Bth 303 genetic engineering

for Bangalore University M.Sc Biotechnology 3 rd sem students

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FOR MSC BIOTECHNOLOGY STUDENTS
2014 ONWARDS

Biochemistry scanner
THE IMPRINT
BTH – 303: GENETIC ENGINEERING
As per Bangalore University (CBCS) Syllabus
2016 Edition

BY: Prof. Balasubramanian Sathyamurthy

Supported By:
Ayesha Siddiqui
Kiran K.S.



THE MATERIALS FROM “THE IMPRINT (BIOCHEMISTRY SCANN ER)” ARE NOT
FOR COMMERCIAL OR BRAND BUILDING. HENCE ONLY ACADEM IC CONTENT
WILL BE PRESENT INSIDE. WE THANK ALL THE CONTRIBUTO RS FOR
ENCOURAGING THIS.
BE GOOD – DO GOOD & HELP OTHERS

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DEDICATIONDEDICATIONDEDICATIONDEDICATION

I dedicate thI dedicate thI dedicate thI dedicate this material to my spiritual guru Shri Raghavendra swamigal, is material to my spiritual guru Shri Raghavendra s wamigal, is material to my spiritual guru Shri Raghavendra s wamigal, is material to my spiritual guru Shri Raghavendra s wamigal,
parents, teachers, well wishers and students who always increase my morale parents, teachers, well wishers and students who always increase my morale parents, teachers, well wishers and students who always increase my morale parents, teachers, well wishers and students who always increase my morale
and confidence to share my and confidence to share my and confidence to share my and confidence to share my knowledge knowledge knowledge knowledge to to to to reach reach reach reach all beneficiariesall beneficiariesall beneficiariesall beneficiaries....

PREFACEPREFACEPREFACEPREFACE

Biochemistry scanner ‘THE IMPRINT’ consists of last ten years solved question
paper of Bangalore University keeping in mind the s yllabus and examination
pattern of the University. The content taken from t he reference books has been
presented in a simple language for better understan ding.

The Author Prof. Balasubramanian Sathyamurthy has 1 5 years of teaching
experience and has taught in 5 Indian Universities including Bangalore
University and more than 20 students has got univer sity ranking under his
guidance.
THE IMPRINT is a genuine effort by the students to help their peers with their
examinations with the strategy that has been succes sfully utilized by them.
These final year M.Sc students have proven their me ttle in university
examinations and are College / University rank hold ers.
This is truly
for the students, by the students. We thank all the contributors for
their valuable suggestion in bringing out this book . We hope this will be
appreciated by the students and teachers alike. Suggestions are welcomed.
For any comments, queries, and suggestions and to g et your free copy write us
at [email protected]
or call 9980494461

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CONTRIBUTORS:
CHETAN ABBUR ANJALI TIWARI
AASHITA SINHA ASHWINI BELLATTI
BHARATH K CHAITHRA
GADIPARTHI VAMSEEKRISHNA KALYAN BANERJEE
KAMALA KISHORE
KIRAN KIRAN H.R
KRUTHI PRABAKAR KRUPA S
LATHA M MAMATA
MADHU PRAKASHHA G D MANJUNATH .B.P
NAYAB RASOOL S NAVYA KUCHARLAPATI
NEHA SHARIFF DIVYA DUBEY
NOOR AYESHA M PAYAL BANERJEE
POONAM PANCHAL PRAVEEN
PRAKASH K J M PRADEEP.R
PURSHOTHAM PUPPALA DEEPTHI
RAGHUNATH REDDY V RAMYA S
RAVI RESHMA
RUBY SHA SALMA H.
SHWETHA B S SHILPI CHOUBEY
SOUMOUNDA DAS SURENDRA N
THUMMALA MANOJ UDAYASHRE. B
DEEPIKA SHARMA

EDITION : 2016
PRINT : Bangalore
CONTACT : [email protected] or 9980494461

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M. SC. BIOTECHNOLOGY – SECOND SEMESTER
BTH – 303: GENETIC ENGINEERING

4 units (52 hrs)
UNIT : 1 Introduction to Genetic Engineering 2 hrs
Scope and importance of genetic engineering
UNIT : 2 Tools of Genetic Engineering 14hrs
Enzymes, Non-specific endo and exo nuclease, DNase, RNase. Restriction modification;
restriction endonuclease- types, nomenclature, recongnition sequence and mechanism
of action. Methylation, RNA modification. Role of k inases, phosphatases,
polynuclcleotide phosphorylase, polynucleotide kinase ligases – types and mechanism
of action
VECTORS : General characteristics of vectors , brief account of naturally occurring
plasmids promoters, MCS, Ori and maker gene-lac Z. construction of pBR 322,
pBR325, pUC18 and 19 vectors and expression vectors E.coli promoters, lac promoters,
lac promoters, trp promoters, lambda pL promoters, hybrid tac promoters, ribosome
binding site, codon selection. M13 derived vectors, Lambda based vectors, cosmids,
phagemids, minichromosomes, BAC’s, YAC’s , shuttle vectors, Ti plasmids, vectors for
animals-SV40 and Bovine papilloma virus.
UNIT : 3 Gene Cloning Strategies and Construction o f Gene Libraries 14hrs
Cloning from mRNA, isolation and purification of RNA, synthesis of cDNA, Isolation of
plasmids, cloning cDNA in plasmid vectors, cloning cDNA in bactriaophage vectors.
cDNA library.
Cloning of genomic DNA: Isolation and purification of DNA, preparation of DNA,
preparation of DNA, fragments and cloning. Constriction of genomic libraries (Using
lambda gt 10 and 11 vectors ) In vitro packaging of lambda phage and amplification of
libraries
Advanced cloning strategies synthesis and cloning of cDNA, PCR amplified DNA, use of
adaptors and linkers, homopolymers tailing in cDNA cloning, expression of cloned DNA
molecule.
Selection, screening and analysis of recombinant
Genetic selection, insertional inactivation, chromogenic substances, complementation
of defined mutations, nucleic acid hybridization, screening methods for cloned libraries,

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PCR screening protocols, immunological screening, restriction mapping of cloned
genes, blotting techniques, sequencing methods. Purification strategies of expressed
His-tagged proteins.
UNIT : 4 Transformation Techniques 8hrs
Purification of vectors DNA, restriction digestion, end modification, cloning of foreign
genes ( from mRNA, genomic DNA ) transformation screening, selection, expression and
preservation. Transformation and transfection techniques, preparation of competent
cells of bacteria, chemical methods-calcium phosphate precipitation methods, liposome
mediated method, physical methods-Electroporation, gene gun method. Method of DNA
transfer to yeast, mammalian and plant cells, trans formation and transfection
efficiency.
UNIT : 5 Labelling and Detection Techniques 8hr s
Labeling of DNA, RNA and proteins by radioactive isotopes, non-radioactive labelling, in
vivo labeling, autoradiography and autofluorography. DNA sequencing by enzymatic
and chemical methods, Agarose gel electrophoresis, PAGE, PFGE. Methods of nucleic
acid hybridization; southern, northern and western blotting techniques.
UNIT : 6 Chemical Synthesis Of Genes And PCR 6h rs
Phosphodiester, phosphotriester and phophite ester methods, principles and strategies.
Oligonucleotide synthesis and application, synthesisof complete gene.
PCR, methodology, essential feature of PCR, primers , Taq polymerases, reverse
transcriptase-PCR, types of PCR-Nested, inverse, RAPD-PCR, RT-PCR (real time PCR),
Application of PCR.

References:
1.
Nicholl D.S.T. Introduction to Genetic Engineering Cambridge (3
rd
Ed.) University
press.UK. 2008
2.
Old R.W., Primrose S.B. Principles of gene manipulation - An introduction to genetic
engineering (5
th
Ed.), Blackwell Scientific Publications, UK. 1996.
3.
David S L. Genetics to Gene Therapy – the molecular pathology of human disease(1
st

Ed.) BIOS scientific publishers, 1994.

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4. Ernst-L Winnacker, From Genes to Clones: Introduction to Gene Technology. WILEY-
VCH Verlag GmbH, Weinheim, Germany Reprinted by Pan ima Publishing Corporation,
New Delhi. 2003
5.
Benjamin Lewis, Genes VIII (3
rd
Ed.) Oxford University & Cell Press,NY.2004
6.
Robert Williamson.Genetic Engineering (1
st
Ed.) Academic Press.1981.USA
7.
Rodriguez. R.L (Author), Denhardt D.T. Vectors: A Survey of Molecular Cloning Vectors
and Their Uses (1
st
Ed.) Butterworth-Heinemann publisher.UK. 1987
8.
Ansubel F.M., Brent R., Kingston R.E., Moore D.D. et al. Short protocols in molecular
biology(4
th
Ed), Wiley publishers. India. 1999.
9.
Sambrook J et al. Molecular cloning Volumes I, II and III. Cold Spring Harbor laboratory
Press, New York, USA. (1989, 2000)
10.
Terence A Brown. Genomes, (2
nd Ed.) BioScientific Publishers.UK.2002
11.
Anthony JF Griffiths, William M Gelbart, Jeffrey H Miller, and Richard CLewontin
Modern Genetic Analysis (1
st
Ed.)W. H. Freeman Publishers.NY. 1999
12.
S. B. Primrose, Richard M. Twyman.Principles of gene manipulation and genomics (7
th

Ed.) John Wiley & Sons publishers.2006

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UNIT: 1 Introduction to Genetic Engineering
Scope and importance of genetic engineering

SCOPE AND IMPORTANCE OF GENETIC ENGINEERING
Introduction
Over the past 12,000 years humans have gradually de veloped greater understanding
and control over life. Agriculture, including plant and animal husbandry, were early
important developments. Medicine also contributed to the control of life by fighting
disease and more recently through technologies to control and manipulate fertility.
Knowledge and technologies from physics and chemistry provide the tools to investigate
biological processes at a molecular and even atomic level. Late 20th century and 21st
century genetic science heralds remarkable advances in our understanding of life and
our ability to control and manipulate it for our teleological endeavours. Emerging
biotechnologies are in the foreground of modern sci entific research. Evolutionary
theory, Mendel’s laws of inheritance, the discovery of DNA, the mapping of the human
genome, genetic engineering (GE) of organisms, gene therapy, synthetic biology, cloning,
stem cell therapies, epigenetics, and life extension research are theories and
technologies providing powerful new insights into the nature of life and the development
of technologies to manipulate all aspects of life. This knowledge is deconstructing and
reconstructing our knowledge of what life is and what it means to be human, and where
humans sit in the order of nature. Table 1 lists a brief selection of important milestones
in humanity’s understanding and control of life along with some loosely associated
worldviews.
Genetic technology has the potential to change biol ogical and social reality. Its
development and application have consequences for h umans, other animals and the
planetary biosphere. These consequences are open to moral evaluation, questions that
may be asked include:
What are the likely social and moral impacts? Is this progress? Are these consequences
good or bad? Does the potential good outweigh the potential bad? For whom? How fair
are theconsequences? How easily can they be accesse d or avoided? And how do
different social and biophysical contexts affect their moral status? Another relevant
question is, can the positive consequences obtained by use of genetic technology be

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obtained using alternative technologies (perhaps wi th less potential for negative
consequences)? These questions demonstrate that the practice of genetic science (and
indeed science in general) is inextricably bound to moral reasoning, moral behaviour
and technological foresighting.
The rise of genetic science
Darwin’s Theory of Evolution completely revised our notions of the nature of life and its
origins. Species were no longer created individually by God, nor once ‘created’, were
they fixed and immutable. No longer were we a unique and special creature, made in
the image of a miraculous supernatural creator, rat her, it became apparent that
humans were one of approximately ten millions species inhabiting earth, evolving to fit
selection pressures in a similar fashion to the other animals on the planet. Gregor
Mendel’s laws of inheritance statistically demonstrated that characteristics could be
passed on from one generation to the next. The discovery, in the early twentieth century
by Thomas Hunt Morgan, of chromosomes and the genet ic diversity engendered by
sexual reproduction, and the mid century discovery of DNA by Crick and Watson
provided a causal mechanism for inheritance and a m olecular level mechanism for
Darwinian natural selection.
Biological Milestones Year Associated worldview

Agriculture – plant and animal
Husbandry
10,000BC Animistic/magical/mythological

Ancient medicine (e.g., Imhotep,
Hippocrates, Galen)
2500BC –
180AD
Animistic/magical/mythological
/religious/Ptolemaic
Medieval medicine (e.g.,Avicenna,
Ibn an-Nafis, Paracelsus)
1000-
1500AD
Religious/Ptolemaic

Renaissance medicine (e.g.,
Vesalius to Jenner)
1500-
1800
Religious/Copernican/scientific

