1. Unit 3
Gene transfer techniques: Physical, chemical and Biological methods, Competent
cells: Chemical and Electro-competent. Introduction of DNA into host cells.
Screening and characterization of transformants; Selectable marker genes, reporter
genes. Expression of recombinant proteins using bacterial, animal and plant vectors
and their purification. Transformation/ transfection in plants and animals.
2. GENE TRANSFER TECHNIQUES: BIOLOGICAL
METHODS
The main gene transfer methods using biological means are as follows:
• Bacterial gene delivery i.e. bactofection.
• Delivery using a viral vector i.e. transduction
Bactofection
It is a method of direct gene transfer using bacteria into the target cell, tissue,
organ or organism.
Various bacterial strains that can be used as vectors in gene therapy are listed in the
following table. The genes located on the plasmids of the transformed bacterial
strains are delivered and expressed into the cells.
The gene delivery may be intracellular or extracellular.
It has a potential to express various plasmid-encoded heterologous proteins
(antigens, toxins, hormones, enzymes etc.) in different cell types. Strains that are
invasive and having better cell to cell spread are more efficient.
3. Bactofection in various disease models.
The main problem of bactofection is the possibility of unwanted side effects related to
the host–bacteria interactions. The response of the immune system might cause rapid
clearance of bacteria or even autoimmune reactions. On contrary, the bacterial strains
can acquire the virulence factors back and might cause serious infections. Therefore,
to reduce the risk of clinically symptomatic infections to minimum, the bacteria are
genetically modified.
4. The process of
bactofection
The process of bactofection
(a) the transformed bacterial strain with plasmid containing transgene is transferred to
target cell
(b) genetically engineered bacteria penetrates into the cell
(c) In the cytoplasm, the vector undergoes lysis and get destructed releasing plasmids
(d)The released plasmids enter into the nucleus where the transgene is expressed by
eukaryotic transcription and translation machinery.
5. Transduction
This method involves the introduction of genes into host cell’s genome
using viruses as carriers.
The viruses are used in gene transfer due to following features-
• Efficiency of viruses to deliver their nucleic acid into cells
• High level of replication and gene expression.
The foreign gene is packaged into the virus particles to enter the host cell.
The entry of virus particle containing the candidate gene sequences into
the cell and then to the nuclear genome is a receptor- mediated process.
The vector genome undergoes complex processes ending up with ds-DNA
depending on the vector that can persist as an episome or integrate into the
host genome followed by the expression of the candidate gene
7. Viral vectors as therapeutic
agents
Viruses have paved a way into clinical field in order to treat
cancer, inherited and infectious diseases.
They can be used as vectors to deliver a therapeutic gene into the
infected cells.
They can be genetically engineered to carry therapeutic gene
without having the ability to replicate or cause disease.
8. GENE TRANSFER TECHNIQUES: CHEMICAL METHODS
• Cell membrane is a sheet like assembly of amphipathic molecules that separate
cells from their environment.
• These physical structures allow only the controlled exchange of materials among
the different parts of a cell and with its immediate surroundings.
• DNA is an anionic polymer, larger molecular weight, hydrophilic and sensitive to
nuclease degradation in biological matrices. They cannot easily cross the physical
barrier of membrane and enter the cells unless assisted.
Various charged chemical compounds can be used to facilitate DNA transfer directly
to the cell. These synthetic compounds are introduced near the vicinity of recipient
cells thereby disturbing the cell membranes, widening the pore size and allowing the
passage of the DNA into the cell.
An ideal chemical used for DNA transfer should have the ability to-
• Protect DNA against nuclease degradation.
• Transport DNA to the target cells.
• Facilitate transport of DNA across the plasma membrane.
• Promote the import of DNA into the nucleus.
9. The commonly used methods of chemical
transfection use the following,
1. Calcium phosphate
2. DEAE dextran
3. Cationic Lipid
4.Other polymers - poly-L-lysine (PLL),
polyphosphoester, chitosan, dendrimers
10. Calcium phosphate
transfection
This method is based on the precipitation of plasmid DNA and calcium ions by
their interaction.
In this method, the precipitates of calcium phosphate and DNA being small and
insoluble can be easily adsorbed on the surface of cell.
This precipitate is engulfed by cells through endocytosis and the DNA gets
integrated into the cell genome resulting in stable or permanent transfection.
Optimal factors (amount of DNA in the precipitate, the length of time for
precipitation reaction and exposure of cells to the precipitate) need to be
determined for efficient transfection of the cells.
This technique is simple, expensive and has minimal cytotoxic effect but the low
level of transgene expression provoked development of several other methods of
transfection.
https://www.nature.com/articles/nmeth0405-319
12. Advantages
• Simple and inexpensive
• Applicability to generate stably transfected cell lines
•Highly efficient (cell type dependent) and can be
applied to a wide range of cell types.
• Can be used for stable or transient transfection
Disadvantages
• Toxic especially to primary cells
•Slight change in pH, buffer salt concentration and
temperature can compromise the efficacy
•Relatively poor transfection efficiency compared to
other chemical transfection methods like
13. DEAE-Dextran (Diethylaminoethyl Dextran) mediated DNA
transfer
Diethylaminoethyl dextran (DEAE-dextran) is a soluble polycationic carbohydrate that promotes
interactions between DNA and endocytotic machinery of the cell.
