Gene Transfer Technique

Introduction 

Gene transfer technique is used very widely both in basic research and applied biology. The delivery of DNA into animal cells is a fundamental and established procedure. It has become an indispensable tool for gene cloning, the study of gene function and regulation and the production of small amounts of recombinant proteins for analysis and verification. Gene transfer experiment helps to express the introduced genetic construct (or transgene) in the recipient cells or to disrupt or inactivate particular endogenous genes (resulting in a loss of function. There are many applications of gene transfer like large-scale commercial production of recombinant antibodies and vaccines and gene medicine or gene therapy. They range from the use of mammalian and insect cell cultures to transfer of DNA into human patients for the correction or prevention of disease.

In the organisms or genetically modified whole animals created by Gene transfer every cell or a specific target population of cells carries a particular alteration. Such animals are used to study gene function and expression, model human diseases, produce recombinant proteins in their milk and other fluids, and to improve the quality of livestock herds and other domestic species. The technology has contributed to understand the functions of the many genes discovered in the genome projects (functional genomics). Examples of such experiments include systematic DNA-mediated mutagenesis and gene trap programs in the mouse and in the fruit fly, Drosophila melanogaster, genome-wide RNA interference experiments in the nematode, Caenorhabditis elegans, and novel protein interaction screens based on the yeast two hybrid system but performed using mammalian cells.

Genetic engineering of food is the science which involves deliberate modification of the genetic material of plants or animals. Introduction of DNA into plants is of great agricultural potential and medical importance. The gene transfer results can be transient and stable transfection.

Gene therapy can be defined as the deliberate transfer of DNA for therapeutic purposes. Gene transfer is one of the key factors in gene therapy, and it is one of the key purposes of the clone. Gene transfer can be targeted to somatic (body) or germ (egg and sperm) cells. In somatic gene transfer the recipient’s genome is changed, but the change will not be passed on to the next generation. In germline gene transfer, the parents’ egg and sperm cells are changed with the goal of passing on the changes to their offspring.

 History

The concept of gene transfer between cells was first demonstrated in bacteria, which are capable of at least four natural forms of genetic exchange. The first mechanism was discovered in 1928 by Frederick Griffith named as transformation in bacterium Streptococcus pneumoniae. However, he was unable to determine the nature of the transforming principle. In 1944 Oswald Avery established that the substance transferred between cells was DNA. A second form of gene transfer was named conjugation discovered by Joshua Lederberg and Edward Tatum in 1946 in Escherichia coli. This process involved the transfer of DNA through a direct link between the bacterial cells. In E. coli this conduit between cells took the form of a proteinaceous tube known as a pilus. The ability of the cells to construct the pilus and pass DNA through it was encoded on a large plasmid known as the F (for fertility) factor.

In most cases, the act of conjugation involved transfer of the plasmid alone, which became established in the recipient cell thereby converting it from an F− to an F+ phenotype. In some cases, however, the F plasmid could integrate into the bacterial chromosome, and conjugation could result in the transfer of chromosomal genes. This process, which was used to construct the first genetic map of E. coli, was termed sexduction. Then in 1951, transduction, a new type of gene transfer mediated by bacteriophage was discovered by Joshua Lederberg and Norton Zinder in Salmonella. They found that newly formed phage could occasionally package some of the host cell’s DNA and then transfer it to a second host cell in a subsequent infection.

Two forms of transduction were identified – generalized and specialized transduction. In generalized transduction the phage head was mistakenly stuffed completely with host cell DNA. Whereas in specialized transduction the phage genome integrated into the bacterial chromosome and became linked to host DNA. The fourth mechanism of gene transfer in bacteria is mediated by complete cell fusion, and occurs in several genera of bacteria including Bacillus and Streptomyces. Under natural conditions, the viral capsid is needed to introduce the oncogene and the rest of the genome into an animal host cell. The unusual process of gene transfer without the viral capsid was named transfection to distinguish it from normal infection.

In bacterial genetics, the term transformation continued to be used to describe the uptake of naked plasmid or genomic DNA (essentially any DNA which had the potential to transform the phenotype of the recipient cell) while transfection was used specifically to describe the uptake of naked phage DNA (or RNA), i.e. nucleic acid which had the potential to initiate a phage replication cycle. The term transfection became generally accepted to mean the introduction of any sort of DNA— phage, plasmid, genomic or otherwise—into an animal cell, in the absence of a biological vector.

Different Gene Transfection Techniques

There are two types of transfection transient and stable. In transient transfection, the transfected DNA is not integrated into host chromosome. DNA is transferred into a recipient cell in order to obtain a temporary but high level of expression of the target gene. Stable transfection is also called permanent transfection. By the stable transfection, the transferred DNA is integrated (inserted) into chromosomal DNA and the genetics of recipient cells is permanent changed.

