar ratio. Include a no-ligase control and a vector-only control.
6. Transform: Use high-efficiency competent cells. Include a positive control (supercoiled plasmid) and a negative control (no DNA).
7. Screen: Pick 8–12 colonies for colony PCR. Confirm positive clones by restriction digest and sequencing.
8. Archive: Prepare glycerol stocks of confirmed clones and store at −80°C.

Final Tips for Beginners

• Always include controls: A ligation without insert, a transformation without DNA, and a transformation with a known plasmid are essential for interpreting results.
• Use fresh reagents: Restriction enzymes, ligase, and ATP in buffers degrade over time. Check expiration dates and store at −20°C.
• Be patient with screening: Not every white colony contains the correct insert. Screen enough colonies (at least 8–12) to find a correct one.
• Read the manufacturer's protocols: Restriction enzyme buffers differ in salt composition and pH. Using the wrong buffer can result in star activity (non-specific cleavage).
• Keep a detailed lab notebook: Record all plasmid names, primer sequences, digestion conditions, and results. This is invaluable for troubleshooting and for reproducing experiments.

Frequently Asked Questions

What is plasmid cloning?

Plasmid cloning is the process of inserting a foreign DNA fragment into a circular plasmid vector, introducing the recombinant plasmid into a host cell (typically E. coli), and propagating it to produce many copies of the inserted DNA. The technique enables gene amplification, protein expression, and genetic manipulation.

What are the steps of plasmid cloning?

The steps are: (1) isolate vector and insert DNA, (2) digest both with restriction enzymes, (3) ligate the insert into the vector using T4 DNA ligase, (4) transform the ligation mixture into competent bacterial cells, and (5) select and screen for recombinant clones using antibiotic resistance, blue-white screening, colony PCR, restriction digest, and sequencing.

How does plasmid cloning work?

Plasmid cloning works by exploiting the ability of restriction enzymes to cut DNA at specific sequences and DNA ligase to join DNA fragments. The vector and insert are cut with the same restriction enzymes to generate complementary ends. The insert is ligated into the vector, and the recombinant plasmid is introduced into bacteria, where it replicates independently of the chromosome. Antibiotic selection ensures that only cells carrying the plasmid survive.

What are examples of plasmid cloning?

Examples include: cloning the human insulin gene into a pET vector for expression in E. coli; constructing a GFP fusion plasmid to study protein localization; creating a knockout construct with a kanamycin resistance cassette flanked by homology arms; and building a DNA vaccine plasmid carrying an antigen gene for immunization studies.

Why is antibiotic selection used in plasmid cloning?

Antibiotic selection is used because plasmid uptake during transformation is inefficient. The plasmid carries an antibiotic resistance gene, and plating on antibiotic-containing medium ensures that only cells that have acquired the plasmid can grow. This eliminates the vast majority of cells that did not take up DNA.

What is blue-white screening?

Blue-white screening is a method to distinguish recombinant clones from non-recombinants. The vector carries lacZα, a fragment of β-galactosidase, with the MCS inserted within it. On medium containing X-gal and IPTG, intact lacZα produces blue colonies. When an insert disrupts lacZα, no functional enzyme is produced, and colonies remain white.

How do you troubleshoot low ligation efficiency?

Check the insert:vector molar ratio (increase to 5:1), verify that the insert has 5′ phosphate groups (phosphorylate PCR primers or treat with PNK), use fresh ligase and buffer, confirm that the restriction enzyme digestion was complete, and ensure the ligation is performed at the correct temperature (16°C for sticky ends).

What is the difference between sticky and blunt end cloning?

Sticky end cloning uses restriction enzymes that generate complementary single-stranded overhangs, which anneal during ligation and increase efficiency. Blunt end cloning uses enzymes that cut straight across both strands, producing no overhangs. Blunt end ligation requires higher concentrations of ligase and insert DNA and is less efficient, but it allows joining of any two blunt-ended fragments regardless of sequence.

Key Takeaways

• Plasmid cloning is the insertion of foreign DNA into a plasmid vector, followed by propagation in a host organism to amplify the DNA or express the encoded gene.
• A functional cloning vector requires an origin of replication, a selectable marker (usually antibiotic resistance), and a multiple cloning site; optional elements include promoters, fusion tags, and reporter genes.
• The classical workflow involves restriction digestion, ligation, transformation, and screening; each step has defined conditions and controls that must be optimized.
• Restriction enzymes generate sticky or blunt ends; sticky-end ligation is more efficient, and dephosphorylation of the vector prevents self-ligation.
• Transformation is achieved by chemical heat shock or electroporation; antibiotic selection and blue-white screening identify cells containing recombinant plasmids.
• Verification of clones requires colony PCR, restriction digest analysis, and DNA sequencing to confirm insert presence, orientation, and sequence accuracy.
• Plasmid cloning underpins protein production, gene knockout studies, vaccine development, and biopharmaceutical manufacturing, making it an essential skill in molecular biology.

Further Reading

• Mead DA, Kemper B. Chimeric single-stranded DNA phage-plasmid cloning vectors. Biotechnology (Reading, Mass.). 1988. PubMed 3061525
• Schmidhauser TJ, Ditta G, Helinski DR. Broad-host-range plasmid cloning vectors for gram-negative bacteria. Biotechnology (Reading, Mass.). 1988. PubMed 2850044
• Chen F et al. Simplified plasmid cloning with a universal MCS design and bacterial in vivo assembly. BMC biotechnology. 2021. PubMed 33722223
• Ding T et al. Reversed paired-gRNA plasmid cloning strategy for efficient genome editing in Escherichia coli. Microbial cell factories. 2020. PubMed 32156270
• Linder JE et al. Sequencing human rhinoviruses: direct sequencing versus plasmid cloning. Journal of virological methods. 2015. PubMed 25286177
• Miura K, Takishima K. Reliable PCR-based method for cloning cDNA in plasmid vectors by frequency estimation. BioTechniques. 1999. PubMed 10090969

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