The cationic DEAE-dextran molecule tightly associates with the negatively charged backbone of the
nucleic acid, and the net positive charge of the resulting nucleic acid-DEAE-dextran complex (A slight
excess of DEAE – dextran in mixture results in net positive charge in the DEAE – dextran/ DNA complex
formed) allows it to adhere to the cell membrane and enter into the cytoplasm via endocytosis or
osmotic shock induced by DMSO or glycerol.
Several parameters such as number of cells, polymer concentration, transfected DNA concentration and
duration of transfection should be optimized for a given cell line.
Advantages
• Simple and inexpensive, • More sensitive, • Can be applied to a wide range of cell types
• Can be used for transient transfection.
Disadvantages
•Toxic to cells at high concentrations, • Transfection efficiency varies with cell type, • Can only be used
for transient transfection but not for stable transfection, • Typically produces less than 10% delivery in
primary cells.
Another polycationic chemical, the detergent Polybrene, has been used for the transfection of Chinese
hamster ovary (CHO) cells, which are not amenable to calcium phosphate transfection.
14. Lipofection
Lipofection is a method of transformation first described in 1965 as a model
of cellular membranes using liposomes.
Liposomes are artificial phospholipid vesicles used for the delivery of a
variety of molecules into the cells. They may be multi-lamellar or
unilamellar vesicles with a size range of 0.1 to 10 micrometer or 20-25
nanometers respectively.
They can be preloaded with DNA by two common methods- membrane-
membrane fusion and endocytosis thus forming DNA- liposome complex.
This complex fuses with the protoplasts to release the contents into the
cell.
Animal cells, plant cells, bacteria, yeast protoplasts are susceptible to
lipofection method.
Liposomes can be classified as either cationic liposome or pH-sensitive
liposomes
15. Cationic liposomes
Cationic liposomes are positively charged liposomes which associate with the
negatively charged DNA molecules by electrostatic interactions forming a stable
complex.
Neutral liposomes are generally used as DNA carriers and helpers of cationic liposomes
due to their non-toxic nature and high stability in serum. A positively charged lipid is often
mixed with a neutral co-lipid, also called helper lipid to enhance the efficiency of gene
transfer by stabilizing the liposome complex (lipoplex).
Dioleoylphosphatidyl ethanolamine (DOPE) or dioleoylphosphatidyl choline (DOPC)
are some commonly used neutral co-lipids.
•The negatively charged DNA molecule interacts with the positively charged groups of the
DOPE or DOPC. DOPE is more efficient and useful than DOPC due to the ability of its inverted
hexagonal phase to disrupt the membrane integrity.
•The overall net positive charge allows the close association of the lipoplex with the
negatively charged cell membrane followed by uptake into the cell and then into nucleus.
•The lipid: DNA ratio and overall lipid concentration used in the formation of these
complexes is particularly required for efficient gene transfer which varies with application.
16. Negatively charged liposomes
Generally pH-sensitive or negatively-charged liposomes are not efficient for gene
transfer. They do not form a complex with it due to repulsive electrostatic
interactions between the phosphate backbone of DNA and negatively charged
groups of the lipids. Some of the DNA molecules get entrapped within the aqueous
interior of these liposomes.
However, formation of lipoplex, a complex between DNA and anionic lipids can
occur by using divalent cations (e.g. Ca2+, Mg2+, Mn2+, and Ba2+) which can
neutralize the mutual electrostatic repulsion. These anionic lipoplexes comprise
anionic lipids, divalent cations, and plasmid DNA which are physiologically safe
components.
They are termed as pH sensitive due to destabilization at low pH.
The efficiency of both in vivo and in vitro gene delivery using cationic liposomes is
higher than that of pH sensitive liposomes. But the cationic liposomes get
inactivated and unstable in the presence of serum and exhibit cytotoxicity. Due to
reduced toxicity and interference from serum proteins, pH-sensitive liposomes
are considered as potential gene delivery vehicles than the cationic liposomes.
17. Schematic representation of liposome
action in gene transfer
Advantages
• Economic
•Efficient delivery of nucleic acids to
cells in a culture dish.
• Delivery of the nucleic acids with
minimal toxicity.
• Protection of nucleic acids from
degradation.
•Measurable changes due to
transfected nucleic acids in sequential
processes.
• Easy to use, requirement of minimal
steps and adaptable to high-
throughput systems.
Disadvantages
• It is not applicable to all cell types.
• It fails for the transfection of some
cell lines with lipids.
18. GENE TRANSFER TECHNIQUES: PHYSICAL OR
MECHANICAL METHODS
It has been discussed earlier that due to amphipathic
nature of the phospholipid bilayer of the plasma
membrane, polar molecules such as DNA and protein
are unable to freely pass through the membrane.
Various physical or mechanical methods are employed
to overcome this and aid in gene transfer as listed
below-
1. Electroporation
2. Microinjection
3. Particle Bombardment
4. Sonoporation
5. Laser induced
6. Bead transfection
19. Electroporation
•Electroporation is a mechanical method used for the introduction
of polar molecules into a host cell through the cell membrane.
•This method was first demonstrated by Wong and Neumann in
1982 to study gene transfer in mouse cells.
• It is now a widely used method for the introduction of transgene
either stably or transiently into bacterial, fungal, plant and animal
cells.
•It involves use of a large electric pulse that temporarily disturbs
the phospholipid bilayer, allowing the passage of molecules such as
DNA.