Darwin’s Theory of Evolutionary 1860
Mendel’s Laws of inheritance 1865
Pasteur invents vaccines 1880
Morgan’ discovery of the 1915

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chromosomes
Fleming invents antibiotics 1928 Religious/Copernican/scientific/
modernist Watson and Crick discover DNA 1953
Fertility control – oral
contraceptive, in vitro fertilisation
1960
Genetic engineering 1971
Tissue engineering 1987 Copernican/scientific/post
modernist Gene therapy (1970)
1990
Epigentics 1990
Animal cloning 1996
Stem cells therapy 1998
Life extension 2000
Synthetic biology 2000
Technology has enabled the genomes of organisms to be ‘read’ and compared, showing
that humans share more than 98% of our genes in com mon with the chimpanzee
(Jones, 2006), giving us new insights into our biological and moral position within
nature.
The Human Genome Project (HGP) achieved three major goals. First, it sequenced the
order of all the 2.9 billion base pairs in the genome. Second, it developed maps locating
genes for major section of all our chromosomes. Thi rd, it produced ‘linkage maps’
enabling inherited traits to be tracked over generations. Francis Collins, the director of
the HGP described the results and meaning of the HGP as:
It's a history book - a narrative of the journey of our species through time. It's a shop
manual, with an incredibly detailed blueprint for building every human cell. And it's a
transformative textbook of medicine, with insights that will give health care providers
immense new powers to treat, prevent and cure disea se. (Cited by National Human
Genome Research Institute, 2009).
As the relationship between genes and individual health and behaviour becomes more
apparent, moral questions arise as to who may have access to an individual’s genome,
and what will they be able to do with this information. As significant a milestone as it
is, sequencing of the genome merely marks a beginning. It will take many decades (and

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massive computer power) to understand how the appro ximately 20,000 genes in the
human genome interact with one another to produce o ver two hundred thousand
different proteins.
A great deal is not currently understood about how the genome works. Long held
theories continue to be questioned. For example, contrary to the last hundred years of
scientific belief, Mendel’s Laws have recently been challenged. Although still believed to
be fundamentally correct, it has been claimed that Mendel’s Laws are not absolute and
exceptions occur (Lolle, Victor, Young, & Pruitt, 2005). Likewise, the idea of inherited
acquired characteristics was for a long time considered biological and scientific heresy,
but the received scientific dogma has been challenged by the new science of epigenetics
(Jablonka & Raz, 2009; Kaati, Bygren, & Edvinsson, 2002; Lumey, 1992). Similarly, a
dozen years ago, with perhaps a little scientific a rrogance, molecular biologists
designated long stretches of organisms’ genomes as “junk DNA” claiming that these
non-coding segments served no purpose. However, it is logically obvious that human
lack of knowledge about the function of elements of nature does not mean they lack
function.
Recently, research has shown important roles for junk DNA (Nowacki, et al., 2009),
demonstrating the hubris of the junk DNA assumption . Indeed, it now appears that
junk DNA plays a vital role in evolution (in particular enabling fast genetic adaptation to
changing environmental circumstances) and will be c rucial for the refining of GE
techniques and for gene therapy (Feng, Naiman, & Co oper, 2009; Vinces, Legendre,
Caldara, Hagihara, & Verstrepen, 2009). New evidenc e also suggests that the rDNA
repeats known as “junk DNA” are essential for repairing the DNA damage caused by
factors such as UV light (Ide, Miyazaki, Maki, & Ko bayashi, 2010). The use of
technologies with powerful potential to affect the physical and social worlds, without a
good understanding of the science involved, has the potential for unexpected and
unforeseen negative social and moral impacts.
Developments in genetic science and moral questions
Genetic engineering
The breeding of promising individuals over generations in order to create desirable
phenotypic characteristics in plants and animals ha s long been practiced in
horticulture and animal husbandry. This is a relatively slow process with progressive

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changes made over many generations, not by nature or natural selection, but by human
intervention in the evolutionary progress of the species. Racehorses, domestic cattle,
show dogs and the staple grains are prime examples of centuries and even millennia of
breeding to slowly bend nature to the aesthetic tas tes and teleological desires of
humans.
In the past forty years, with the discovery of recombinant DNA, humans have gained
the power to make changes to an organism’s genome i n a single generation. Genetic
engineering (GE) involves the chemical addition or deletion of a specific gene from an
organism’s genome in order to bring about a desired change in the organism’s
phenotype. With this process organisms can have current characteristics enhanced or
removed and even entirely new characteristics, not evident in the organism’s species,
added. Thus, a gene from one species (or a synthetic analogue of the gene), may be
spliced into the genome of the same or a different species, or even an organism from a
different biological kingdom, giving the new GE organism phenotypic characteristics
from the donor species (Small, 2004a).
In this way GE can create organisms with desired attributes much more quickly than
traditional breeding (i.e., in a single generation). This amounts to a speeding up of
evolution in a direction decided by humans. This also differs from normal evolution and
animal and plant husbandry in that the new organism does not co-evolve, in little steps,
over time with the other organisms in its environment. Instead an evolutionary leap is
engineered within a single generation. Another difference between GE and selective
breeding is that organisms can be created that could not possibly have come about
naturally, as organisms generally cannot breed with others from different species or
kingdoms. Proponents see great hope for the common good of humanity in GE
technology, and often claim that the technology will be necessary to produce enough
food to feed the future population (Borlaug, 1997; Fedoroff, et al., 2010; Ortiz, 1998).
While GE offers the potential to further bend natur e to our desires, critical
commentators express concern about negative extrinsic moral impacts. These include
the potential to develop dangerous organisms, the impossibility of reversibility once
such organisms are loose in the environment, and the potential for negative impacts on
humans, other animals and the environment (Antoniou , 1996; Fox, 1999; Ho, 2000;
Rifkin, 1998; Straughan, 1995b). Others criticise the technology from an intrinsic moral

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perspective; creating life is the province of ‘God’ or nature – human attempts to usurp
the role of God or nature are seen as acts of hubris – against God or disrespectful to
nature (Appleby, 1999; Straughan, 1995a).
Currently GE is being used to engineer micro-organisms and bacteria (particularly for
the production of medicines such as insulin, factor 9 clotting agent, human growth
hormone, etc.), plants and animals for food product ion, production of medicines,
industrial production and phytoremediation. An example of a potential GE food animal
is the ‘ecofriendly’ GE pig, engineered to contain bacteria which help pigs remove
phosphate from their food, thus stopping it from passing through into the environment,
where it causes harm to life in streams and rivers (Golovan, et al., 2001). Pigs have also
been genetically engineered to contain human genes, so that their organs will be less
susceptible to immune system rejection when used fo r xenotransplantation (White,
Langford, Cozzi, & Young, 1995); the replacement of failing human organs with those
from animals.
Advocates of GE claim that the technology is safe. In 2008 GE crops were grown on 300
million acres worldwide. GE crops have been consumed for over 13 years without any
incident, it is claimed. Furthermore, production has increased and so have farmers’
profits, while pesticide and herbicide use have been reduced and the use of the no-till
method of agriculture (helpful for reducing soil erosion) increased (Fedoroff, et al.,
2010). However, so far the principal use of GE in food crops has been to engineer insect
resistance (bt crops) or to make the crops resistant to a specific herbicide used to
eliminate weeds from fields of growing crops – a major beneficiary being the company
selling the proprietary herbicide and seeds (one and the same company – Monsanto).
On the positive side, the herbicide for which resistance is engineered (Roundup or
glyphosate) is relatively environmentally benign and the whole process eliminates the
need for further applications of less environmentally benign herbicides.
One possibility presented by GE is the enhancement of nutritional qualities of crops, as
for example, the much heralded golden rice. Golden rice has been engineered to contain
extra beta-carotene which converts to vitamin A when consumed by humans. Many
people in developing countries, where rice is the primary staple, suffer from vitamin A
deficiency (Tang, Qin, Dolnikowski, Russell, & Grusak, 2009). Foods with genetically
enhanced health qualities or with healthy additives are referred to as functional foods

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and the science of developing them and studying the relationship between food plant
genes, health and the individual human genome is called nutrigenomics. Of course, the
societal benefits of functional foods will be dependent upon the public’s acceptance of
GE food.
Genetic engineering for medical purposes is considerably more acceptable to the general
public than GE of food crops (Small, Parminter, & Fisher, 2005). Proponents hope that
numerous medicines will be able to be grown in GE p lants and/or GE animals and
produced more cheaply than through current techniqu es. A biotech company,
SemBioSys, has submitted an Investigational New Dru g application for safflower-
produced recombinant human insulin to the U.S. FDA (SemBioSys, 2008). Edible
vaccines (e.g., potatoes, tomatoes, bananas etc) are being developed for a range of
diseases (e.g., cholera, measles, malaria, hepatitis B, type 1 diabetes etc) and are
proposed as a logistically simpler resolution of the problem of getting vaccines to those
in need in developing countries (Chowdhury & Bagasra, 2007; Levi, 2000). However, it
remains unclear how vaccine dosages would be controlled and how accepting the public
will be of the conflation of food and medicine. Nonetheless, biotech and pharmaceutical
companies have high hopes for rich profit streams from genetically enhanced medical
foods and functional foods.
GE animals have been used as ‘bioreactors’ to produ ce medicines and industrial
products. Cows, sheep and goats have been genetically engineered to produce human
proteins in their milk for medical purposes (Wells, 2010). Silk worms have been
genetically engineered to produce a form of the human protein collagen which scientists
hope to harvest for applications such as artificial skin and wound dressings (Tomita, et
al., 2003).
The industrial sector also contains many potential applications for GE technology in
terms of new methods of producing currently available materials, new materials with
desirable qualities, and the production of chemicals and biofuels. For example, spider
silk is stronger than steel and as resilient as kevlar, but it is very expensive to produce.
Scientist have placed artificial versions of silk genes in various plants (potatoes,
tobacco) and animals (goats) and, using this technology, hope to be able to mass
produce silk protein for the development of new bio degradable ‘super-materials’
(Scheller, Guhrs, Grosse, & Conrad, 2001).

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Gene engineered viruses have even been used to manu facture a ‘green battery’ which
the authors claim is capable of powering an iPod three times as long as current iPod
batteries (Lee, et al., 2009). However, some GE ani mals seem largely for human
entertainment, for example, the first GE pet commercially available in the U.S. was a
florescent red zebrafish called a GloFish (GloFish.com, 2010). A company called
Lifestyle Pets has marketed a genetically engineered hypoallergenic cat. Given the
history of animal breeding for traits of interest to humans, further such applications
seem highly probable. Indeed, GE pets suggest mythological sized possibilities; anyone
for a pet gryphon? Chimeras are indeed possible using genetic technologies, with a
number of research projects having already created them (however, a gryphon might be
a bit of a stretch). Of particular concern to some is the possibility of human-animal
chimeras (Robert & Baylis, 2003). Robert and Baylis imagine a fusion between a chimp
and a human. They suggest that there might be confu sion over the status of such a
creature and that it might lead to social disorder. However, Savulescu (2003) argues
that there might be good reasons to create human ch imeras. He suggests medical
reasons (e.g., to confer resistance to specific diseases such as AIDS), to delay aging, or
to enhance human capabilities.
Clearly, a range of ethical questions are opened by the creation of chimeras.
Undoubtedly, there will be a range of different responses to these questions. Another
question some ethicists have raised regarding GE animals concerns respect for the telos
of the animal. Telos refers to the “genetically based drives or instincts that, if frustrated,
would result in a significant compromise to the welfare of an animal” (Thompson, 2010,
p. 817). Some ethicists claim that it may be morally acceptable to alter an animal’s telos
using GE so long as it enhances wellbeing (Rollin, 1998), while others have argued that
it is not (Fiester, 2008).
AREA Technology /
Product
Potential Benefits Potential Harm
Food GE crops Less pesticides and
Herbicides.
Less fertilizer.
No till Agriculture
(soil conservation)
Extrinsic
Resistant pests (evolve)
Super weeds (outcrossing
and escape)
Irreversibility

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Environmentally
resilient crops
Crops with enhanced
nutritional value
Single generation
evolutionary impacts
Conflation of food and
medicine
Lack of knowledge
Accidental or incidental
negative impacts on
humans, animals, and
environment
Intrinsic and emotional
Playing God
Disrespectful to nature
Morally/spiritually wrong
Emotional yuk factor
GE animals Increased production
Healthier meat
More resilient
animals
(less medicines,
increased
environmental
tolerance)
Extrinsic
Reduced species diversity
Single generation
evolutionary impacts
Conflation of food and
medicine
Lack of knowledge
Accidental or incidental
negative impacts on
humans, animals, and
environment
Intrinsic and emotional
Playing God
Disrespectful to nature
Disrespectful to animal
telos
Morally/spiritually wrong
Emotional yuk factor