Regardless of the delivery method, gene transfer into animal cells must accomplish three distinct goals. First, the exogenous genetic must be transported across the cell membrane. In physical transfection methods, transport across the membrane is achieved by direct transfer, where the membrane is breached during delivery through which DNA and RNA can diffuse. In other delivery methods, the nucleic acid must form some sort of complex which binds to the cell surface before internalization. For example, in chemical transfection methods the complex is formed between nucleic acid and a synthetic compound, while in transduction methods the complex comprises nucleic acid packaged inside a viral capsid.

Once across the cell membrane, the genetic material must be released in the cell and transported to its site of expression or activity. Again, the nucleic acid is passive at this stage. In most transfection methods, DNA or RNA complexes are deposited in the cytoplasm, following escape from the endosomal vesicle. DNA must be transported to the nucleus, while RNA can function directly in the cytoplasm. In methods such as particle bombardment and microinjection, it is possible to deliver DNA directly into the nucleus, so intrinsic transport pathways are not required. Many viruses also deliver their nucleic acid cargo to the nucleus as part of the infection cycle, often after interaction with cell surface receptors and either internalization within endosomes or direct fusion with the plasma membrane. However, few exceptions are there like poxviruses (e.g. Vaccinia virus) and alphaviruses (e.g. Sindbis virus) which replicate in the cytoplasm. In the final stage of gene transfer, the exogenous genetic material must be activated. It must be released from its complex and rendered competent for expression and/or interaction with the host genome. Exogenous RNA exist only transiently in the host, whereas exogenous DNA can exist transiently or permanently.

The gene transfer methods normally include three categories: 1. transfection by biochemical methods; 2. transfection by physical methods; 3. virus-mediately transduction. The first gene transfer protocols used naked DNA which was mixed with particular chemicals to form synthetic complexes. These synthetic complexes either interact with the cell membrane and promote uptake by endocytosis, or fuse with the membrane and deliver the DNA directly into the cytoplasm. Such chemical transfection methods have been widely used. However they are generally inefficient for gene transfer in vivo. In contrast, physical transfection methods are efficient for both in vitro and in vivo gene transfer.

Physical transfection methods involve breaching the cell membrane and introducing the nucleic acid directly into the cell or nucleus. Although there are advantages and disadvantages to both sets of procedures, some of the most efficient transfection methods in use today involve a combination of chemical and physical processes. Chemical and physical transfection methods were first used for the transfer of naked, wild-type viral DNA into animal cells. Now both techniques are more widely used for the introduction of plasmid vectors and recombinant viral genomes carrying specific transgenes of interest.

Chemical Transfection

Chemical transfection methods have to overcome a number of boundaries to deliver active DNA into the nucleus. They have to persuade the cell to interact with and process exogenous DNA, and eventually deliver at least some intact DNA molecules to the nucleus. The first boundary to gene transfer is the cell membrane, which is hydrophobic and negatively charged whereas DNA, is hydrophilic and negatively charged. DNA can only interact with the cell membrane by a synthetic complex, in combination with DNA that carries a net positive charge or if it is either enclosed in a fusogenic capsule.

The function of the synthetic complex is to form a positively charged complex. The second boundary to successful transfection is DNA transfer to the nucleus, and it is the most difficult step in chemical transfection methods. DNA encapsulated within fusogenic particles is deposited in the cytoplasm under the cell membrane following membrane fusion, and is thought to find its way to the nucleus via an intrinsic transport pathway. However the process is still poorly understood. In contrast, complexes taken up by endocytosis are transported in acidic endosomes, eventually to be degraded in lysosomes. To achieve a high level of transfection efficiency, the DNA must find its way to the nucleus.

Chloroquine are known to help the DNA escape into the cytoplasm by disrupting endosomes and sabotaging the endosomal transport pathway. Also peptides with endosome-disrupting properties or DNA sequence itself can be important for transfer to the nucleus. Vectors carrying the SV40 virus origin of replication are more efficiently transported to the nucleus than similar vectors lacking this sequence. The inclusion of peptides with canonical nuclear localization sequences can also promote the nuclear import of exogenous DNA. The final barrier to efficient transfection is activation of the exogenous DNA by dissociation from the complex once within the cell. As only free DNA can interact with hosts genome so dissociation becomes important.

In an experiment animal oocyte nuclei were injected with naked DNA and DNA lipid complexes to check for the association with hosts genome and the former were capable of transgene expression. It is thought that dissociation occurs either by simple diffusion or neutralization of positively charged complex with negatively charged intracellular lipids and other molecules.