The basis of electroporation is the relatively weak hydrophobic
/hydrophilic interaction of the phospholipids bilayer and ability to
spontaneously reassemble after disturbance. A quick voltage shock
may cause the temporary disruption of areas of the membrane and
allow the passage of polar molecules. The membrane reseals leaving
the cell intact soon afterwards.
20. Procedure
The host cells and the DNA molecules to be transported into the cells are
suspended in a solution.
When the first switch is closed, the capacitor charges up and stores a high voltage
which gets discharged on closing the second switch.
Typically, 10,000-100,000 V/cm in a pulse lasting a few microseconds to a
millisecond is essential for electroporation which varies with the cell size.
This electric pulse disrupts the phospholipid bilayer of
the membrane causing the formation of temporary
aqueous pores.
When the electric potential across the cell membrane is
increased by about 0.5-1.0 V, the charged molecules
e.g. DNA migrate across the membrane through the
pores in a similar manner to electrophoresis.
The initiation of electroporation generally occurs when
the transmembrane voltage reaches at 0.5-1.5 V. The
cell membrane discharges with the subsequent flow of
the charged ions and molecules and the pores of the
membrane quickly close reassembling the phospholipid
bilayer
21. Applications
Electroporation is widely used in many areas of molecular biology and in medical field.
Some applications of electroporation include:
DNA transfection or transformation
Electroporation is mainly used in DNA transfection/transformation which
involves introduction of foreign DNA into the host cell (animal, bacterial or plant
cell).
Direct transfer of plasmids between cells
It involves the incubation of bacterial cells containing a plasmid with another strain
lacking plasmids but containing some other desirable features. The voltage of
electroporation creates pores, allowing the transfer of plasmids from one cell to another.
This type of transfer may also be performed between species. As a result, a large number of
plasmids may be grown in rapidly dividing bacterial colonies and transferred to yeast cells
by electroporation.
Gene transfer to a wide range of tissues
Electroporation can be performed in vivo for more efficient gene transfer in a wide range of
tissues like skin, muscle, lung, kidney, liver, artery, brain, cornea etc. It avoids the vector-
specific immune-responses that are achieved with recombinant viral vectors and thus are
promising in clinical applications.
22. Advantages
•It is highly versatile and effective for nearly all cell types and
species.
•It is highly efficient method as majority of cells take in the
target DNA molecule.
•It can be performed at a small scale and only a small
amount of DNA is required as compared to other
methods.
Disadvantages
•Cell damage is one of the limitations of this method caused
by irregular intensity pulses resulting in too large pores
which fail to close after membrane discharge.
•Another limitation is the non-specific transport which may
result in an ion imbalance causing improper cell function
and cell death.
23. Microinjection
DNA microinjection was first proposed by Dr. Marshall A. Barber in the early of
nineteenth century.
This method is widely used for gene transfection in mammals.
It involves delivery of foreign DNA into a living cell (e.g. a cell, egg, oocyte,
embryos of animals) through a fine glass micropipette. The introduced DNA may
lead to the over or under expression of certain genes.
It is used to identify the characteristic function of dominant genes.
Procedure
The delivery of foreign DNA is done under a powerful microscope using a glass
micropipette tip of 0.5 mm diameter.
Cells to be microinjected are placed in a container. A holding pipette is placed in
the field of view of the microscope that sucks and holds a target cell at the tip.
The tip of micropipette is injected through the membrane of the cell to deliver the
contents of the needle into the cytoplasm and then the empty needle is taken out.
24. Advantages
• No requirement of a marker
gene.
•Introduction of the target gene
directly into a single cell.
•Easy identification of transformed
cells upon injection of dye along
with the DNA.
•No requirement of selection of
the transformed cells using
antibiotic resistance or
herbicide resistance markers.
•It can be used for creating
transgenic organisms, particularly
mammals.
25. Particle bombardment
Prof Sanford and colleagues at Cornell University (USA) developed the original
bombardment concept in 1987 and coined the term “biolistics” (short for “biological
ballistics”) for both the process and the device.
Also termed as particle bombardment, particle gun, micro projectile bombardment
and particle acceleration.
It employs high-velocity micro projectiles to deliver substances into cells and
tissues.
Applications:
This method is commonly employed for genetic transformation of plants and
many organisms.
This method is applicable for the plants having less regeneration capacity and
those which fail to show sufficient response to Agrobacterium- mediated gene
transfer in rice, corn, wheat, chickpea, sorghum and pigeon-pea.
26. APPARATUS
The biolistic gun employs the principle of conservation of momentum and uses the
passage of helium gas through the cylinder with arrange of velocities required for optimal
transformation of various cell types. It consists of a bombardment chamber which is
connected to an outlet for vacuum creation. The bombardment chamber consists of a
plastic rupture disk below which macro carrier is loaded with micro carriers. These micro
carriers consist of gold or tungsten micro pellets coated with DNA for transformation.
The apparatus is placed in Laminar flow while
working to maintain sterile conditions. The target
cells/tissue is placed in the apparatus and a
stopping screen is placed between the target
cells and micro carrier assembly. The passage of
high pressure helium ruptures the plastic rupture
disk propelling the macro carrier and micro
carriers.
The stopping screen prevents the passage of
macro projectiles but allows the DNA coated
micro pellets to pass through it thereby,
delivering DNA into the target cells
27. Advantages
•Simple and convenient method involving coating DNA or RNA on to gold
microcarrier, loading sample cartridges, pointing the nozzle and firing the device.