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Medicine Therapy
Medicines
derived from
GE
microorganisms,
plants, animals.
Gene therapy
Stems cells
Tissue
engineering
New medicines for
curing illness, and
injury
Organ replacement
Elimination of some
diseases
Increased life
expectancy
Outcrossing (and/or
escape)- Irreversibility -
Lack of knowledge
Accidental or incidental
negative impacts on
humans, animals, and
environment - Zoonotic
disease (e.g. from
xenotransplantations)
Overpopulation
Malevolent actions (GE
virus developed as weapon)
Intrinsic and emotional
Same as for GE food
animals
Enhancement
Somatic and
germline
therapy
(enhanced
physical,
social mental
capabilities, life
extension)
Chimeras
Enhanced human
(and non-human)
capabilities
Increased human
resilience
Disease elimination
Promotion of human
wellbeing
Much increased life
expectancy
Extrinsic
Super warriors - Eugenics
Lack of knowledge
Accidental or incidental
negative impacts on
humans, Fairness/justice
Autonomy - Species
divergence - Potential
enforcement -
Overpopulation
Intrinsic and emotional
Playing God - Disrespect to
nature - Disrespectful to
human telos - Morally/
spiritually wrong
Emotional yuk factor
Industry GE Pets, GE Pets with reduce Extrinsic

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plants, animals
microorganism
for
manufacturing
Chemicals and
materials
Energy and fuels
Synthetic biology
Bioinformatics
Biomimetics
allergic potential
New and existing
chemicals and
materials with a
range of new or
enhanced properties
Mitigation of peak oil
New production
methods and
processes
Outcrossing or escape
Dangerous organisms
Irreversibility - Competition
between food and fuel for
land and water - Lack of
knowledge - Accidental or
incidental negative impacts
on humans, animals, and
environment - Malevolent
bioweapons
Intrinsic and emotional
Same as for GE food crops
Ecosystem
services
Phytoremediation
Trees with
enhanced
carbon
absorption
Remediation of
pollution and toxic
sites
Climate change
mitigation
Extrinsic
Outcrossing or escape
Irreversibility
Lack of knowledge
Unforeseen or incidental
negative impacts on
humans, animals, and
environment
Accidental or incidental
negative impacts on
humans, animals, and
environment
Malevolent application as
bioweapons
Intrinsic and emotional
Playing God
Disrespectful to nature
Morally/spiritually wrong

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UNIT : 2 TOOLS OF GENETIC ENGINEERING
Enzymes, Non-specific endo and exo nuclease, DNase, RNase. Restriction
modification; restriction endonuclease- types, nome nclature, recongnition
sequence and mechanism of action. Methylation, RNA modification. Role of
kinases, phosphatases, polynuclcleotide phosphoryl ase, polynucleotide kinase
ligases – types and mechanism of action
VECTORS : General characteristics of vectors , brie f account of naturally
occurring plasmids promoters, MCS, Ori and maker ge ne-lac Z. construction of
pBR 322, pBR325, pUC18 and 19 vectors and expressio n vectors E.coli promoters,
lac promoters, trp promoters, lambda pL promoters, hybrid tac promoters,
ribosome binding site, codon selection. M13 derived vectors, Lambda based
vectors, cosmids, phagemids minichrosomes, BAC’s, Y AC’s , shuttle vectors, Ti
plasmids, vectors for animals-SV40 and Bovine papilloma virus.
ENZYMES, NON-SPECIFIC ENDO AND EXO NUCLEASE, DNASE, RNASE.
DNases: Deoxyribonuclease I cleaves double-stranded or single stranded DNA. Cleavage
preferentially occurs adjacent to pyrimidine (C or T) residues, and the enzyme is
therefore an endonuclease. Major products are 5'-ph osphorylated di, tri and
tetranucleotides.
In the presence of magnesium ions, DNase I hydrolyz es each strand of duplex DNA
independently, generating random cleavages. In the presence of manganese ions, the
enzyme cleaves both strands of DNA at approximately the same site, producing blunt
ends or fragments with 1-2 base overhangs. DNase I does not cleave RNA.
Some of the common applications of DNase I are:
• Eliminating DNA (e.g. plasmid) from preparations of RNA.
• Analyzing DNA-protein interactions via DNase footprinting.
• Nicking DNA prior to radiolabeling
by nick translation.
Exonuclease III(E. coli): Removes mononucleotides from the 3' termini of duplex DNA.
The preferred substrates are DNAs with blunt or 5' protruding ends. It will also extend
nicks in duplex DNA to create single-stranded gaps. In works inefficiently on DNA with
3' protruding ends, and is inactive on single-stranded DNA.
Mung Bean Nuclease : Digests single-stranded DNA to 5'-phosphorylated mono or
oligonucleotides. High concentrations of enzyme will also degrade double-stranded

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nucleic acids. Used to remove single-stranded extensions from DNA to produce blunt
ends.
Nuclease BAL 31 : Functions as an exonuclease to digest both 5' and 3' ends of double-
stranded DNA. It also acts as a single-stranded endonuclease that cleaves DNA at
nicks, gaps and single stranded regions. Does not cleave internally in duplex DNA.Used
for shortening fragments of DNA at both ends.
Nuclease S1: The substrate depends on the amount of enzyme used. Low
concentrations of S1 nuclease digests single-stranded DNAs or RNAs, while double-
stranded nucleic acids (DNA:DNA, DNA:RNA and RNA:RN A) are degraded by large
concentrations of enzyme. Moderate concentrations can be used to digest double
stranded DNA at nicks or small gaps.
Used commonly to analyze the structure of DNA:RNA hybrids (S1 nuclease mapping),
and to remove single-stranded extensions from DNA to produce blunt ends.
Ribonuclease A is an endoribonuclease that cleaves single-stranded RNA at the 3' end
of pyrimidine residues. It degrades the RNA into 3'-phosphorylated mononucleotides
and oligonucleotides.
RNases, which play important roles in nucleic acid metabolism, are found in both
prokaryotes and eukaryotes, and in practically every cell type. The human body uses
RNases to defend against invading microorganisms by secreting these enzymes in fluids
such as tears, saliva, mucus, and perspiration.
RNase H: RNase H (Ribonuclease H ) is an endoribonuclease that specifically hydrolyzes
the phosphodiester bonds of RNA which is hybridized to DNA. This enzyme does not
digest single or double-stranded DNA.
Applications:
Removal of poly(A) tails of mRNA hybridized to poly(dT)
Removal of mRNA during second strand cDNA synthesis
Ribonuclease I
f (RNase If) is a single strand specific RNA endonuclease which will cleave
at all RNA dinucleotide bonds leaving a 5´ hydroxyl and 2´, 3´ cyclic monophosphate (1).
RNase I
fis a recombinant protein fusion of RNase I (from E. coli) and maltose-binding
protein. It has identical activity to RNase I.
Applications:
Degradation of single-stranded RNA to mono-, di- and trinucleotide (3)

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Used in ribonuclease protection assays
RNase A:
RNase A is a pancreatic ribonuclease that cleaves s ingle-stranded RNA. Bovine
pancreatic RNase A is one of the classic model systems of protein science. The positive
charges of RNase A lie mainly in a deep cleft between two lobes. The RNA substrate lies
in this cleft and is cleaved by two catalytic histidine residues, His12 and His119, to
form a 2',3'-cyclic phosphate intermediate that is stabilized by nearby Lys41.
Eliminating or reducing RNA contamination in preparations of plasmid DNA.
Mapping mutations in DNA or RNA by mismatch cleavag e. RNase will cleave the RNA in
RNA:DNA hybrids at sites of single base mismatches, and the cleavage products can be
analyzed.
RESTRICTION MODIFICATION; RESTRICTION ENDONUCLEASE- TYPES,
NOMENCLATURE, RECONGNITION SEQUENCE AND MECHANISM O F ACTION.
Restriction Modification System
Phage (or viruses) invade all types of cells. Bacteria are one favorite target. Defense
mechanisms have been developed by bacteria to defen d themselves from these
invasions. The system they possess for this defense is the restriction-modificiation
system. This system is composed of a restriction en donuclease enzyme and a
methylase enzyme and each bacterial species and strain has their own combination of
restriction and methylating enzymes.
Restriction enzyme - an enzyme that cuts DNA at internal phosphodiest er bonds;
different types exist and the most useful ones for molecular biology (Type II) are those
which cleave at a specific DNA sequence
Methylase - an enzyme that adds a methyl group to a molecule ; in restriction-
modification systems of bacteria a methyl group is added to DNA at a specific site to
protect the site from restriction endonuclease cleavage
Several different types of restriction enzymes have been found but the most useful ones
for molecular biology and genetic engineering are the Type II restriction enzymes. These
enzymes cut DNA at specific nucleotide sequences. F or example, the enzyme EcoRI
recognizes the sequence:
5' - G A A* T T C - 3'
3' - C T T *A A G - 5'

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*The site of methylation protection from restriction enzyme cleavage is the 3'
adenine.This enzyme always cuts between the 5' G and A residues. But if we look at the
sequence we can see that both strands will be cut and leave staggered or overlapping
ends.
5' - G A A T T C - 3'
3' - C T T A A G - 5'
Not all Type II restriction enzymes generate staggered ends at the target site. Some cut
and leave blunt ends. For example, the enzyme BalI.
5' - T G G C* C A - 3'
3' - A C *C G G T - 5'
is cut at the point of symmetry to produce:
5' - T G G C C A - 3'
3' - A C C G G T - 5'
(Note: * The site of methylation protection from restriction enzyme cleavage; 5' cytosine)
The bacterial cell uses the restriction enzyme to cut the invading DNA of the virus at the
specific recognition site of the enzyme. This prevents the virus from taking over the
cellular metabolism for its own replication. But bacterial DNA will also contain sites
that could be cleaved by the restriction enzyme.
How is the bacterial cell protected? This protection is offered by the action of the
methylase. The methylase recognizes the same target site as the restriction enzyme and
adds a methyl group to a specific nucleotide in the restriction site. Methylated sites are
not substrates for the restriction enzyme. The bacterial DNA is methylated immediately
following replication so it will not be a suitable substrate for restriction endonuclease
cleavage. But it is unlikely that the invading viral DNA will have been methylated so it
will be an appropriate target for cleavage. Thus, the viral DNA is restricted in the
bacterial cell by the restriction enzyme, and the bacterial DNA is modified by the
methylase and is provided protection from its own restriction enzyme.
Isoschizomers are pairs of restriction enzymes specific to the s ame recognition
sequence.
Restriction endonucleases that recognize the same sequence are isoschizomers.

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For example, Sph I (CGTAC/G) and Bbu I (CGTAC/G) ar e isoschizomers of each other.
The first enzyme to recognize and cut a given sequence is known as the prototype, all
subsequent enzymes that recognize and cut that sequence are isoschizomers.
HpaII (recognition sequence: C↓CGG) and MspI (recognition sequence: C↓CGG).
Neoschizomer: An enzyme that recognizes the same sequence but cuts it differently is
a neoschizomer. Neoschizomers are a specific type (subset) of Isoschizomers.
For example, Sma I (CCC/GGG) and Xma I (C/CCGGG) ar e neoschizomers of each
other.
Thus, AatII (recognition sequence: GACGT↓C) and Zra I (recognition sequence:
GAC↓GTC) are neoschizomers of one another,
METHYLATION
Among these mechanisms, DNA methylation, or the enz ymatically mediated addition of
a methyl group to cytosine or adenine dinucleotides, serves as an inherited epigenetic
modification that stably modifies gene expression in dividing cells. The unique
methylomes are largely maintained in differentiated cell types, making them critical to
understanding the differentiation potential of the cell.
In the DNA methylation process, cytosine residues in the genome are enzymatically
modified to 5-methylcytosine, which participates in transcriptional repression of genes
during development and disease progression. 5-methy lcytosine can be further
enzymatically modified to 5-hydroxymethylcytosine by the TET family of methylcytosine
dioxygenases. DNA methylation affects gene transcription by physically interfering with
the binding of proteins involved in gene transcription.
Methylated DNA may be bound by methyl-CpG-binding d omain proteins (MBDs) that
can then recruit additional proteins. Some of these include histone deacetylases and
other chromatin remodeling proteins that modify histones, thereby forming compact,
inactive chromatin, or heterochromatin. While DNA m ethylation doesn’t change the
genetic code, it influences chromosomal stability and gene expression
.
RNA MODIFICATION.
PHOSPHATASES
Alkaline Phosphatase is an important tool in molecular biological processes like cloning.
It removes 3’- phosphate groups from a variety of substrates. Although in laboratory, it
is used to catalyze the removal of terminal 5’-(P), residues from single stranded or