Calcium phosphate Transfection

It was the first chemical transfection method to be used with animal cells. Calcium phosphate is probably the most widely used transfection method. This is a simple, reliable method applicable to many cultured cell lines, and the reagents are inexpensive. It can be used both for transient and stable transformation. The principle of the technique is that DNA in a buffered phosphate solution is mixed gently with calcium chloride, which causes the formation of a fine DNA-calcium phosphate coprecipitate. The precipitate settles onto the cells and some of the particles are taken up by endocytosis. The most efficient transfection occurs in cells growing as a monolayer, because these cells become evenly coated with the precipitate. The procedure was developed in 1973 by Graham and van der Erb for the introduction of adenovirus DNA into rat cells. In a report by Szybalska and Szybalski published in 1962 the presence of calcium was shown to be responsible for the successful transformation of human cells with genomic DNA. The first mammalian cell lines stably transfected with plasmid DNA were also produced by calcium phosphate transfection, in 1978.

Transfection with DEAE-dextran

DEAE-dextran was the first transfection reagent to be developed and was very widely used until the advent of lipofection reagents in the 1990s. It is a soluble polycationic carbohydrate that forms aggregates with DNA through electrostatic interactions. It provides the entire complex with a net positive charge, which allows it to interact with the negatively charged cell membrane and promotes uptake by endocytosis. The complexes are very small compared to the particles formed during calcium phosphate transfection. Also much less DNA can be used in each transfection experiment. Like the calcium phosphate method, the reagents are inexpensive and the procedure is simple and efficient. DEAE-dextran-mediated transfection is not particularly efficient for the production of stably transformed cell lines.

Lipofection

Gene transfer mediated by liposomes was first described by Fengler in 1980. Liposomes are used to form a fusogenic particle with DNA. They are hydrophobic, unilaminar phospholipid vesicles. Such vesicles when mixed with cells in culture, fuse with the cell membrane and deliver DNA directly into the cytoplasm. Lipofection is the most widely used chemical gene transfer method for gene therapy. The disadvantage is the preparation of DNA-containing liposomes is complicated and labor-intensive. One particular advantage of the method was the ability to transform mouse cells in vivo by injecting liposomes into the tail vein. Liposome-mediated transfection was the first non-viral technique devised specifically for in vivo DNA transfer. The efficiency of liposome-mediated gene transfer can be enhanced by incorporating viral proteins that facilitate the active fusion between viral envelopes and cell membranes. Such fusogenic particles have been termed virosomes. Lipofection, is highly reproducible and extremely efficient for both transient and stable transfection.

It allows up to 90% of cells in culture to be transiently transfected. Also demonstrates stable transfection efficiencies up to 20-fold greater than standard chemical transfection methods. One drawback to this approach is that the lipids are usually difficult to prepare in the laboratory. They must therefore be purchased from a commercial source, and they are very expensive. Lipid-based transfection reagents include a positively charged head group, a linker, and a hydrophobic ‘anchor’. The head group generally contains between one and four amine groups, and this is joined either via a glycerol linkage to an aliphatic hydrocarbon chain anchor, or through a variety of linkages to a cholesterol anchor.

Physical Transfection

In physical methods the DNA is delivered directly into either the cytoplasm or the nucleus using some kind of physical force. There is no requirement for interaction with the plasma membrane. This avoids involvement with the endosomal pathway and thus limits the amount of damage sustained by the exogenous DNA. The physical transfection methods are usually expensive s they employ some sort of apparatus, which is required to administer the physical force. The DNA may be introduced free in solution, but it may be beneficial to protect it by forming chemical complexes (e.g. with polyamines) to reduce the damaging effects of shear forces during the transfer process.

Electroporation

This method was first used with animal cells by Neumann and colleagues in 1982. Electroporation is the transfection of cells following their exposure to a pulsed electric field. This causes a number of nanometer-sized pores to open in the plasma membrane for up to 30 minutes, allowing the uptake of free DNA from the surrounding medium. Afterwards, the pores close spontaneously with no noticeable adverse effects on the treated cells. It is ideal for many established cell lines, especially those recalcitrant to chemical transfection methods. The standard electroporation procedure is very simple. Cells are suspended in or flooded with an electroporation buffer and exposed to a brief, high voltage electrical pulse.

The magnitude and duration of the pulse determine the transfection efficiency, and these conditions must be established empirically for different cell lines. It is efficient, highly reproducible, suitable for both stable and transient transfection, and has the added advantage that transgene copy number can be at least partially controlled. Disadvantages of this technique include the requirement for specialized capacitor discharge equipment capable of accurately controlling pulse length and voltage the requirement for larger numbers of cells and higher DNA concentrations than used in chemical transfection methods, and the rather high level of cell death that accompanies this procedure. Electroporation has also been explored as a method for in vivo gene transfer. It has been used with success to introduce DNA into surface or near-surface tissues such as skin, muscle and melanoma, and also into internal organs such as the liver. Among the commercial devices available for electroporation, at least one has been designed specifically for electroporation-mediated gene therapy in humans.