• No need to obtain protoplast as the intact cell wall can be penetrated.
• Manipulation of genome of sub-cellular organelles can be done.
•Eliminates the use of potentially harmful viruses or toxic chemical treatment as
gene delivery vehicle.
• This device offers to place DNA or RNA exactly where it is needed into any
organism.
Disadvantages
•The transformation efficiency may be lower than Agrobacterium- mediated
transformation.
• Specialized equipment is needed. Moreover the device and consumables are
costly.
• Associated cell damage can occur.
• The target tissue should have regeneration capacity.
• Random integration is also a concern.
• Chances of multiple copy insertions could cause gene silencing
28. Sonoporation
Sonoporation involves the use of ultrasound for temporary
permeabilization of the cell membrane allowing the uptake of
DNA, drugs or other therapeutic compounds from the extracellular
environment.
This method leaves the compound trapped inside the cell after
ultrasound exposure.
It employs the acoustic cavitation of micro bubbles for enhancing
the delivery of large molecules like DNA. The micro bubbles form
complex with DNA followed by injection and ultrasound treatment
to deliver DNA into the target cells.
Unlike other methods of transfection, sonoporation combines the
capability to enhance gene and drug transfer.
29. Sonoporation
Rupture of microbubbles by
ultrasound resulting in enhanced
membrane permeability caused
by shear stress,
increased temperature and
activation
of reactive oxygen species. Drug
delivery by microbubbles by
(a)transient holes induced by shear
stress for drug delivery
(b) increase in membrane fluidity
(c) endocytosis of microbubbles
(d) microbubble- cell
membrane fusion.
30. Laser induced transfection
• It involves the use of a brief pulse of focused laser beam.
•In this method, DNA is mixed with the cells present in the culture and then
a fine
focus of laser beam is passed on the cell surface that forms a small pore
sufficient
for DNA uptake into the cells. The pore thus formed is transitory and
repairs soon.
Bead transfection
•Bead transfection combines the principle of physically producing breaks in
the cellular membrane using beads.
• In this method, the adherent cells are incubated for a brief period with
glass beads in a solution containing the DNA.
•The efficiency of this rapid technique depends on:
o Concentration of DNA in a solution.
o Timing of the addition of DNA.
o Size and condition of the beads and the buffers utilized.
31. Introduction of DNA into host cells
The delivery of DNA into the host is required for generation
of genetically modified organism. DNA delivery to host is a 3
stage process, DNA sticking to the host cell, internalization
and release into the host cell. As a result, it depends on 2
parameters-
• Surface chemistry of host cell-Host cell surface charges
either will attract or repel DNA as a result of opposite
or similar charges. Presence of cell wall (in the case of
bacteria, fungus and plant) causes additional physical
barrier to the up-take and entry of DNA.
• Charges on DNA-Negative charge on DNA
modulates interaction with the host cell especially
cell surface.
Modulation of these two properties is achieved in
32. Transformation
• Transformation is the natural process, through which bacterial population
transfer the genetic material to acquire phenotypic features.
• The event of transformation was first time demonstrated by Frederick
Griffith in 1928.
The schematic presentation of the experiment is given in Figure 20.1. Griffith has
used two different Streptococcus pneumonia strains, virulent (S, causes disease
and death of mice) and avirulent (R, incapable of causing disease or death of
mice). In a simple experiment he injected 4 different combination of bacterial
mixture,
(1) live S, (2) heat killed S, (3) live R, (4) mixture of live R and heat killed S in to
the mice.
(2) The observation indicates that live S has killed the mice where as mice were
healthy with heat killed S or live R. Surprisingly, mice injected with mixture of
live R with heat killed S were found dead, and bacteria isolated from these
dead mice were virulent. Based on these observations, Griffith hypothesized
the existence of a transforming agent (Protein, DNA) being transferred from
heat killed virulent strain to the avirulent strain and proposed the concept of
transformation. Later, Oswald has proved that the transforming factor is DNA
rather than protein.
34. Mechanism of
Transformation
• Transformation is the
process by which cell free
DNA is taken up by another
bacteria.
• The DNA from donor
bacteria binds to the
competent recipient cell and
DNA enters into the cell.
• The DNA enters into the
recipient cell through a
uncharacterized
mechanism.
• The DNA is integrated into
the chromosomal DNA
through a homologous
recombination. Naturally
transformation is common
between closely related
35. Procedure for
Transformation
• Natural ability of a bacterium to take up cell free DNA present in
extracellular environment is low and only 1% bacteria are capable to take
DNA.
• Hence, a number of (chemical/physical) treatments make the cell
competent to take DNA through transformation.
• A list of agent used for different organism is given in Table 20.1.
The most popular reagent for making E.coli competent cell is calcium
chloride
36. The complete procedure of
transformation
The complete procedure of transformation is given in Figure 20.3 and it has following
steps:
1.Bacterial Culture- The growth stage of the bacteria has a significant impact for its
ability to take up foreign DNA. The bacterium at log phase is more active and efficient
to perform DNA damage and repair than stationary phase. As a result, it is preferred to
use a bacteria of log phase for making competent cells for transformation.
2.Preparation of Competent Cell-Bacteria is incubated with divalent cation (Calcium
chloride,Manganese chrloride or Rubidium chloride) for 30mins at 400
C. During this
process, cell wall of treated bacteria swell and gathers factors required for intake of
DNA docked on the plasma membrane.