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double stranded DNA and RNA. The resulting 5’ -OH termini can no longer take part in
ligation reactions, thus prevents self religation of vectors, reducing the background of
transformed bacterial colonies that carry empty plasmids. This enzyme works optimally
at alkaline pH (range of 8-9 in the presence of low Zn+2 concentrations) and hence
derived the name.
Alkaline Phosphatase is isolated from various sources:
Bacterial Alkaline phosphatase
Secreted in monomeric form into the Periplasmic space of E.coli, where it form dimers
and gets catalytically activated. It’s a remarkably stable enzyme and is resistant to
inactivation by heat and detergent. Thus, bacterial alkaline phosphatase is the most
difficult to destroy in the reaction mix.
Calf Intestinal Phosphatase
Calf intestinal phosphatase is a dimeric glycoprotein isolated from bovine intestine. This
has much more practical significance than bacterial alkaline phosphatase, since it can
be readily inactivated from the reaction mixture using proteinase K or by heating at
65˚C for 30 minutesor 75˚C for 15 minutes in the presence of 10mM EGTA.
Shrimp alkaline phosphatase
Extracted from cold water shrimp, can be inactivated readily by heating at 65˚C for 15
min.
POLYNUCLEOTIDE PHOSPHORYLASE
Polynucleotide Phosphorylase (PNPase) is a bifunctional enzyme with a
phosphorolytic 3' to 5' exoribonuclease activity and a 3'-terminal oligonucleotide
polymerase activity. That is, it dismantles the RNA chain starting at the 3' end and
working toward the 5' end. It also synthesizes long, highly heteropolymeric tails in vivo.
It accounts for all of the observed residual polyadenylylation in strains of Escherichia
coli missing the normal polyadenylylation enzyme.
It is involved on mRNA processing and degradation in bacteria, plants, and in humans.
In humans, the enzyme is encoded by the PNPT1 gene. In its active form, the protein
forms a ring structure consisting of three PNPase molecules. Each PNPase molecule
consists of two RNase PH domains, an S1 RNA binding domain and a K-homology
domain. The protein is present in bacteria and in the chloroplasts and mitochondria of
some eukaryotic cells.

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POLYNUCLEOTIDE KINASE
A kinase from bacteriophage T4 that transfers the γ-phosphate of ATP to the 5' end of
DNA or RNA. Used to end label DNA and to phosphorylate synthetic DNA
LIGASES – TYPES AND MECHANISM OF ACTION
DNA ligases close nicks in the phosphodiester backbone of DNA. Biologically, DNA
ligases are essential for the joining of Okazaki fragments during replication, and for
completing short-patch DNA synthesis occurring in DNA repair process.
The smallest known ATP-dependent DNA ligase is the one from the bacteriophage T7 (at
41KdA). Eukaryotic DNA ligases may be much larger (human DNA ligase I is >
100KDA) but they all appear to share some common sequences and probably structural
motifs.
DNA Ligase Mechanism
The reaction occurs in three stages in all DNA ligases:
Formation of a covalent enzyme-AMP intermediate linked to a lysine side-chain in the
enzyme.
Transfer of the AMP nucleotide to the 5’ phosphate of the nicked DNA strand.
Attack on the AMP-DNA bond by the 3’-OH of the nick ed DNA sealing the phosphate
backbone and resealing AMP.
The following figure illustrates the three reaction stages:
DNA ligases are Mg++-dependent enzymes that catalyze the formation of phosphodiester
bonds at single-strand breaks in double-stranded DNA. The first step in the reaction is
the formation of a covalent enzyme/adenylate intermediate.
The ATP is cleaved to AMP and pyrophosphate with th e adenylyl residue linked by a
phosphoramidate bond to the &-amino group of a spec ific lysine residue at the active
site of the protein. The reaction is readily revers ed in vitro by addition of
pyrophosphate.
The activated AMP residue of the DNA ligase/adenylate intermediate is transferred to
the 5 -phosphate terminus of a single-strand break in double-stranded DNA to generate
a covalent DNA-AMP complex with a 5'-5' phosphoanhy dride bond. In the final step of
DNA ligation, unadenylylated DNA ligase is required for the generation of a
phosphodiester bond and catalyzes displacement of the AMP residue through attack by
the adjacent 3 -hydroxyl group on the adenylylated site.

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DNA LIGASE I
It is present in all the eukaryotic cells. The main function of human DNA ligase I is
probably the joining of Okazaki fragments during lagging-strand DNA replication. The
enzyme is also involved in DNA excision repair. Similarly to DNA polymerase-A, DNA
ligase I is induced 10- to 15-fold in S phase in mammalian cells. Thus, it is present at
much higher concentrations in proliferating tissues.
DNA ligase I is a phosphoprotein, and most or all o f the phosphate residues are
localized to the amino-terminal region. Furthermore, the amino-terminal part is highly
susceptible to proteolysis, so a 78-kD active fragment of mammalian DNA ligase I,
comprising the catalytic domain of the enzyme, is often generated as a preparation
artifact due to endogenous degradation during enzyme purification.
The human gene encoding DNA ligase I is located at chromosome 19q13.2-13.3. The
gene covers 53 kb and contains 28 exons. DNA ligase I is more effective at blunt-end
joining than mammalian DNA ligases I1 and 111 but is less efficient in this regard than
bacteriophage T4 DNA ligase.
DNA LIGASE II
This approximately 69-kD DNA ligase can be distingu ished from DNA ligase I by its
ability to join an oligo(dT)*poly(rA) substrate. DNA ligase I1 is not induced on cell
proliferation, and its cellular role is not clear. It is the major DNA ligase activity in
certain nonproliferating tissues, e.g., adult liver.
The enzyme is more firmly retained in cell nuclei than DNA ligase I and requires buffers
of moderate or high salt concentration for efficient extraction.
DNA LIGASE III
DNA ligase 111 is a mammalian DNA ligase of 103 kD. The enzyme res embles DNA
ligase I in having a protease-sensitive amino-terminal region not required for DNA

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ligation activity in vitro, but it resembles DNA ligase I1 in its ability to join an
oligo(dT)*poly(rA) substrate.
DNA LIGASE IV
A fourth mammalian DNA ligase has been detected rec ently by a search of expressed
sequence tags with short sequence motifs highly conserved in eukaryotic DNA ligases
(Wei et al. 1995). The complete cDNA encodes a 96-kD protein with DNA ligase activity
that exhibits partial sequence homology with ligases 1-111.
The human gene for DNA ligase IV is localized on chromosome 13q33-34. Northern
blots indicate that the enzyme is expressed in thymus and testis and at a very low level
in several other tissues, but its physiological role is not known. An unusual feature of
DNA ligase IV, in comparison with the other mammalian DNA ligases, is an extended
carboxy-terminal region of more than 300 amino acids that shows no homology with
other proteins in databases. This region may conceivably be involved in protein-protein
interactions that could functionally distinguish this enzyme from the other DNA ligases.
VIRUS-ENCODED DNA LIGASES
Herpesviruses and smaller DNA and RNA animal viruse s do not encode a DNA ligase.
However, a distinct virus-encoded enzyme is produced by vaccinia virus and a number
of other poxviruses.
The 63-kD vaccinia protein only shows weak homology (-30%) with DNA ligase I from
human cells, S. pombe, or S. cerevisiae. Although poxviruses replicate in the host-cell
cytoplasm, the vaccinia DNA ligase is nonessential for viral DNA replication and growth.
However, a virus DNA ligase-deficient mutant shows attenuated virulence in vivo and is
anomalously sensitive to DNA-damaging agents during infection, implying a role for the
enzyme in viral DNA repair.
T4 DNA ligases:
T4 DNA ligase is an enzyme encoded by bacteriophage T4. It also catalyzes the covalent
joining of two segments to one uninterrupted strand in a DNA duplex, provided that no
nucleotides are missing at the junction (repair reaction). For its catalytic activity the
enzyme requires the presence of ATP and Mg
++
. DNAs that lack the required phosphate
residues can be rendered capable of ligation by phosphorylation with T4 polynucleotide
kinase.

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Catalyzes the formation of a phosphodiester bond between juxtaposed 5' phosphate and
3' hydroxyl termini in duplex DNA or RNA. This enzyme will join blunt end and cohesive
end termini as well as repair single stranded nicks in duplex DNA, RNA or DNA/RNA
hybrids.
The catalytic activity of the enzyme requires the presence of ATP and Mg++. DNAs that
lack the required phosphate residues can be rendere d capable of ligation by
phosphorylation with T4 polynucleotide kinase. The enzyme also catalyzes an addition
reaction of phosphate between pyrophosphate and ATP . The ligation and the repair
catalyzed reactions of T4 DNA ligases are illustrated in the following:
T4 DNA ligase is mainly used in joining DNAZ molecu les with compatible cohesive
termini, or blunt ended double stranded DNA to one another or to synthetic linkers.
It catalyzes a joining reaction between DNA molecules involving the 3' - hydroxy and the
5' - phosphate termini.
In addition the enzyme catalyzes an exchange reacti on of phosphate between
pyrophosphate and ATP.
Applications:
Cloning of restriction fragments
Joining linkers and adapters to blunt-ended DNA



VECTORS: GENERAL CHARACTERISTICS OF VECTORS
Vector is an agent that can carry a DNA fragment into a host cell in which it is capable
of replication. If it is used only for reproducing the DNA fragment, it is called a cloning
vector. If it is used for expression of foreign gene, it is called an expression vector.
Properties of a good vector:

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It should be autonomously replicating i.e. it should have ori region.
It should contain at least one selectable marker e. g. gene for antibiotic resistance.
It should have unique restriction enzyme site (only one site for one RE) for different REs
to insert foreign DNA.
It should be preferably small in size for easy handling.
It should have relaxed control of replication so that multiple copies can be obtained.
Vectors are of different types depending on the host. These are as follows:
Bacterial vectors
Yeast vectors
Plant vectors
Animal vectors
BRIEF ACCOUNT OF NATURALLY OCCURRING PLASMIDS PROMO TERS, MCS, ORI
AND MAKER GENE-LAC Z.
Protein synthesis, which is responsible for trait characteristics, requires genes to
undergo two steps: 1) transcription, or production of a messenger RNA (mRNA); and 2)
translation of mRNA into a protein. A gene has three major regions: the promoter,
coding region, and terminator. The promoter acts as the regulator for the level of gene
expression i.e. when, where and how much of the gene product (protein) is produced.
The coding region contains the information for making mRNA, which in turn specifies
the protein to be produced; while the terminator indicates the end of the gene.

Promoters regulate level of gene expression by spec ifying how many mRNAs are
produced (transcribed) for a given gene. The DNA se quence of the promoter region
interacts with transcription factor proteins that serve to recruit the cellular machinery
needed to produce the RNA transcripts. Transcription is performed by the enzyme, RNA
polymerase. The resultant RNA transcript is processed into mRNA, and then translated
into protein. The number of mRNAs produced is a pri mary factor determining the
amount of protein synthesized, which plays a role in determining the level of gene
expression.