Electroporation was initially only applicable to cells cultured in suspension, but it can also be used with cells growing in monolayers using an adaptation of the technique. Most recently, a modification of the technique known as nucleofection has been described. This method combines specific electroporation conditions with different transfection reagents to promote direct electroporation-mediated gene transfer to the nucleus, resulting in the efficient transfection of traditionally difficult targets including primary cells. Another in vitro transfection technology called as laser poration is based on pore formation, using laser treatment. This involves a similar DNA uptake mechanism to electroporation, i.e. free DNA is taken up directly from the surrounding medium through transient pores created by a finely focused laser beam. Like microinjection (see below) this strategy can be applied only to a small number of cells at a time, but with optimal DNA concentration can result in stable transfection frequencies of greater than 0.5%.

Microinjection

The direct microinjection of DNA into the cytoplasm or nuclei of cultured cells is sometimes used as a transfection method. It is highly efficient at the level of individual cells. The most significant use of this technique is introduction of DNA into the oocytes, eggs and embryos of animals, either for transient expression analysis (e.g. in fish or Xenopus) or to generate transgenic animals (e.g. mice, Drosophilathis). The procedure is time consuming and only a small number of cells can be treated. Originally, this technique was used for the transformation of cells that were resistant to any other method of transfection. Stable transfection efficiencies are extremely high, in the order of 20%, and very small quantities of DNA are sufficient.

This technique provides direct nuclear delivery of DNA avoiding the endogenous pathway and also ensures that the DNA is delivered intact. Microinjection is suitable for the introduction of large vectors such as YACs into the pronuclei of fertilized mouse eggs. DNA delivered in this manner must be very pure so it needs a lot of preparation as it is necessary to avoid fragmentation. Shearing can also occur in the delivery needle, and large DNA fragments are often protected by suspension in a high salt buffer and/or mixing with polyamines and other protective agents. Now transfection of cultured cells is automated with computer-controlled micromanipulation and microinjection processes as well as the automated production of injection capillaries and the standardization of cell preparation procedure.

Transfection by particle bombardment

Particle bombardment (also known as biolistics or microprojectile transfection) procedure involves coating micrometer-sized gold or tungsten particles with DNA and then accelerating the particles into cells or tissues. A major advantage of this method is that DNA can be delivered to deep cells in tissue slices, and the depth of penetration can be adjusted by changing the applied force. The size and total mass of the particles and the force of the bombardment are important parameters that balance efficient penetration against cell damage. The technique was developed for the transformation of maize and is now a method of choice for generating transgenic cereal plants. For animal cells, the technique has been less widely used because it is usually simpler to transfect cultured cells by alternative well-established methods. However, the technique has found a role in the transfection of whole organs and tissue slices, and more recently for the transfer of DNA to surface organs in gene therapy.

Transfection by ultrasound

It involves the exposure of cells to a rapidly oscillating probe, such as the tip of a sonicator. The application of ultrasound waves to a dish of cells or a particular tissue results in the formation and collapse of bubbles in the liquid, including the cell membrane, a process known as cavitation. The transient appearance of such cavities allows DNA to cross the membrane into the cytoplasm. It has been shown that the application of low-frequency ultrasound allows the efficient delivery of nucleic acids into mammalian cells both in vitro and in vivo, because the plasmid DNA is left structurally intact. Furthermore, the ultrasound waves appear to have no adverse effects when focused on different anatomic locations in the human body. Hence, ultrasound-mediated gene delivery raises no safety concerns. Gene transfer in vivo is generally achieved by injection followed by the application of a focused ultrasound device.

Virus-mediated transduction

Viruses have evolved to deliver nucleic acids safely into animal cells. The enveloped and non- enveloped viruses follow different way to interacting with the membrane. In the case of enveloped viruses they deliver nucleic acid by binding to specific receptors on the cell surface and then by either fusing directly with the plasma membrane or, following uptake by endocytosis, by fusing to the endosomal membrane.