• As with many breakthroughs in recombinant DNA technology, the key development as
far as transformation is concerned occurred in the early 1970s, when it was observed
that E. coli cells that had been soaked in an ice cold salt solution were more efficient at
DNA uptake than unsoaked cells. A solution of 50-100 mM calcium chloride (CaCl2) is
traditionally used, although other salts, notably rubidium chloride, are also effective.
• Exactly why this treatment works is not understood. Possibly CaCl2
causes the DNA to
precipitate onto the outside of the cells, or perhaps the salt is responsible for some
kind of change in the cell wall that improves DNA binding. In any case, soaking in CaCl2
affects only DNA binding, and not the actual uptake into the cell. When DNA is added
to treated cells, it remains attached to the cell exterior, and is not at this stage
transported into the cytoplasm (Figure 5.3). The actual movement of DNA into
competent cells is stimulated by briefly raising the temperature to 42°C. Once again, the
exact reason why this heat shock is effective is not understood.
37. 3.On the day of transformation, competent cells
are incubated with DNA or circular plasmid
containing appropriate resistance gene such as
ampicillin resistance gene for 30mins on ice.
4.Heat Shock -Competent cells are given a brief heat
shock (420
C for 60 -90 sec) to relax the cell wall
and high temperature stress causes upregulation of
the factor responsible for DNA recombination and
repair.
5.A chilled media is added for faster recovery of
transformed cells and grown for 1-1.5 hours
6.it is plated on the solid media with appropriate
antibiotics such as ampicillin and allowed to
grow for another 12-18 hrs
7.Transformed cells with appropriate resistance will
grow and give colony.
39. TRANSFORMATION IN YEAST
1. Lithium Acetate/ssDNA/PEG Method:
• In this method, yeast cells are incubated with a
transformation mixture of lithium acetate, PEG 3500,
single stranded carrier DNA and foreign plasmid at
420
C for 40 mins.
• The purpose of adding carrier DNA is to block the non-
specific sites on cell wall and make plasmid available
for uptake.
• Post- transformation, cells are pelleted to remove
transformation mixture and re-suspended in 1ml
water.
• It is plated on a solid media with an appropriate
40. 2. Spheroplast Transformation Method:
In this method, yeast cell wall is removed partially to produce
spheroplast.
Spheroplasts are very fragile for osmotic shock but are competent to
takes up free DNA at high rate. In addition, polyethyl glycol (PEG) is
used to facilitate deposition of plasmid and carrier DNA on cell wall
for easier uptake.
The mechanism of DNA uptake in yeast is not very clear. A schematic
of spheroplast method is given in Figure 20.4.
(1) In the spheroplast method, yeast cells are incubated with
zymolyase to partially remove cell wall to produce spheroplast.
(2)They are collected by centrifugation and incubated with carrier DNA
and plasmid DNA for 10mins at room temperature.
(3)It is now treated with PEG and calcium for 10mins with gentle
shaking.
(4)Transformed spheroplast are plated on selective solid media and
0
42. 3. Electroporation
• Plasma membrane is composed of lipid and protein. These
macromolecules give a partial conductance to the cell
membrane.
• When a high electric pulse is given to the cell, the charge
run across the membrane and partially disturbs the
arrangement of lipid molecule.
• As a result, it makes formation of pore and allow easy
passage of macromolecule especially charged molecule
like DNA into the cell.
• After the electroporation, cell is allowed to recover from
the damage and it forms colony on the selective solid
43. DNA Delivery in mammalian cells
Mammalian cell membrane surface chemistry, intracellular compartmentation and uptake mechanism
is different from the prokaryotic cells or yeast. Hence specialized methods have been developed to suit
mammalian cells.
There are many strategies to deliver the DNA in mammalian cells:
1. Chemical transfection techniques-
The principle behind the chemical transfection technique is to coat or complex the DNA with a polymeric
compound to a reasonable size precipitate (Figure 21.1).
It facilitates the interaction of the precipitate with the plasma membrane and uptake through endocytosis.
There are multiple chemical compounds have been discovered which can be able to make complex and
deliver DNA into the mammalian cell.
44. DNA Delivery in mammalian cells
1A : Calcium Phosphate method
In this method, DNA is mixed with calcium chloride in phosphate buffer and
incubated for 20mins.
Afterwards, transfection mixture is added to the plate in dropwise fashion.
DNA-calcium phosphate complex forms a precipitate and deposit on the cells
as a uniform layer.
The particulate matter is taken up by endocytosis into the internal storage of
the cell. The DNA then escapes from the precipitate and reach to nucleus
through a unknown mechanism.
This method suited to the cell growing in monolayer or in suspension but not
for cells growing in clumps.
But the technique is inconsistent and the successful transfection depends on
DNA-phosphate complex particle size and which is very difficult to control.
45. 1B. Polyplexes method
• The disadvantage of calcium phosphate method is severe physical damage to the cellular
integrity due to particulate matter settling on the cell.
• It results in reduced cellular viability and cyto-toxicity to the cell. An alternate method was
evolved where DNA was complexed with chemical agent to form soluble precipitate
(polyplexes) through electrostatic interaction with DNA (Figure 21.2).
• A number of polycationic carbohydrate (DEAE-Dextran), positively charged cationic lipids
(transfectin), polyamines (polyethylenimines) etc are used for this purpose.