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Factors that bind to promoters react to signals fro m the organism or/and the
surrounding environment. The source and type of sig nal determines the type of
promoters that are activated. In genetic engineering, there are three major types of
promoters used, depending on the level of gene expression and specificity required:
Constitutive promoters: These facilitate expression of the gene in all ti ssues
regardless of the surrounding environment and development stage of the organism.
Such promoters can turn on the gene in every living cell of the organism, all the time,
throughout the organism’s lifetime. These promoters can often be utilized across
species. Examples of constitutive promoters that are commonly used for plants include
Cauliflower mosaic virus (CaMV) 35S, opine promoters, plant ubiquitin (Ubi), rice actin
1 (Act-1) and maize alcohol dehydrogenase 1 (Adh-1). CaMV 35S is the most commonly
used constitutive promoter for high levels of gene expression in dicot plants. Maize Ubi
and rice Act-1 are the currently the most commonly used constitutive promoters for
monocots.
Tissue-specific or development-stage-specific promoters:
These facilitate expression of a gene in specific tissue(s) or at certain stages of
development while leaving the rest of the organism unmodified. In the case of plants,
such promoters might specifically influence expression of genes in the roots, fruits, or
seeds, or during the vegetative, flowering, or seed-setting stage. If the developer wants a
gene of interest to be expressed in more than one tissue type for example the root,
anthers and egg sac, then multiple tissue-specific promoters may have to be included in
the gene construct.
An example of a tissue-specific promoter is the phosphoenolpyruvate (PEP) carboxylase
promoter which induces gene expression only in cell s that are actively involved in
photosynthesis. In plant genetic engineering, this promoter is used for traits desired in
the shoot, leaves and sometimes the stem. Expressio n of genes controlled by this
promoter is reduced later in the growing season as the plant approaches senescence.
Inducible promoters:
These are activated by exogenous (i.e., external) factors. Exogenous factors may be
abiotic such as heat, water, salinity, chemical, or biotic like pathogen or insect attack.
Promoters that react to abiotic factors are the most commonly used in plant genetic
engineering because these can easily be manipulated . Such promoters respond to

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chemical compounds such as antibiotics, herbicides or changes in temperature or light.
Inducible promoters can also be tissue or development stage specific.
Promoters can be derived directly from naturally oc curring genes, or may be
synthesized to combine regulatory sequences from di fferent promoter regions. The
promoters interact with other regulatory sequences (enhancers or silencers) and
regulatory proteins (transcription factors) to influence the amount of gene
transcription/expression.
These are of two types:
Chemically-regulated promoters: including promoters whose transcriptional activity
is regulated by the presence or absence of alcohol, tetracycline, steroids, metal and
other compounds.
The activity of this class of promoters is modulated by chemical compounds that either
turn off or turn on gene transcription. As prerequisites, the chemicals influencing
promoter activity typically.
Should not be naturally present in the organism where expression of the transgene is
sought;
Should not be toxic;
Should affect only the expression of the gene of interest;
Should be easy to apply or removal; and
Should induce a clearly detectable expression pattern of either high or very low gene
expression.
Preferably, chemically-regulated promoters should be derived from organisms distant in
evolution to the organisms where its action is required. For example, promoters to be
used in plants are mostly derived from organisms such as yeast, E. coli, Drosophila or
mammals.
Alcohol-regulated: Syngenta has several patents and patent applications in Europe
and Australia directed to a transcriptional system containing the alcohol dehydrogenase
I (alcA) gene promoter and the transactivator prote in AlcR. Different agricultural
alcohol-based formulations are used to control the expression of a gene of interest
linked to the alcA promoter.

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Tetracycline-regulated: Yale University and BASF AG have several patents and
patent applications filed in the United States, Europe, Australia and Canada covering
aspects of tetracycline-responsive promoter systems, which can function either to
activate or repress gene expression system in the presence of tetracycline. Some of the
elements of the systems include a tetracycline repressor protein (TetR), a tetracycline
operator sequence (tetO) and a tetracycline transactivator fusion protein (tTA), which is
the fusion of TetR and a herpes simplex virus protein 16 (VP16) activation sequence.
Steroid-regulated: Numerous patent and patent applications are directed to steroid-
responsive promoters for the modulation of gene expression in plant and animal cells.
Metal-regulated: Promoters derived from metallothionein (proteins t hat bind and
sequester metal ions) genes from yeast, mouse and human are the subject matter of
several United States patents granted to Genentech, University Patents Inc .
and The University of California (Berkeley). DNA constructs having metal-regulated
promoters and eukaryotic cells transformed with them are claimed.
Pathogenesis-related (PR) proteins are induced in plants in the presence of particular
exogenous chemicals in addition to being induced by pathogen infection. Salicylic acid,
ethylene and benzothiadiazole (BTH) are some of the inducers of PR proteins. Promoters
derived from Arabidopsis and maize PR genes are the subject matter of patents granted
to Novartis and Pioneer Hi-Bred in the United States, Australia and Europe.
Physically-regulated promoters: including promoters whose transcriptional activity is
regulated by the presence or absence of light and low or high temperatures.
Promoters induced by environmental factors such as water or salt stress, anaerobiosis,
temperature, illumination and wounding have potential for use in the development of
plants resistant to various stress conditions. These promoters contain regulatory
elements that respond to such environmental stimuli.
Temperature-induced promoters include cold- and hea t-shock-induced promoters. In
many cases, these promoters are able to operate under normal temperature conditions,
which vary according to the organism, but when eith er cold or heat is applied, the
promoters maintain activity. In addition, expression can be enhanced by the application
of higher or lower temperature as compared to the normal temperature conditions. One
of the best studied eukaryotic heat-shock systems is the one found in Drosophila (fruit
fly).

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Heat-inducible promoters
Mycogen Plant Sciences, The United States Department of Health and Human Services
and The General Hospital Corporation have granted p atents and patent applications
that relate in general to DNA sequences of heat shock promoters and methods for
expressing a gene of interest under the control of such promoters. Some of the
inventions relate to the use of the heat shock promoters in transformed plants, while
others do not specify the organism to be transformed. For some properties of heat shock
promoters please also refer to the plant ubiquitin promoters.
Light-regulated promoters
The IP portfolio of Calgene Inc includes a United States patent that claims the use of
light responsive promoters in plant cells.
The other patents presented in this section relate to light-regulated promoters isolated
from genes of specific organisms. The University of Warwick in UK, Suntory LTD in
Japan and Mycogen Plant Sciences in the USA have filed patents on the use of
promoters whose expression is induced by light, such as a promoter isolated from
myxobacterium and promoters whose expression is inhibited by light exposure, such as
a promoter isolated from a pea gene.
The patents are classified according to whether the promoters are Light-
inducible or Light-repressible.
CONSTRUCTION OF PBR 322, PBR325, PUC18 AND 19 VECTO RS
In early cloning experiments, the cloning vectors used were natural plasmids, such as
Col E1 and pSC101. While these plasmids are small a nd have single sites for the
common restriction endonucleases, they have limited genetic markers for selecting
transformants. For this reason, considerable effort was expended on constructing, in
vitro, superior cloning vectors. The best and most widely used of these early purpose-
built vectors is pBR322. Plasmid pBR322 contains the ApR and TcR genes of RSF2124
and pSC101, respectively, combined with replication elements of pMB1, a Col E1-like
plasmid. The origins of pBR322 and its progenitor, pBR313, are shown in Fig. and
details of its construction can be found in the papers of Bolivar et al. (1977a,b).
The origins of plasmid pBR322.

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A.

B.
The boundaries between the pSC101, pMB1 and RSF2124 -derived material.
The numbers indicate the positions of the junctions in base pairs from the unique
EcoRI site.
The molecular origins of plasmid pBR322. R7268 was isolated in London in 1963
and later renamed R1. 1, A variant, R1drd19, which was derepressed for mating
transfer, was isolated. 2, The ApR transposon, Tn3, from this plasmid was
transposed on to pMB1 to form pMB3. 3, This plasmid was reduced in size by
EcoRI* rearrangement to form a tiny plasmid, pMB8, whi ch carries only colicin
immunity. 4, EcoRI* fragments from pSC101 were combined with pMB8 o pened at
its unique EcoRI site and the resulting chimeric molecule rearranged by EcoRI*
activity to generate pMB9. 5, In a separate event, the Tn3 of R1drd19 was
transposed to Col E1 to form pSF2124. 6, The Tn 3 element was then transposed
to pMB9 to form pBR312. 7, EcoRI* rearrangement of pBR312 led to the
formation of pBR313, from which (8) two separate fragments were isolated and
ligated together to form pBR322. During this series of constructions, R1 and Col
E1 served only as carries for Tn3. (Reproduced by courtesy of Dr G. Sutcliffe and
Cold Spring Harbor Laboratory.)

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Plasmid pBR322 has been completely sequenced. The o riginal published sequence
(Sutcliffe 1979) was 4362 bp long. Position O of the sequence was arbitrarily set
between the A and T residues of the EcoRI recognition sequence (GAATTC). The
sequence was revised by the inclusion of an additional CG base pair at position 526,
thus increasing the size of the plasmid to 4363 bp (Backman & Boyer 1983, Peden
1983). More recently, Watson (1988) has revised the size yet again, this time to 4361
bp, by eliminating base pairs at coordinates 1893 and 1915. The most useful aspect of
the DNA sequence is that it totally characterizes pBR322 in terms of its restriction sites,
such that the exact length of every fragment can be calculated. These fragments can
serve as DNA markers for sizing any other DNA fragment in the range of several base
pairs up to the entire length of the plasmid.
There are over 40 enzymes with unique cleavage sites on the pBR322 genome.
The target sites of 11 of these enzymes lie within the TcR gene, and there are sites for a
further two (ClaI and HindIII) within the promoter of that gene. There are unique sites
for six enzymes within the ApR gene. Thus, cloning in pBR322 with the aid of any one
of those 19 enzymes will result in insertional inactivation of either the ApR or the TcR
markers. However, cloning in the other unique sites does not permit the easy selection
of recombinants, because neither of the antibiotic resistance determinants is
inactivated.
Following manipulation in vitro, E. coli cells transformed with plasmids with inserts in
the TcR gene can be distinguished from those cells transformed with recircularized
vector. The former are ApR and TcS, whereas the lat ter are both ApR and TcR. In
practice, transformants are selected on the basis of their Ap resistance and then
replica-plated on to Tc-containing media to identify those that are TcS.
Cells transformed with pBR322 derivatives carrying inserts in the ApR gene can be
identified more readily (Boyko & Ganschow 1982). Detection is based upon the ability of
the β-lactamase produced by ApR cells to convert penicillin to penicilloic acid, which in
turn binds iodine. Transformants are selected on rich medium containing soluble
starch and Tc. When colonized plates are flooded with an indicator solution of iodine
and penicillin, β-lactamase-producing (ApR) colonies clear the indicator solution
whereas ApS colonies do not.

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The PstI site in the ApR gene is particularly useful, because the 3′ tetranucleotide
extensions formed on digestion are ideal substrates for terminal transferase. Thus this
site is excellent for cloning by the homopolymer tailing method.


If oligo(dG.dC) tailing is used, the PstI site is regenerated and the insert may be cut out
with that enzyme. Plasmid pBR322 has been a widely used cloning vehicle. In addition,

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it has been widely used as a model system for the study of prokaryotic transcription
and translation, as well as investigation of the effects of topological changes on DNA
conformation.
The popularity of pBR322 is a direct result of the availability of an extensive body of
information on its structure and function. This in turn is increased with each new
study. The reader wishing more detail on the struct ural features, transcriptional
signals, replication, amplification, stability and conjugal mobility of pBR322 should
consult the review of Balbás et al. (1986).
Example of the use of plasmid pBR322 as a vector: i solation of DNA fragments
which carry promoters
Cloning into the HindIII site of pBR322 generally results in loss of tetracycline
resistance. However, in some recombinants, TcR is retained or even increased. This is
because the HindIII site lies within the promoter rather than the coding sequence.
Thus whether or not insertional inactivation occurs depends on whether the cloned
DNA carries a promoter-like sequence able to initiate transcription of the TcR gene.
Widera et al. (1978) have used this technique to search for promo ter-containing
fragments.
Four structural domains can be recognized within E. coli promoters. These are:
• Position 1, the purine initiation nucleotide from which RNA synthesis begins;
• Position −6 to −12, the Pribnow box;
• The region around base pair −35;
• The sequence between base pairs −12 and −35.
Although the HindIII site lies within the Pribnow box (Rodriguez et al. 1979) the box is
re-created on insertion of a foreign DNA fragment. Thus when insertional inactivation
occurs it must be the region from −13 to −40 which is modified.
Restriction map of plasmid pBR322 showing the location and direction of transcription
of the ampicillin (Ap) and tetracycline (Tc) resistance loci, the origin of replication (ori)
and the Col E1-derived Rop gene. The map shows the restriction sites of those enzymes
that cut the molecule once or twice. The unique sites are shown in bold type. The
coordinates refer to the position of the 5′ base in each recognition sequence with the
first T in the EcoRI site being designated as nucleotide number 1. The exact positions of

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the loci are: Tc, 86–1268; Ap, 4084–3296; Rop, 1918–2105 and the origin of replication,
2535.
Much of the early work on the improvement of pBR322 centred on the insertion of
additional unique restriction sites and selectable markers, E.g. pBR325 encodes
chloramphenicol resistance in addition to ampicillin and tetracycline resistance and has
a unique EcoRI site in the CmR gene. Initially, each new vector was constructed in a
series of steps analogous to those used in the generation of pBR322 itself.
Then the construction of improved vectors was simplified (Vieira & Messing 1982, 1987,
Yanisch-Perron et al. 1985) by the use of polylinkers or multiple cloning sites (MCS), as
exemplified by the pUC vectors.
Over the years, numerous different derivatives of pBR322 have been constructed, many
to fulfil special-purpose cloning needs. A compilation of the properties of some of these
plasmids has been provided by Balbás et al. (1986).
An MCS is a short DNA sequence, 2.8 kb in the case of pUC19, carrying sites for many
different restriction endonucleases. An MCS increases the number of potential cloning
strategies available by extending the range of enzymes that can be used to generate a
restriction fragment suitable for cloning. By combining them within an MCS, the sites
are made contiguous, so that any two sites within it can be cleaved simultaneously
without excising vector sequences.
Improved vectors derived from pBR322
The pUC vectors also incorporate a DNA sequence that permits rapid visual detection of
an insert. The MCS is inserted into the lacZ′ sequence, which encodes the promoter
and the α-peptide of β- galactosidase. The insertion of the MCS into the lacZ′ fragment
does not affect the ability of the α-peptide to mediate complementation, but cloning
DNA fragments into the MCS does. Therefore, recombi nants can be detected by
blue/white screening on growth medium containing Xgal.
The usual site for insertion of the MCS is between the iniator ATG codon and codon 7, a
region that encodes a functionally non-essential part of the α-complementation peptide.
Recently, Slilaty and Lebel (1998) have reported that blue/white colour selection can be
variable. They have found that reliable inactivation of complementation occurs only
when the insert is made between codons 11 and 36.