In contrast, non-enveloped viruses penetrate or disrupt the plasma or endosomal membranes with specific virion proteins. In each case, the viral genome ends up being uncoated in the cytoplasm and transported to its normal replication site which may or may not be nucleus. The transfer of exogenous nucleic acid into animal cells as part of a recombinant viral particle is known as transduction. The advantages of viral transduction methods over transfection with naked plasmid DNA include the high efficiency of gene transfer as it follows a natural delivery process. Different viruses are used for different transformation objectives –

1. Adenoviruses for short-term infections with high-level transient expression
2. erpesviruses for long-term expression
3. etroviruses for stable integration of DNA into the host cell genome

The adenoviruses are non-enveloped DNA viruses. They possess an icosahedral capsid about 100 nm in diameter containing a double-stranded 36kbp linear genome. Of the two known genera, only the mastadenoviruses can infect mammalian cells. Six subgroups of human adenoviruses have been defined based on cellular agglutination assays and genomic GC content. The major advantages of adenoviral vectors are that they can be purified to extremely high titers, which makes them highly suited for in vivo applications, and the efficiency of gene transfer approaches 100% if the target cells bear the appropriate receptors.

The herpesviruses are large, enveloped viruses with linear, double-stranded DNA genomes varying in length from 100–200 kbp. The virion structure consists of an outer lipid envelope studded with a large number of glycoproteins and other proteins, surrounding a matrix of proteins known as the tegument, which overlies the capsid proper. The complexity of the lipid envelope means that the herpesviruses demonstrate considerable variability in their host range and cell tropism. The two herpesviruses developed as vectors are Epstein-Barr virus (EBV) and herpes simplex virus (HSV1). EBV has a very narrow host range and tropism (limited mainly to human Blymphocytes and a few other cell types), so it is not much use as a general transduction vector. However, the EBV genome will replicate in many cells following transfection, wherein it is maintained as an episomal replicon.

The EBV genome has therefore been used as a basis of a series of large-capacity plasmid vectors for episomal transgene expression in human cells. In contrast, HSV1 has a very wide host range and cell tropism, and has been developed into a versatile transduction vector. Following infection, the HSV genome remains within the nucleus of the host cell indefinitely, i.e. it has a life-long latent infection cycle. Other advantages of HSV1 as a vector include its large capacity for foreign DNA, its efficient infection of neurons, and the ability of replication-competent viruses to cross synapses. HSV–1 has a broad host range and cell tropism because glycoproteins C and B, embedded in the envelope, interact with ubiquitous glucosaminoglycans on the cell surface, e.g. heparan sulfate and dermatan sulfate.

Retroviruses are enveloped RNA viruses, which comprise a proteinacious virion approximately 100 nm in diameter surrounded by a lipid bilayer. Each viral particle carries two copies of a single-stranded, positive-sense RNA genome as well as several proteins required for infection. There are seven genera of retroviruses based on data from sequence comparisons. The conventional oncoretroviruses, such as murine leukemia virus (MLV) are simple. Whereas lentiviruses, such as human immunodeficiency virus (HIV) and spumaviruses are complex.

Lentiviruses contain additional genes compared to the basic oncoretrovirus genome. The replication strategy of the retroviruses is unique. After entering the cell, the virus is uncoated and the genomic RNA is transported to the nucleus where it is converted into a terminally redundant double-stranded cDNA copy by the virion protein reverse transcriptase. A second virion protein, integrase, then inserts this cDNA copy into the host genome. The host range of each retrovirus species is determined by proteins in the lipid envelope. Some retroviruses have a very restricted host range while others have a broad host range because the envelope proteins interact with more widely distributed receptors.

Retroviruses are advantageous vectors they have the ability to produce high viral titers (106 –108 particles ml−1) using readily available packaging lines, the incredible efficiency of stable transduction (approaching 100% in vitro, but also very high in vivo) and the ability to pseudotype viral particles and thus engineer the host range of each vector. The small viral genome is easy to manipulate in the laboratory once it has been converted into a cDNA copy, and it carries a useful promoter/enhancer system, which can be used to drive transgene expression.

 The basic technique for introducing DNA into E. coli have inspired procedures for the introduction of DNA into cells from a wide variety of organisms, including mammalian cells.  Gene transfer techniques for higher plants and animals are complex and costly. It is generally not done in laboratory scale experiment. However, the techniques of gene transfer in bacteria like Escherichia coli (E. coli) are simple and appropriate for the teaching and learning laboratory. Plasmid-based genetic transformation is the most common used technology for genetic transfer. It enables manipulation of genetic information in a laboratory setting and it makes the understanding of how DNA operates. In the following protocol, we will look at the tools to transform E. coli bacteria to express new genetic information using a plasmid system and apply mathematical routines to determine transformation efficiency.

Preparation

Materials and Equipment

Commercial suppliers for plasmid transformation systems may be purchased in kits. These plasmids should contain the gene for ampicillin resistance (pBLU), as experimental procedures typically use ampicillin to select transformed cells. In addition, plasmids with colored marker genes like beta-GAL and fluorescence markers like green fluorescent protein (GFP) and its cousins make it possible to measure gene expression directly, to follow cell populations as they grow or move, and to find cells that have taken up a second plasmid that we cannot see easily.