• The soluble aggregates of DNA with polycationic complex is readily been taken up by the
cell and it reaches to the nucleus for expression
46. DNA Delivery in mammalian cells
1C. Liposome and lipoplex method
• Another approach of DNA transfection in animal cell is to pack the DNA
in a lipid vesicle or liposome. In this approach, DNA containing vesicle will
be fused with the cell membrane and deliver the DNA to the target cell.
• Preparation of liposome and encapsulating DNA was a crucial step to
achieve good transfection efficiency.
• Liposome prepared with the cationic or neutral lipid facilitates DNA
binding to form complex (lipoplex) and allow uptake of these complexes
by endocytosis.
• The lipoplex method was applicable to a wide variety of cells, and found
to transfect large size DNA as well.
• Another advantage of liposome/lipoplexes is that with the addition of
ligand in the lipid bilayer, it can be used to target specific organ in the
47. DNA Delivery in mammalian cells
Bactofection
This mode of gene transfer is very popular in plant where
agrobacterium tumefaciens is used.
In animal cell, bacteria is actively been taken up by the host cell through
phagocytosis and entrapped in a membranous vesicle known as phagosome.
Then bacteria escapes from phagosome and get lysed to release the DNA into
the cytosol.
In alternate mechanism, bacteria get lysed inside the phagosome and DNA is
released into the cytosol.
The bacterial species used in this methods are salmonella, shigella etc. Most of
the strain used to deliver the DNA are attenuated so that they should not harm
host cell
48. DNA Delivery in mammalian cells
Transduction (Virus mediated)-
Viral particle has a natural tendency to attack and deliver the DNA into the
eukaryotic cells.
As discussed previously, cloning gene of interest in to the viral vectors is a
innovative way to deliver the DNA into the host cell.
If the recombination sequences are available, the delivered DNA is integrated into
the host and replicate.
Virus has essential components for expression of proteins required for DNA
replication, RNA polymerase and other ligand for attachment onto the host cell. In
addition, it has additional structural components to regulate infection cycle.
If the virus vector contains cassettes to perform all these functions then it is fully
sufficient to propagate independently. Few virus strains may cause disease if their
propagation will be uncontrolled. A mechanism has been devised to keep a check
on the uncontrolled propagation of virus in cell.
49. 3. Through Viral vectors:
Particular viruses have been selected as gene delivery vehicles
because of their capacities to carry foreign genes and their ability
to efficiently deliver these genes associated with efficient gene
expression.
Principle:
> Generation of recombinant virus containing the transgene via gene
cloning.
> Amplification of viral particles in packaging cell line and virus isolation.
> Purification and titration of viral particles
> Infection of cell type of interest.
> Depending on the virus, integration of transgene in host genome.
Package the DNA inside an animal virus, since viruses have evolved
mechanisms to naturally infect cells and introduce their own nucleic acid. The
transfer of foreign DNA into a cell by this route is termed transduction.
Whichever route is chosen, the result is transformation, i.e. a change of the
recipient cell’s genotype by integration of the transgene. Transformation can
be transient or stable, depending on how long the foreign DNA persists in the
cell.
50. The first step in the infection cycle of a virus is the interaction
between the virus and a cellular receptor on the surface of a target
cell, resulting in the fusion of the viral and the cellular membrane.
Cells which are not carrying such a receptor cannot be infected by
the virus.
Limitations of viral gene transfer are the time-consuming and
laborious production of vectors, elevated laboratory costs due to
the high level of safety requirements, limitation of insert size (~ 10
kb for most viral vectors versus ~ 100 kb for non-viral vectors).
Variability in infection potencies of the generated virus particle
preparations and possible immunogenic reaction in animal or
clinical trials.
51. Alternatively, using different recombinant viruses each expressing a
different protein, a co-infection of the desired cell lines has been
performed. However, this is very time consuming and laborious.
Well-known examples for viral gene transfer vectors are
recombinant adenoviruses, retroviruses, adenoassociated viruses,
herpes simplex viruses and vaccinia viruses.
Adenoviruses have a broad cell tropism and can infect both dividing
cells and non dividing cells. The packaging capacity of adenoviral
vectors is 7 to 8 kb.
Retroviruses make excellent gene therapy vectors because they have
the ability to integrate their genome into a host cell genome thus
enabling stable expression of the transgene. However, most
retroviruses are limited by the requirement of replicating
cells for infection.
52. An exception are the lentiviruses (subgroup of retroviruses), which
have the ability to infect and integrate into non dividing cells.
Retroviruses can carry foreign genes of around 8 kb. Among the
disadvantages are the instability of some retroviral vectors, possible
insertional mutagenesis by random integration into the host DNA.
Adenoassociated viruses need helper viruses like adenovirus or
herpes virus for lytic infection.
Furthermore the adenoassociated viruses have only limited
capacity for insertion of foreign genes ranging up to 4.9 kb. Wild
type viruses have the ability to integrate into a specific region
of the human chromosome thus avoiding insertional mutagenesis.
An advantage is the low immunogenicity of adenoassociated viruses
which is important for the application in human gene therapy.
53. Besides retroviruses, adenoviruses and adenoassociated viruses
herpes simplex viruses and vaccinia viruses are frequently used for
viral gene transfer. They have the ability to carry large inserts up to
50 kb.
Herpes simplex viruses have been used for gene transfer into
neurons, brain tumors, various tumor cells and B cells.