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Cloning foreign DNA using the PstI site of pBR322.

Cut both the plasmid and the insert (yellow) with PstI, and then join them through
these sticky ends with DNA ligase.
Next, transform bacteria with the recombinant DNA a nd screen for tetracycline-
resistant, ampicillin-sensitive cells.
The recombinant plasmid no longer confers ampicillin resistance because the foreign
DNA interrupts that resistance gene (blue).
CONSTRUCTION OF EXPRESSION VECTORS
Expression vectors are required if one wants to prepare RNA probes from the cloned
gene or to purify large amounts of the gene product. In either case, transcription of the
cloned gene is required. Although it is possible to have the cloned gene under the
control of its own promoter, it is more usual to utilize a promoter specific to the vector.
Such vector-carried promoters have been optimized f or binding of the E.coli RNA
polymerase and many of them can be regulated easily by changes in the growth
conditions of the host cell.

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E. coli RNA polymerase is a multi-subunit enzyme. The core enzyme consists of two
identical α subunits and one each of the β and β′ subunits. The core enzyme is not
active unless an additional subunit, the σ factor, is present. RNA polymerase
recognizes different types of promoters depending on which type of σ factor is attached.
The most common promoters are those recognized by the RNA polymerase with σ70. A
large number of σ70 promoters from E. coli have been analysed and a compilation of
over 300 of them can be found in Lisser and Margalit (1993). A comparison of these
promoters has led to the formulation of a consensus sequence

The base sequence −10 and −35 regions of four natur al promoters, two hybrid
promoters and the consensus promoter.
If the transcription start point is assigned the position +1 then this consensus sequence
consists of the −35 region (5′-TTGACA-) and the −10 region, or Pribnow box (5′-TATAAT).
RNA polymerase must bind to both sequences to initiate transcription. The strength of a
promoter, i.e. how many RNA copies are synthesized per unit time per enzyme molecule,
depends on how close its sequence is to the consensus. While the −35 and −10 regions
are the sites of nearly all mutations affecting promoter strength, other bases flanking
these regions can affect promoter activity (Hawley & McClure 1983, Dueschle et al.
1986, Keilty & Rosenberg 1987). The distance between the −35 and −10 regions is also
important. In all cases examined, the promoter was weaker when the spacing was
increased or decreased from 17 bp. Upstream (UP) el ements located 5′ of the −35
hexamer in certain bacterial promoters are A+T-rich sequences that increase
transcription by interacting with the α subunit of RNA polymerase. Gourse et al. (1998)
have identified UP sequences conferring increased activity to the rrn core promoter. The
best UP sequence was portable and increased heterologous protein expression from the
lac promoter by a factor of 100.

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Once RNA polymerase has initiated transcription at a promoter, it will polymerize
ribonucleotides until it encounters a transcription-termination site in the DNA.
Bacterial DNA has two types of transcription-termination site: factor-independent and
factor-dependent. As their names imply, these types are distinguished by whether they
work with just RNA polymerase and DNA alone or need other factors before they can
terminate transcription. The factor-independent transcription terminators are easy to
recognize because they have similar sequences: an inverted repeat followed by a string
of A residues.

Structure of a factor-independent transcriptional terminator.
Transcription is terminated in the string of A residues, resulting in a string of U
residues at the 3′ end of the mRNA. The factordependent transcription terminators have
very little sequence in common with each other. Rather, termination involves interaction
with one of the three known E. coli termination factors, Rho (ρ), Tau (τ) and NusA. Most
expression vectors incorporate a factor-independent termination sequence downstream
from the site of insertion of the cloned gene.
Vectors for making RNA probes
Although single-stranded DNA can be used as a seque nce probe in hybridization
experiments, RNA probes are preferred.

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Method for preparing RNA probes from a cloned DNA m olecule using a phage SP6
promoter and SP6 RNA polymerase.
The reasons for this are that the rate of hybridization and the stability are far greater for
RNA–DNA hybrids compared with DNA–DNA hybrids. To m ake an RNA probe, the
relevant gene sequence is cloned in a plasmid vector such that it is under the control of
a phage promoter. After purification, the plasmid is linearized with a suitable restriction
enzyme and then incubated with the phage RNA polyme rase and the four
ribonucleoside triphosphates. No transcription terminator is required because the RNA
polymerase will fall off the end of the linearized plasmid. There are three reasons for
using a phage promoter. First, such promoters are very strong, enabling large amounts
of RNA to be made in vitro. Secondly, the phage promoter is not recognized by the E. coli
RNA polymerase and so no transcription will occur inside the cell. This minimizes any
selection of variant inserts. Thirdly, the RNA polymerases encoded by phages such as
SP6, T7 and T3 are much simpler molecules to handle than the E. coli enzyme, since
the active enzyme is a single polypeptide. If it is planned to probe RNA or single-
stranded
DNA sequences, then it are essential to prepare RNA probes corresponding to both
strands of the insert. One way of doing this is to have two different clones
corresponding to the two orientations of the insert.
An alternative method is to use a cloning vector in which the insert is placed between
two different, opposing phage promoters (e.g. T7/T3 or T7/SP6) that flank a multiple
cloning sequence.
Since each of the two promoters is recognized by a different RNA polymerase, the
direction of transcription is determined by which polymerase is used.
Structure and use of the LITMUS vectors for making RNA probes. (a) Structure of
the LITMUS vectors showing the orientation and rest riction sites of the four
polylinkers.

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A further improvement has been introduced by Evans et al. (1995). In their LITMUS
vectors, the polylinker regions are flanked by two modified T7 RNA polymerase
promoters. Each contains a unique restriction site (SpeI or AflII) that has been
engineered into the T7 promoter consensus sequence such that cleavage with the
corresponding endonuclease inactivates that promote r. Both promoters are active
despite the presence of engineered sites. Selective unidirectional transcription is
achieved by simply inactivating the other promoter by digestion with SpeI or AflII prior
to in vitro transcription.
Since efficient labelling of RNA probes demands that the template be linearized prior to
transcription, at a site downstream from the insert, cutting at the site within the
undesired promoter performs both functions in one s tep. Should the cloned insert
contain either an SpeI or an AflII site, the unwanted promoter can be inactivated by
cutting at one of the unique sites within the polylinker.
Method of using the LITMUS vectors to selectively s ynthesize RNA probes from
each strand of a cloned insert.

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E.COLI PROMOTERS
LAC PROMOTERS
It is usually advantageous to keep a cloned gene repressed until it is time to express it.
One reason is that eukaryotic proteins produced in large quantities in bacteria can be
toxic. Even if these proteins are not actually toxic, they can build up to such great levels
that they interfere with bacterial growth. In either case, if the cloned gene were allowed
to remain turned on constantly, the bacteria bearing the gene would never grow to a
great enough concentration to produce meaningful quantities of protein product. The

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solution is to keep the cloned gene turned off by placing it downstream of an inducible
promoter that can be turned off. The lac promoter is inducible to a certain extent,
presumably remaining off until stimulated by the sy nthetic inducer
isopropylthiogalactoside (IPTG). However, the repression caused by the lac repressor is
incomplete (leaky), and some expression of the cloned gene will be observed even in the
absence of inducer. One way around this problem is to express a gene in a plasmid or
phagemid that carries its own lacI (repressor) gene, as pBS does.
The pBluescript vector:

This plasmid is based on pBR322 and has that vector ’s ampicillin resistance gene
(green) and origin of replication (purple). In addition, it has the phage f1 origin of
replication (orange). Thus, if the cell is infected by an f1 helper phage to provide the
replication machinery, single-stranded copies of the vector can be packaged into
progeny phage particles. The multiple cloning site (MCS, red) contains 21 unique
restriction sites situated between two phage RNA polymerase promoters ( T7 and T3).
Thus, any DNA insert can be transcribed in vitro to yield an RNA copy of either strand,
depending on which phage RNA polymerase is provided. The MCS is embedded in an E.
coli lacZ′ gene (blue), so the uncut plasmid will produce the β-galactosidase N-terminal
fragment when an inducer such as isopropylthiogalac toside (IPTG) is added to
counteract the repressor made by the lacI gene (yellow). Thus, clones bearing the uncut
vector will turn blue when the indicator X-gal is added. By contrast, clones bearing
recombinant plasmids with inserts in the MCS will have an interrupted lacZ′ gene, so no
functional β-galactosidase is made. Thus, these clones remain white. The excess

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repressor produced by such a vector keeps the cloned gene turned off until it is time to
induce it with IPTG.
TRP PROMOTERS
The main function of an expression vector is to yield the product of a gene—usually, the
more products the better. Therefore, expression vectors are ordinarily equipped with
very strong promoters; the rationale is that the more mRNA that is produced, the more
protein product will be made. One such strong promoter is the trp (tryptophan operon)
promoter. It forms the basis for several expression vectors, including ptrpL1.
Two uses of the ptrpL 1 expression vector:


The vector contains a ClaI cloning site, preceded by a Shine–Dalgarno ribosome binding
site (SD) and the trp operator–promoter region (trpO,P). Transcription occurs in a
counterclockwise direction as shown by the arrow (top). The vector can be used as a
traditional expression vector (left) simply by inserting a foreign coding region (X, green)
into the unique ClaI site. Alternatively (right), the trp control region (purple) can be cut

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out with ClaI and HindIII and inserted into another plasmid bearing the coding region
(Y, yellow) to be expressed.
It has the trp promoter/operator region, followed by a ribosome binding site, and can be
used directly as an expression vector by inserting a foreign gene into the ClaI site.
Alternatively, the trp control region can be made “portable” by cutting it out with ClaI
and HindIII and inserting it in front of a gene to be expressed in another vector.
LAMBDA PL PROMOTERS
Another strategy is to use a tightly controlled promoter such as the λ phage promoter
PL. Expression vectors with this promoter–operator system are cloned into host cells
bearing a temperature-sensitive λ repressor gene (c1857). As long as the temperature of
these cells is kept relatively low (32° C), the repressor functions, and no expression
takes place. However, when the temperature is raised to the nonpermissive level (42° C),
the temperaturesensitive repressors can no longer function and the cloned gene is
derepressed.
A very popular method of ensuring tight control, as well as high-level induced
expression, is to place the gene to be expressed in a plasmid under control of a T7
phage promoter. Then this plasmid is placed in a cell that contains a tightly regulated
gene for T7 RNA polymerase. For example, the T7 RNA polymerase gene may be under
control of a modified lac promoter in a cell that also carries the gene for the lac
repressor. Thus, the T7 polymerase gene is strongly repressed unless the lac inducer is
present. As long as no T7 polymerase is present, transcription of the gene of interest
cannot take place because the T7 promoter has an ab solute requirement for its own
polymerase. But as soon as a lac inducer is added, the cell begins to make T7
polymerase, which transcribes the gene of interest. And because many molecules of T7
polymerase are made, the gene is turned on to a very high level and abundant amounts
of protein product are made.
HYBRID TAC PROMOTERS
When maximizing gene expression it is not enough to select the strongest promoter
possible: the effects of overexpression on the host cell also need to be considered. Many
gene products can be toxic to the host cell even when synthesized in small amounts.
Examples include surface structural proteins (Beck & Bremer 1980), proteins, such as
the PolA gene product, that regulate basic cellular metabolism (Murray & Kelley 1979),

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the cystic fibrosis transmembrane conductance regulator (Gregory et al. 1990) and
lentivirus envelope sequences (Cunningham et al. 1993). If such cloned genes are
allowed to be expressed there will be a rapid selection for mutants that no longer
synthesize the toxic protein.
Even when overexpression of a protein is not toxic to the host cell, high-level synthesis
exerts a metabolic drain on the cell. This leads to slower growth and hence in culture
there is selection for variants with lower or no expression of the cloned gene because
these will grow faster.
To minimize the problems associated with high-level expression, it is usual to use a
vector in which the cloned gene is under the control of a regulated promoter.
Many different vectors have been constructed for regulated expression of gene inserts
but most of those in current use contain one of the following controllable promoters: λ
PL, T7, trc (tac) or BAD.