The following materials are included in a typical eight-station ampicillin-resistant plasmid system.

Materials

1. E.coli (1 vial or slant)
2. Plasmid (pBLU), hydrated (20 μg)
3. Ampicillin, lyophilized (30 μg)
4. Transformation solution (50mM CaCl2, pH 6), sterile (15 mL)
5. LB nutrient agar powder, sterile (to make 500 mL) (20 g) or prepared agar
6. LB nutrient broth, sterile (10 mL)
7. Pipettes, sterile (50)
8. Inoculation loops, sterile (10 μL, packs of 10 loops)
9. Petri dishes, sterile, 60 mm (packs of 20)
10. Multicolor 2.0 mL microcentrifuge tubes (60)
11. Microcentrifuge tube holders
12. Clock or watch to time 50 seconds
13. Microwave oven/water bath
14. Thermometer that reads 42°C
15. 1 L flask
16. 500 mL graduated cylinder
17. Distilled water
18. Crushed ice and containers
19. 10% solution household bleach
20. Permanent marker pens
21. Masking tape
22. Biohazardous waste disposal bags or plastic trash bags
23. Micropipettes, adjustable volume, 2–20 μL (and pipette tips)
24. Parafilm laboratory sealing film
25. 37°C incubator oven*

Preparation

Advance Preparation Quick Guide for Teachers

Step       Objective                                            Time Required                          When

Step 1    Prepare agar plates.                           1 hr.                                           3–7 days prior

Step 2   Rehydrate E. coli.                                 2 min                                         24–36 hours prior

Streak starter plates.                                         2 min                                         24–36 hours prior

Rehydrate plasmid DNA                                   15 min.                                       24–36 hours prior.

Step 3   Aliquot solutions.                                10 min.                                        Immediately prior

Advance Preparation for Step 1: 3–7 Days before the Transformation

1. Prepare nutrient agar (autoclave-free).

The agar plates should be prepared at least three days before the investigation(s) are performed. Plates should be left out at room temperature for two days and then refrigerated until use. (Two days at room temperature allows the agar to cure, or dry, sufficiently to readily take up the liquid transformation solution.) If time is short, incubate the plates at 37°C overnight. This will dry them out as well, but it shortens their shelf life.  Refrigerated plates are good for up to 30 days.

To prepare the agar, add 500 mL of distilled water to a one liter or larger Erlenmeyer flask. Add the entire content of the LB nutrient agar packet. Swirl the flask to dissolve the agar and heat to boiling in a microwave or water bath or by using a hot plate with stir bar. Heat and swirl until all the agar is dissolved.

When all the agar is dissolved, allow the LB nutrient agar to cool so that the outside of the flask is just comfortable to hold (approximately 50°C.). While the agar is cooling, you can label the plates and prepare the ampicillin as outlined below in Step 3.

Pre-prepared nutrient agar also can be purchased. However, it will have to be melted before it can be poured into plates. To do this, the plastic bottles containing solid agar can be microwaved at a low temperature (such as using the “poultry defrost” option) for several minutes. Be sure to loosen the cap slightly to expel any air. At high microwave temperatures, the agar can boil over. Another option is to  place the bottles in a hot water bath; however, this will take up to 45 minutes or so to melt the agar.

2. Prepare ampicillin.

Ampicillin is either shipped dry in a small vial or already hydrated. If shipped dry, you need to hydrate the ampicillin. Do this by adding 3 mL of transformation solution to the vial to rehydrate the antibiotic. Use a sterile pipette. The nutrient agar solidifies at 27°C, so you must be careful to monitor the cooling of the agar and then pour the plates from start to finish without interruption. Keeping the flask with liquid agar in a water bath set to 45–50°C can help prevent the agar from cooling too quickly. Before adding ampicillin to the flask of agar, make sure you can hold the flask in your bare hand (approximately 50°C). If your hand tolerates the temperature of the flask, so will the antibiotic!

3. Label plates.

While the agar is cooling, reduce preparation time by labeling the plates. Label with a permanent marker on the bottom of each plate close to the edge. For each class using an eight-station kit, label 16 plates LB and 16 plates LB/amp.

4. Pour nutrient agar plates.    

 First, pour LB nutrient agar into the 16 plates that are labeled LB. If you do not do this and add ampicillin to the flask with agar, you will not be able to make control plates containing just nutrient agar. Fill each plate to about one-third to one-half (approximately 12 mL) with agar and replace the lid. You may want to stack the plates and let them cool in the stacked configuration. Add the hydrated ampicillin to the remaining LB nutrient agar. Swirl briefly to mix. Pour into the 16 plates labeled LB/amp using the same technique. Plates should set within 30 minutes.