54. Screening of recombinant clone
Chromogenic Substrate-
The use of chromogenic substrate to detect a particular enzymatic activity is the basis
to screen the desired clone. The most popular system to exploit this feature is “Blue
white screening” where a coloroless substrate is processed to a colored compound.
The colorless compound X-gal or 5-bromo-4-chloro-3-indolyl-β- D-galactoside used
in this screening method is a substrate for β-galactosidase (Figure 22.1).
The enzyme β-galactosidase is the product of lacZ gene of the lac operon. It is a
tetrameric protein and an initial N-terminal region (11-41) of the protein is important
for activity of the protein.
In this system, host contains lacZ gene without the initial region where as vector
contains α-peptide to complement the defect to form active enzyme. As a result, if a
vector containing α-peptide will be transformed into the host containing remaining
lacz, the two fragment will reconstitute to form active enzyme. In addition, the α-
peptide region in vector contains multiple cloning site and as a result of insertion of
gene fragment, consequently α-peptide will not be synthesized to give fully active β-
galactosidase. The enzyme β-galactosidase oxidizes x-gal to form 5-bromo-4-chloro-
indoxyl and galactose. The indoxyl derivative is oxidized in air to give a blue colored
dibromo-dichloro derivative (Figure 22.2). Hence, blue colored colonies indicate the
presence of an active enzyme or absence of insert where as colorless colonies
indicate presence of an insert.
56. Insertional inactivation
In this approach a foreign DNA is cloned within the coding gene
responsible for a phenotype. As a result of insertion, the gene
product is not available to modulate the phenotype of the host.
This approach is known as insertional inactivation, and it can be
used with a suitable genetic system.
(1) Insertional Inactivation of antibiotic resistance gene- As
discussed in an earlier lecture, bacterial plasmid PBR322 has
two antibiotic resistance gene, Apr
and Tcr.
If a gene fragment
will be cloned in ScaI, it will disrupt the Apr
gene. As a result,
the clone will be ampicillin sensitive and Tcr,
where as the
original plasmid will be Apr
and Tcr
.
To select the clone, first the transformed e.coli is plated on
tetracycline containing media. Subsequently, a replica plate will be
made on ampicillin containing media to identify the clone growing
57. Figure 22.3: Insertional Inactivation of antibiotic resistance gene in pBR322 to screen
recombinant clone.
58. Insertional
inactivation
(2) Insertional Inactivation of LacZ gene- LacZ is a
part of lac operon and responsible for synthesis of β-
galactosidase. As discussed earlier, X-gal system can
be used to detect the insertional inactivation of LacZ
gene to screen the cloned fragment. If the gene is
inserted into the lacz, the clone will not be able to
produce a functional β-galactosidase. Hence, blue
colored colonies indicate the presence of an active
enzyme or absence of insert where as colorless
colonies indicate presence of an insert.
59. Insertional
inactivation
(3) Insertional Inactivation of cI gene-
During an infection cycle, virus undergoes a lytic and lysogenic stages.
The lytic phase is responsible for lysis of host to release the virus
particle where as lysogenic stage allow the replication of virus without
lysis of host.
The cI gene encodes for cI repressor and which is responsible for the
formation of lysogens.
In the presence of functional cI, the plaques contains unlysed host cells
and has a turbid appearance where as in the absence it will clear.
This feature can be use to screen the clone to detect functional cI
(absence of clone) or absence of cI (presence of insert). This
approach is schematically depicted in Figure 22.5.
60.
61. Screening of recombinant clone
Antibiotic sensitivity-
Vector carries a functional selection marker such as antibiotic resistance gene
to be use to select the clone.
The antibiotic resistance gene product has multiple mechanism to provide
resistance in host cell (Table 22.1).
In this approach, a circular plasmid containing antibiotic resistance can be
able to replicate into the host cell plated on a antibiotic containing media. In
the cloning of a fragment into this plasmid, the plasmid is cut with restriction
enzymes and a fragment in ligated to give circular plasmid with insert.
The transformation of both DNA species; cut plasmid and circularized clone
into the host and plated onto the antibiotic containing solid media.
Only circularized clone will give colonies where as cut plasmid will not grow as
it has lost antibiotic resistance gene.
62. TABLE 22.1: ANTIBIOTICS RESISTANCE GENE AND THEIR MODE OF
MECHANISM.
Antibiotic Gene product Mechanism
Ampicillin β-lactamase Degradation of ampicillin
Kanamycin Neomycin
phosphotransferase II
Covalent modification of
kanamycin
Tetracycline Ribosomal
protection proteins
Efflux of tetracycline
outside of the bacteria
Chloramphenicol Chloramphenicol
acetyl transferase
Chloramphenicol to
acetyl Chloramphenicol
63. Complementation of
mutation
In this approach, a mutant host strain can be used to screen the
plasmid containing the missing gene and the transformant will
grow only if the gene product from clone will complement the
function. In general genes taking part in metabolic pathway or
biosynthetic pathway are routinely been used for this purpose.
There are 3 important requirement for this approach to work-
1.Host strain deficient in a particular gene. If the gene belongs to
the biosynthetic pathway, the mutant host in this case are called as
auxotroph as host dependent on the gene product or final product
of the biosynthetic pathway as a supplement in media for growth.