Table shows the different levels of expression that can be achieved when the gene for
chloramphenicol transacetylase (CAT) is placed under the control of three of these
promoters.
The trc and tac promoters are hybrid promoters derived from the lac and trp promoters
(Brosius 1984). They are stronger than either of the two parental promoters because
their sequences are more like the consensus sequenc e. Like lac, the trc and tac
promoters are inducibile by lactose and isopropyl-β-d-thiogalactoside (IPTG). Vectors
using these promoters also carry the lacO operator and the lacI gene, which encodes the
repressor.
The pET vectors are a family of expression vectors that utilize phage T7 promoters to
regulate synthesis of cloned gene products (Studier et al. 1990). The general strategy for
using a pET vector is shown in Fig

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Strategy for regulating the expression of genes cloned into a pET vector.

The gene for T7 RNA polymerase (gene 1) is inserted into the chromosome of E.
coli and transcribed from the lac promoter; therefore, it will be expressed only if
the inducer
IPTG is added. The T7 RNA polymerase will then tran scribe the gene cloned into
the pET vector. If the protein product of the clone d gene is toxic, it may be
necessary to further reduce the transcription of the cloned gene before induction.
The T7 lysozyme encoded by a compatible plasmid, pL ysS, will bind to any
residual T7 RNA polymerase made in the absence of i nduction and inactivate it.
Also, the presence of lac operators between the T7 promoter and the cloned ge ne
will further reduce transcription of the cloned gene in the absence of the inducer
IPTG.
To provide a source of phage-T7 RNA polymerase, E. coli strains that contain gene 1 of
the phage have been constructed. This gene is cloned downstream of the lac promoter,
in the chromosome, so that the phage polymerase wil l only be synthesized following
IPTG induction. The newly synthesized T7 RNA polyme rase will then transcribe the
foreign gene in the pET plasmid. If the protein product of the cloned gene is toxic, it is
possible to minimize the uninduced level of T7 RNA polymerase. First, a plasmid
compatible with pET vectors is selected and the T7 lysS gene is cloned in it. When

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introduced into a host cell carrying a pET plasmid, the lysS gene will bind any residual
T7 RNA polymerase (Studier 1991, Zhang & Studier 19 97). Also, if a lac operator is
placed between the T7 promoter and the cloned gene, this will further reduce
transcription of the insert in the absence of IPTG (Dubendorff & Studier 1991).
Improvements in the yield of heterologous proteins can sometimes be achieved by use of
selected host cells (Miroux & Walker 1996).
The λ PL promoter system combines very tight transcriptional control with high levels of
gene expression. This is achieved by putting the cloned gene under the control of the PL
promoter carried on a vector, while the PL promoter is controlled by a cI repressor gene
in the E. coli host. This cI gene is itself under the control of the tryptophan (trp)
promoter,

In the absence of exogenous tryptophan, the cI gene is transcribed and the cI repressor
binds to the PL promoter, preventing expression of the cloned gene. Upon addition of
tryptophan, the trp repressor binds to the cI gene, preventing synthesis of the cI
repressor. In the absence of cI repressor, there is a high level of expression from the
very strong PL promoter. Many of the vectors designed for high-level expression also
contain translation-initiation signals optimized for E. coli expression.

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RIBOSOME BINDING SITE

Translation of mRNA into protein is a complex process which involves interaction of the
messenger with ribosomes. For translation to take place the mRNA must carry a
ribosome binding site (rbs) in front of the gene to be translated. After binding, the
ribosome moves along the mRNA and initiates protein synthesis at the first AUG codon
it encounters and continues until it encounters a stop codon (UAA, UAG or UGA). If the
cloned gene lacks a ribosome binding site, it is necessary to use a vector in which the
gene can be inserted downstream from both a promoter and an rbs.
CODON SELECTION
The hallmark of bacterial adaptation to novel envir onments is physiological
differentiation, whereby evolved organisms interact with their environments differently
than did their ancestors. Such physiological differentiation often involves
change in biochemical activities as the result of gene gain, gene loss, or the occurrence
of mutations that change the biochemical activities of existing gene products. These
adaptive shifts can be readily identified as changes in gene inventory (Ochman et al.
2000; Hacker and Carniel 2001) or as sites showing evidence of positive selection for
change (Nielsen and Yang 1998; Suzuki and Gojobori 1999). However, exploration of the
novel ecological niches afforded by these changes may also demand expression changes
among genes not involved in qualitative physiologic al adaptations. For example,
changes in the abundance of a familiar nutrient will result in a concomitant change in
the demand for the enzymes to metabolize that nutrient. Here, adaptation can occur

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through synonymous changes affecting the nature of mRNA/tRNA interactions. Such
codon selection is common
in genes of both prokaryotic and eukaryotic taxa (Sharp et al. 1988), most likely due to
the influence of codon identity on the duration for which a ribosome is occupied
synthesizing a particular polypeptide and/or its influence on the accuracy of translation
(Plotkin and Kudla 2010).
Selection on synonymous codons produces systemic biases in codon usage among the
open reading frames (ORFs) found in a genome, where the frequencies of certain codons
increase relative to their synonyms. Although codon selection is not the only selective
force that affects the nucleotide identity of synonymous sites, it is the primary selective
force in many bacteria, with the less-preferred codons existing as a result of mutation
and genetic drift (Bulmer 1991).
This bias increases in tandem with the expression level of the gene (Ikemura 1981),
indicating stronger selection in these ORFs (Sharp and Li 1987a,b). The genes encoding
core physiological processes often exhibit high frequencies of preferred codon usage
(Sharp and Li 1987b; Karlin and Mrazek 2000). Aside from widely conserved, highly
expressed genes (e.g., those encoding ribosomal proteins), enrichment for preferred
codon usage is also seen in genes that are distinctive to particular groups of bacteria
(e.g., photosynthesis genes in cyanobacteria [Mrazek et al. 2001]), indicating that codon
selection acts beyond those genes that are essentia l for all organisms. Although
differences in preferred codon usage have been noted among orthologous genes (Karlin
and Mrazek 2000), these differences have not been examined quantitatively; therefore,
the extent to which change in selection is responsible for such differences is unknown.
However, such changes are likely to be common as di fferences in gene expression
among lineages may arise from either regulatory cha nges or simple environmental
changes, thereby resulting in different levels of codon optimization in the orthologous
ORFs.
M13 DERIVED VECTORS
Another phage used as a cloning vector is the filamentous (long, thin, filament-like)
phage M13. Joachim Messing and his coworkers endowe d the phage DNA with the
same β-galactosidase gene fragment and multiple cloning sites found in the pUC family

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of vectors. In fact, the M13 vectors were engineered first; then the useful cloning sites
were simply transferred to the pUC plasmids.
Advantages of the M13 vectors:
The main factor is that the genome of this phage is a single-stranded DNA, so DNA
fragments cloned into this vector can be recovered in single-stranded form.
Single-stranded DNA can be an aid to sitedirected m utagenesis, by which we can
introduce specific, premeditated alterations into a gene. It also makes it easier to
determine the sequence of a piece of DNA.
Obtaining single-stranded DNA by cloning in M13 pha ge.
Foreign DNA (red), cut with HindIII, is inserted into the HindIII site of the double-
stranded phage DNA. The resulting recombinant DNA is used to transform E. coli cells,
whereupon the DNA replicates, producing many single -stranded product DNAs. The
product DNAs are called positive (+) strands, by convention. The template DNA is
therefore the negative (–) strand.

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Figure illustrates how to clone a double-stranded piece of DNA into M13 and harvest a
single-stranded DNA product.
The DNA in the phage particle itself is single-stranded, but after infecting an E. coli cell,
the DNA is converted to a double-stranded replicative form (RF). This double-stranded
replicative form of the phage DNA is used for cloning. After it is cut with one or two
restriction enzymes at its multiple cloning site, foreign DNA with compatible ends can
be inserted. This recombinant DNA is then used to transform host cells, giving rise to
progeny phages that bear single-stranded recombinan t DNA. The phage particles,
containing phage DNA, are secreted from the transformed cells and can be collected
from the growth medium.
LAMBDA BASED VECTORS - COSMIDS
Plasmids become unstable after a certain amount of DNA has been inserted into them,
because their increased size is more conducive to recombination. To circumvent this,
phage transduction is used instead. This is made possible by the cohesive ends, also
known as cos sites. In this way, they are similar to using the lambda phage as a vector,
but only that all the lambda genes have been deleted with the except ion of the cos
sequence.
It is possible to create an artificial plasmid consisting of two cos sites plus a plasmid
origin of replication and up to 46 kb of foreign DNA. This will replicate as a plasmid and
then can be packaged into lambda-phage heads for infection into bacteria.
These highly engineered vectors are a sort of cross between a plasmid and a
lambda phage and are capable of carrying 30-46 kb o f foreign genes with only a
little genetic material of their own. Like plasmids, these cosmids perpetuate in
bacteria and do not carry the genes for lytic development.
A cosmid, first described by Collins and Hohn in 1978.
A typical cosmid has:
1.
Replication origin
2.
Unique restriction sites.
3.
Selectable markers from plasmid.

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Structure of pJBB

A typical vector containing pBR322 module and a lambda segment having cos site.

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The cosmidpJB8 contains:
An origin of replication (ori) that allows it to replicate as a bacterial plasmid,
A cos gene for packing phage DNA into protein coats,
An ampicillin resistance gene (amp),
A region containing restriction sites for cloning (BamHI, EcoRI, ClaI, and HindIII).
Cos sequences are essential for packaging. They contain a cosN site where DNA is
nicked at each strand, 12bp apart, by terminase. Th is causes linearization of the
circular cosmid with two "cohesive" or "sticky ends" of 12bp. (The DNA must be linear to
fit into a phage head.) The cosB site holds the terminase while it is nicking and
separating the strands. The cos site of next cosmid is held by the terminase after the
previous cosmid has been packaged, to prevent degradation by cellular DNases.
Advantages of cosmids:
They can be used to clone DNA inserts of upto 40kb.
They can be packaged into lambda particles that infect host cells (which is many folds
more efficient than plasmid transformation).
The advantage of the use of cosmids for cloning is that its efficiency is high enough to
produce a complete genomic library of 10
6
- 10
7
clones from a. mere 1 pg of insert DNA.


Problems associated with lambda and cosmid cloning.

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Cosmids are difficult to maintain in a bacterial cell because they are somewhat
unstable.
The disadvantage, however, is its inability to accept more than 40-50 kbp of DNA.
Concatemers: These are long continuous DNA molecules that contain multiple copies
of the same DNA sequences linked in series. These p olymeric molecules are usually
copies of an entire genome linked end to end and se parated by cos sites (a protein
binding nucleotide sequence that occurs once in each copy of the genome).
A DNAsegment made up of repeated sequences linked e nd to end. Concatemers are
frequently the result of rolling circle replication, and may be seen in the late stage of
bacterial infection by phages. As an example, if the genes in the phage DNA are
arranged ABC, then in a concatemer the genes would be ABCABCABCABC and so on.
They are further broken by ribozymes.
PHAGEMIDS
Phagemids are very similar to M 13 and replicate in a similar fashion.One of the first
phagemid vectors to be developed, pEMBL, was constructed by inserting a fragment of
another phage, termed fl, containing a phage origin of replication and elements for its
morphogenesis into a pUC8 plasmid. Following superinfection with helper phage, the fl
origin is activated, allowing single-stranded DNA to be produced. The phage is
assembled into a phage coat extruded through the pe riplasm and secreted into the
culture medium in a similar way to M 13. Without su perinfection the phagemid
replicates as a pUC type plasmid and in the replicative form (RF) the DNA isolated is
double stranded. This allows further manipulation such as restriction digestion, ligation
and mapping analysis to be performed. The pBluescript SK vector is also a phagemid
and can be used in its own right as a cloning vector and manipulated as if it were a
plasmid. It can, like M 13, be used in nucleotide s equencing and site-directed
mutagenesis and it is also possible to produce RNA transcripts which may be used in
the production of labelled cRNA probes or riboprobes.