4. Store the plates.

After the plates have cured for two days at room temperature, they may be either used or stored by stacking them in a plastic sleeve bag slipped back down over them. The stack is then inverted, the bag taped closed, and the plates stored upside down at 4°C until used. (The plates are inverted to prevent condensation on the lid, which may drip onto the agar.)

Advance Preparation for Step 2: 24–36 Hours before the Transformation

1. Rehydrate bacteria.

Some E. coli cultures come prepared in a slant and will not have to be rehydrated. For bacteria that must be rehydrated, use a sterile pipette to add 250 μL of transformation solution directly to the vial. Recap the vial and allow the cell suspension to stand at room temperature for 5 minutes. Then shake the mix before streaking on the LB starter plates. Store the rehydrated bacteria in the refrigerator until used (within 24 hours for best results and no longer than three days).

2. Streak starter plates.

Starter plates are needed to produce bacterial colonies of E. coli on agar plates. Each lab team will need its own starter plate as a source of cells for transformation. LB plates should be streaked for single colonies and incubated at 37°C for 24–26 hours before the transformation investigation begins.

Using E. coli and LB agar plates, streak one starter plate to generate single colonies from a concentrated suspension of bacteria. A small amount of the bacterial suspension goes a long way. Under favorable conditions, one cell multiples to become millions of genetically identical cells in just 24 hours. There are millions of individual bacteria in a single millimeter of a bacterial colony.

1. Insert a sterile inoculation loop straight into the vial of rehydrated bacterial culture. Remove the loop and streak the plates. Streaking takes place sequentially in four quadrants. The first streak spreads out the cells. Go back and forth with the loop about a dozen times in each of the small areas shown. In subsequent quadrants, the cells become more and more dilute, thus increasing the likelihood of producing single colonies.
2. For subsequent streaks, use as much of the surface area of the plate as possible. After the initial streak, rotate the plate approximately 45 degrees and start the second streak. Do not dip into the rehydrated bacteria a second time. Go into the previous streak about two times and then back and forth for a total of about 10 times.
3. Rotate the plate again and repeat streaking.
4. Rotate the plate for the final time and make the final streak. Repeat steps a–c with the remaining LB plates for each student workstation. Although you can use the same inoculation loop for all starter plates, it is recommended that you use a new, sterile loop for each plate if you have enough. When you are finished with each plate, cover it immediately to avoid contamination.
5. Place the plates upside down inside the incubator overnight at 37°C or at room temperature for 2–3 days if an incubator is unavailable. Use for transformation within 24–36 hours because bacteria must be actively growing to achieve high transformation efficiency. (Remember, bacterial growth is exponential.) Do not refrigerate before use. This will slow bacterial growth.
6. coli forms off-white colonies that are uniformly circular with smooth edges. Avoid using plates with contaminant colonies such as mold.

3. Prepare plasmid.

The quantity of DNA is so small that the vial may appear empty. Tap the vial or spin it in a microcentrifuge to ensure that the DNA is not sticking to the cap. If the plasmid is not hydrated, refer to instructions that come with the sample. Store the vial of hydrated DNA in a refrigerator. Rehydrated plasmid should be used within 24 hours.

 Advance Preparation for Step 3: Immediately Before Transformation Investigation

1. Aliquot solutions.
2. Each student workstation will need 1 mL of transformation solution and 1 mL of LB nutrient broth. You might have to aliquot these solutions into separate color-coded 2 mL microtubes. If the LB nutrient broth is aliquoted one day prior to the lab, it should be refrigerated. Make sure to label the tubes with permanent marker.
3. Set up the workstations. See the list of materials required. If the plasmid goes through multiple freeze-thaw cycles in a frost-free freezer, the DNA in the plasmid can degrade. It is recommended that you check the shelf life of materials with the commercial vendor.

Procedure

The plasmid likely will contain the gene for resistance to ampicillin (pBLU) antibiotic that is lethal to many bacteria, including E. coli cells. This transformation procedure involves the following three main steps to introduce the plasmid DNA into the E. coli cells and to provide an environment for the cells to express their newly acquired genes:

1. Adding CaCl2
2. “Heat shocking” the cells
3. Incubating the cells in nutrient broth for a short time before plating them on agar