2. A defined media with missing nutrient.
3.A vector containing the gene to supply the gene product to
64. Selectable
marker
• A selectable marker protects the organism from
a selective agent that would normally kill it or prevent
its growth. In a transformation reaction, depending
on the transformation efficiency, only one in a several
million to billion cells may take up DNA. Rather than
checking every single cell, scientists use a selective
agent to kill all cells that do not contain the foreign
DNA, leaving only the desired ones.
• Antibiotics are the most common selective agents.
In bacteria, antibiotics are used almost exclusively.
In plants, antibiotics that kill the chloroplast are
often used as well, although tolerance to salts and
growth- inhibiting hormones is becoming more
popular. In mammals, resistance to antibiotics that
would kill the mitochondria is used as a selectable
marker.
65. Selectable
marker
• For molecular biology research different types of markers may be
used based on the selection sought. These include:
• Positive or selection markers are selectable markers that confer
selective advantage to the host organism.[1]
An example would be
antibiotic resistance, which allows the host organism to survive
antibiotic selection.
• Negative or counterselectable markers are selectable markers that
eliminate or inhibit growth of the host organism upon
selection.[2]
An example would be thymidine kinase, which makes
the host sensitive to ganciclovir selection.
• Positive and negative selectable markers can serve as both a
positive and a negative marker by conferring an advantage to the
host under one condition, but inhibits growth under a different
condition. An example would be an enzyme that can
complement an auxotrophy (positive selection) and be able to
convert a chemical to a toxic compound (negative selection).
66. Selectable
marker
Examples of selectable markers include:
• Beta-lactamase which confers ampicillin resistance to
bacterial hosts.
• Neo gene from Tn5, which confers resistance to kanamycin in
bacteria and geneticin in eukaryotic cells[3]
• Mutant FabI gene (mFabI) from E. coli genome, which
confers triclosan resistance to the host.[4]
• URA3, an orotidine-5' phosphate decarboxylase from yeast is a
positive and negative selectable marker. It is required for uracil
biosynthesis and can complement ura3 mutants that are
auxotrophic for uracil (positive selection). The enzyme URA3 also
converts 5-fluoroorotic acid (5FOA) into the toxic compound 5-
fluorouracil, so any cells carrying the URA3 gene will be killed in the
presence of 5FOA (negative selection)
67. Reporter gene
Genetic reporters are used as indicators to study gene expression and cellular
events coupled to gene expression. They are widely used in pharmaceutical and
biomedical research and also in molecular biology and biochemistry. Typically, a
reporter gene is cloned with a DNA sequence of interest into an expression vector
that is then transferred into cells. Following transfer, the cells are assayed for the
presence of the reporter by directly measuring the reporter protein itself or the
enzymatic activity of the reporter protein. A good reporter gene can be identified
easily and measured quantitatively when it is expressed
General Characteristics of Ideal Reporter Gene:
Some genes are selected as reporters because the characteristics they confer on
organisms expressing them are easily assayed and measured or because they are
selectable markers.
They are also nontoxic, non- immunogenic and small enough to fit into a delivery
vehicle and biological half-life of the reporter protein is relevant for the studied
phenomenon.
Measurements that are almost instantaneous, exceptional sensitivity, a wide dynamic
range, typically no endogenous activity in host cells to interfere with quantitation.
68. Utilization of Reporter Gene: Normally the reporter gene is
linked to a promoter sequence through an expression vector
that is further transferred into cells. After transfer, the cells
are assayed for the presence of the reporter by directly
testing the amount of either the reporter mRNA, the reporter
proteins, or the enzymatic activities of the reporter proteins
Recently Used Reporter Gene in Diagnostic Assay: Reporter
gene assays continue to be one of the simplest, most robust
ways to analyze the activation of transcription factors and
their associated signaling pathways.
Normally used reporter genes that induce visually identifiable
features usually involve fluorescent and luminescent proteins.
These include the gene that encodes jellyfish green
fluorescent protein (GFP), which causes cells that express it to
glow green under blue light, the enzyme luciferase, which
catalyzes a reaction with luciferin to produce light
69. The Luciferase Reporter Assay
The luciferase reporter assay is commonly used as a tool to study
gene expression at the transcriptional level. It is widely used because
it is convenient, relatively inexpensive, and gives quantitative
measurements instantaneously.
Luciferases make up a class of oxidative enzymes found in several
species that enable the organisms that express them to
“bioluminesce,” or emit light. The most famous one of these enzymes
is the firefly luciferase. Fireflies are able to emit light via a chemical
reaction in which luciferin is converted to oxyluciferin by the
luciferase enzyme. Some of the energy released by this reaction is in
the form of light.
The assay setup is shown in Figure 23.4. The reporter gene
construct containing luciferase is transfected to the mammalian
cells. The cells are washed with PBS and lysed with a lysis buffer.
71. Chimeric Construct with green fluorescent
protein (GFP)
In the live cell, green fluorescent protein is a good choice as
reporter gene to screen cells containing recombinant protein
fluorescently tagged with the GFP at their c-terminus.
The cell receiving recombinant DNA will give green fluorescence and it
can be visualized with an inverted fluorescence microscope and it can
be analyzed in flow cytometer to separate the GFP containing cells
from the un-transfected cells.
Flow cytometer analyses the cell based on its shape, size and
fluorescence level. A non-fluorescent cell gives a separate peak as
compared to the fluorescently labelled cells and with the help of flow
cytometer, both of these peaks can be collected in separate tubes.
Besides, GFP, red fluorescent protein, yellow fluorescent protein, cyan
fluorescent protein are also popular to use to label the protein.