Materials

1. coli starter plate prepared.
2. Poured agar plates prepared.
3. 2 LB agar plates
4. 2 LB/amp agar (LB agar containing ampicillin) plates
5. Transformation solution (CaCl2, pH 6.1) kept ice cold
6. LB nutrient broth
7. Sterile inoculation loops
8. 100–1000 μL sterile bulb pipettes
9. 1–10 μL micropipettes with sterile tips
10. Microcentrifuge tubes
11. Microcentrifuge tube holder/float
12. Container full of crushed ice
13. Marking pen
14. DNA plasmid (0.005 μg/μL)
15. 42°C water bath and thermometer
16. 37°C incubator
17. 20 μL adjustable-volume micropipettes and tips (optional)
18. 10% household bleach
19. Biohazardous waste disposal bags
20. Masking or lab tape

Step 1

Label one closed microcentrifuge tube (micro test tube) “+ plasmid” and one tube “-plasmid.” Label both tubes, and place them in the microcentrifuge tube holder/float.

Step 2

Carefully open the tubes and, using a 100–1000 μL bulb pipette with a sterile tip, transfer 250 μL of the ice cold transformation solution (CaCl2) into each tube.

Step 3

Place both tubes on (into) the ice.

Step 4

Use a sterile inoculation loop to pick up a single colony of bacteria from your starter plate. Be careful not to scrape off any agar from the plate. Pick up the “+ plasmid” tube and immerse the loop into the CaCl2 solution (transforming solution) at the bottom of the tube. Spin the loop between your index finger and thumb until the entire colony is dispersed in the solution.

Step 5

Use a new sterile 100–1,000 μL micropipette to repeatedly pulse the cells in solution to thoroughly resuspend the cells. Place the tube back on the ice.

Step 6

Using a new sterile inoculation loop, repeat Steps 5 and 6 for the “- plasmid” tube.

Step 7

Using a 1–10 μL micropipette with a sterile tip, transfer 10 μL of the plasmid solution directly into the E. coli suspension in the “+ plasmid” tube. Tap tube with a finger to mix, but avoid making bubbles in the suspension or splashing the suspension up the sides of the tube. Do not add the plasmid solution into the “- plasmid” tube.

Step 8

Incubate both tubes (“+ plasmid” and “- plasmid”) on ice for 10 minutes. Make sure the bottom of the tubes make contact with the ice.

Step 9

While the tubes are sitting on ice, label each of your agar plates on the bottom.

Step 10

Following the 10-minute incubation at 0°C, remove the tubes from the ice and “heat shock” the cells in the tubes. It is critical that the cells receive a sharp and distinct shock.Make sure the tubes are closed tightly! Place the tubes into a test tube holder/ float, and dunk the tubes into the water bath, set at 42°C, for exactly 50 seconds. Make sure to push the tubes all the way down in the holder so that the bottom of the tubes with the suspension makes contact with the warm water.

Step 11

When the 50 seconds have passed, place both tubes back on ice. For best transformation results, the change from 0°C to 42°C and then back to 0°C must be rapid. Incubate the tubes on ice for an additional two minutes.

Step 12

Remove the holder containing the tubes from the ice and place on the lab counter. Using a 100–1,000 μL micropipette with sterile tip, transfer 250 μL of LB nutrient broth to the “+ plasmid” tube. Close the tube and gently tap with your finger to mix. Repeat with a new sterile micropipette for the “- plasmid” tube.

Step 13

Incubate each tube for 10 minutes at room temperature.

Step 14

Use a 10–1,000 μL micropipette with sterile tip to transfer 100 μL of the transformation (“+ plasmid”) and control (“- plasmid”) suspensions onto the appropriate LB and LB/Amp plates. Be sure to use a separate pipette for each of the four transfers.

Step 15

Using a new sterile inoculation loop for each plate, spread the suspensions evenly around the surface of the agar by quickly “skating” the flat surface of the sterile loop back and forth across the plate surface. Do not poke or make gashes in the agar. Allow the plates to set for 10 minutes.

Step 16

Stack your plates and tape them together. Place the stack upside down in the 37°C incubator for 24 hours

Analyzing Results

By calculating transformation efficiency, you can measure the success of your transformation quantitatively.

Calculating Transformation Efficiency

Transformation efficiency = Total number of colonies growing on the agar plate/ Amount of DNA spread on the agar plate (in μg)

This quantitative measurement of transformation is the transformation efficiency. In many applications, it is important to transform as many cells as possible. For example, in some forms of gene therapy, cells are collected from the patient, transformed in the laboratory, and then put back into the patient. The more cells that are transformed to produce the needed protein, the more likely the therapy will work..

Calculating transformation efficiency gives you an indication of how effective you were in getting plasmids carrying new information into host bacterial cells. In this example, transformation efficiency is a number that represents the total number of bacterial cells that express the gene for ampicillin resistance divided by the amount of DNA plasmid used in the experiment. The transformation efficiency is calculated using the following formula.