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MINICHROMOSOMES
Miniature chromosomes of 50– 150 kbp found in trypanosomes that carry silent copies
of the VSG gene. Variant surface glycoprotein (VSG) Glycoprotein found on surface of
trypanosomes that is encoded by multiple gene copies and varied to avoid recognition
by the animal immune system.
Parasitic eukaryotes live in hostile environments and must elude the defenses of their
host. Many adaptations have occurred in parasitic eukaryotes to overcome host defense
systems and allow the parasite to thrive in these o ne-sided relationships. Many
parasitic microorganisms attempt to elude the immun e system of their host by
changing their surface proteins.
The idea behind this strategy is straightforward. The immune memory cells recognize
the proteins on the surface of the original generation of invading germs. However, if
each successive wave of invaders changes its than average bacteria) and swim around
in the blood by means of a flagellum. Insects harbor these parasitic eukaryotes in their

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saliva, and when the insect bites a person or animal, the eukaryotic parasite invades
the bloodstream of the new host. For example, tsetse flies carry the trypanosomes that
cause sleeping sickness, whereas malarial parasites are carried by mosquitoes. While
growing and dividing in the insect gut, the trypanosomes are covered with a layer of a
protein called procyclin that protects them from digestion by the insect.
The trypanosomes then move to the salivary glands where they stop dividing and wait
for the insect to bite someone.While waiting, they convert their surface layer to the
variant surface glycoprotein (VSG) , designed to protect against animal immune
systems. The VSG protein has a variable region that is displayed on the trypanosome
surface and a conserved portion that anchors it to the membrane. It is found as a
dimer, as shown in Figure.

After transfer to a human, the trypanosomes grow an d divide in the blood until the
immune system kills most of them. However, a few of the trypanosomes switch their
VSG shape and escape recognition by the immune syst em. Eventually, the immune
system learns about the new surface protein and kills off most of the second wave of
trypanosomes. Meanwhile, some of the invaders have switched their VSG type again.
This continues and the infection therefore goes in waves, each spreading the invaders
further inside each human victim. The immune system never catches up with the
constantly changing outer layer of the trypanosome, and the normal result is death of
the victim.When tsetse flies suck blood from humans or animals with the disease, they
become re-infected. And so the cycle continues.
Trypanosomes alternate between two hosts, humans or other mammals and the tsetse
fly. While residing in the fly, trypanosomes acquire a procyclin coat to protect against
the fly’s digestive enzymes. After moving to the salivary glands of the fly, trypanosomes
start to express VSG proteins on the cell surface. When the tsetse fly bites a human,
the trypanosome enters the mammalian bloodstream an d starts to divide. Changes in
VSG surface expression allow a few of the trypanosomes to evade the immune system. If
a tsetse fly bites an infected human, the cycle starts all over again.

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To understand how the VSG protein is constantly altered, we must first discuss the
genomic structure of the trypanosomes. Both the nuclear and mitochondrial genomes of
trypanosomes are divided up in a peculiar manner. In addition, trypanosomes indulge
in the trans-splicing of many genes at the RNA level as well as RNA-editing
Each trypanosome cell contains one giant mitochondrion, the so-called “kinetoplast”.
This contains about 50 copies of a large circular DNA molecule that ranges from 20 to
80 kbp depending on the species. These “maxi-circles” encode the normal mitochondrial
genes. In addition there are approximately 10,000 mini-circles that encode only the
guide RNAs used in splicing.
The mini-circles range from 1.5 to 10 kbp and are often catenated (i.e. interlocked).
The nuclear genome is divided into 11 pairs of larg e “normal” or “megabase”
chromosomes plus about 100 mini-chromosomes of 50–150 kbp. The only
proteincoding genes found on the mini-chromosomes are silent copies of the VSG gene,
located close to one or both ends.
Trypanosome genomes consist of four different genetic elements. The mitochondria or
kinetoplast has maxi-circles and mini-circles. The nucleus has eleven pairs of megabase
chromosomes, and about 100 mini-chromosomes. The VS G genes are located at the
ends of both the megabase and minichromosomes

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The rest mostly comprises a Another method to switc h expression is called end
swapping.The megabase chromosomes and mini-chromoso mes can exchange ends by
recombination between blocks of repeated sequences just to the inside of the VSG
genes.

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This allows about 200 alternative VSG genes to be exchanged into the telomeric
expression sites of the normal megabase chromosomes. Not all VSG genes are found at
the ends of the chromosomes, some are found as tandem arrays scattered throughout
the megabase chromosomes.These are not accessible b y chromosomal end-swapping
and may only be used by gene conversion.

All unexpressed copies of the VSG gene may be used to supply sequences for splicing
into the VSG genes in the expression sites. Usually, the complete variable region of the
VSG gene in the expression site is replaced with the complete variable region from one
of the 1,000 extra copies (Fig.A).
The constant region stays unchanged, as its name in dicates. Later in infection,
segments of various sizes from the spare VSG genes are used for replacement; anywhere
from just a few base pairs to the whole gene may be used (Fig.B).
Furthermore, just as with the genes encoding mammal ian antibodies, point mutations
occur in the VSG genes at higher than normal frequency. However, in the case of the
VSG genes, the mutations occur during the segment-swapping process, not afterwards.
BAC’S
Multicopy vectors, such as ColE1 derived plasmids, are valuable because they give
higher yields of DNA than single copy vectors. However, they also have disadvantages.
In particular, the inserts may be unstable especially if they are very long and contain
repeated sequences. Many times, unstable inserts are deleted from the plasmid by
recombination events. Eukaryotic DNA is particularly unstable in plasmids.Therefore;

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cloning large segments of eukaryotic DNA in bacteria is now done using bacterial
artificial chromosomes (BACs).
These are single copy vectors based on the F-plasmid of E. coli.They can accept inserts
of 300 kb or more. Electroporation is necessary to transform these large constructs into
E. coli host cells and the yields of DNA are low. Nonethele ss bacterial artificial
chromosomes have been widely used in the human geno me project and other
eukaryotic genome sequencing projects.
Another cloning vector used for larger eukaryotic DNA segments is the P1 artificial
chromosome (PAC). This cloning vector is derived from bacteriophage P1, and has been
used to carry inserts of up to 150 kb. Just like the lambda derived vectors, these PACs
require in vitro packaging. Artificial chromosomes based on P1 have also been made for
use in E. coli host cells.
YAC’S
Analysis of the genomes of higher organisms require s the cloning of much larger
fragments than for bacteria. Because eukaryotic genes contain introns they may be
hundreds of kilobases in length. Such large DNA fragments require special vectors. The
largest capacity vectors derived from bacteriophage can handle at most 100 kb

Consequently, “artificial chromosomes” have been developed to carry huge lengths of
eukaryotic DNA. Huge segments of DNA, up to 2,000 kb or 2 million basepairs may be
carried on yeast artificial chromosomes or YACs.

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For any replicon, whether plasmid or chromosome, to survive, the vector must have a
yeast specific origin of replication and a centrome re recognition sequence (Cen
sequence). The YAC has both of these elements. In a ddition, as required by all
eukaryotic chromosomes, telomere sequences are present on both ends.A yeast cell will
treat this structure, although artificial, as a chromosome. Of course, for practical use a
selectable marker and a suitable multiple cloning sites are also included.
Colossal amounts of cloned DNA can be inserted into a YAC and may thus be replicated
inside yeast cells. Because the recognition sequenc es for replication origins,
centromeres and telomeres are so similar among higher organisms, an added bonus is
that YACs will survive in mice and are even passed on from parent to
offspring.Admittedly, not every baby mouse inherits the YAC, nonetheless, this opens
the way for cloning the huge DNA sequences needed for engineering higher animals and
for sequencing their genomes.
The YAC has two forms, a circular form for growing in bacteria, and a linear form for
growing in yeast. The circular form can be manipula ted and grown like any other

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plasmid in bacteria since it has a bacterial origin of replication and an antibiotic
resistance gene. In order to use this in yeast, the circular form is isolated and linearized
such that the yeast telomere sequences are on each end. This form can accommodate
up to 2,000 kb of cloned DNA inserted into its multiple cloning site (MCS).
SHUTTLE VECTORS
A shuttle vector is a vector (usually a plasmid) constructed so that it can propagate in
two different host species. Therefore, DNA inserted into a shuttle vector can be tested or
manipulated in two different cell types. The main advantage of these vectors is they can
be manipulated in E. coli then used in a system which is more difficult or slower to use
(e.g. yeast, other bacteria).
Shuttle vectors include plasmids that can propagate in eukaryotes and prokaryotes
(e.g. both Saccharomyces cerevisiae and Escherichia coli) or in different species of
bacteria (e.g. both E. coliand Rhodococcus erythropolis). There are also adenovirus
shuttle vectors, which can propagate in E. coli and mammals.
Shuttle vectors are frequently used to quickly make multiple copies of the gene in E.
coli (amplification). They can also be used for in vitro experiments and modifications
(e.g. mutagenesis,PCR).
One of the most common types of shuttle vectors is the yeast shuttle vector. Almost all
commonly used S. cerevisiae vectors are shuttle vectors. Yeast shuttle vectors have
components that allow for replication and selection in both E. coli cells and yeast cells.
The E. coli component of a yeast shuttle vector includes an origin of replication and a
selectable marker, e.g. antibiotic resistance, Beta lactamase. The yeast component of a
yeast shuttle vector includes an autonomously repli cating sequence (ARS), a
yeast centromere (CEN), and a yeast selectable marker(e.g. URA3, a gene that encodes
an enzyme for uracil synthesis, Lodish et al. 2007).
TI PLASMIDS
Genes can also be introduced into plants, using vectors that can replicate in plant cells.
The common bacterial vectors do not serve this purp ose because plant cells cannot
recognize their prokaryotic promoters and replication origins. Instead, a plasmid
containing so-called T-DNA can be used. This is a piece of DNA from a plasmid known
as Ti (tumor-inducing). The Ti plasmid inhabits the bacte rium Agrobacterium
tumefaciens, which causes tumors called crown galls in dicotyledonous plants.

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When this bacterium infects a plant, it transfers its Ti plasmid to the host cells,
whereupon the T-DNA integrates into the plant DNA, causing the abnormal proliferation
of plant cells that gives rise to a crown gall. This is advantageous for the invading
bacterium, because the T-DNA has genes directing the synthesis of unusual organic
acids called opines. These opines are worthless to the plant, but the bacterium has
enzymes that can break down opines so they can serve as an exclusive energy source
for the bacterium.
The T-DNA genes coding for the enzymes that make opines (e.g., mannopine synthetase)
have strong promoters. Plant molecular biologists take advantage of them by putting T-
DNA into small plasmids, then placing foreign genes under the control of one of these
promoters.
Crown gall tumours:
1.
Agrobacterium cells enter a wound in the plant, usually at the crown, or the junction of
root and stem.
2.
The Agrobacterium contains a Ti plasmid in addition to the much large r bacterial
chromosome. The Ti plasmid has a segment (the T-DNA , red) that promotes tumor
formation in infected plants.
3.
The bacterium contributes its Ti plasmid to the plant cell, and the T-DNA from the Ti
plasmid integrates into the plant’s chromosomal DNA.
4.
The genes in the T-DNA direct the formation of a cr own gall, which nourishes the
invading bacteria.

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Using a T-DNA plasmid to introduce a gene into tobacco plants.

a)
A plasmid is constructed with a foreign gene (red) under the control of the mannopine
synthetase promoter (blue). This plasmid is used to transform Agrobacterium cells.
b)
The transformed bacterial cells divide repeatedly.

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c) A disk of tobacco leaf tissue is removed and incubated in nutrient medium, along with
the transformed Agrobacterium cells. These cells infect the tobacco tissue, transferring
the plasmid bearing the cloned foreign gene, which integrates into the plant genome.
d)
The disk of tobacco tissue sends out roots into the surrounding medium.
e)
One of these roots is transplanted to another kind of medium, where it forms a shoot.
This plantlet grows into a transgenic tobacco plant that can be tested for expression of
the transplanted gene.
Outlines the process used to transfer a foreign gene to a tobacco plant, producing a
transgenic plant. One punch out a small disk (7 mm or so in diameter) from a tobacco
leaf and places it in a dish with nutrient medium. Under these conditions, tobacco
tissue will grow around the edge of the disk. Next, one adds Agrobacterium cells
containing the foreign gene cloned into a Ti plasmid; these bacteria infect the growing
tobacco cells and introduce the cloned gene.
VECTORS FOR ANIMALS-SV40
The first mammalian cell viral vector to be developed was based on the simian virus 40
(SV40) (Hamer and Leader, 1979). SV40 is a primate double stranded DNA tumor virus
whose genome is 5243 bp in size. Genes are encoded on both strands of the genome
such that they overlap each other. Virus has two life cycles depending upon host cell
line employed. In permissive cells (Monkey cells) a productive lytic cycle occurs while
in non permissive (rat or mouse cells) viral replication is blocked and host cells are
transformed (no growth as monolayer but proliferate without substratum attachment).
Recombinant SV40 vectors (rSV40) are good candidate s for gene transfer, as they
display some unique features: SV40 is a well-known virus, nonreplicative vectors are
easy-tomake, and can be produced in titers of 10(12 ) IU/ml. They also efficiently
transduce both resting and dividing cells, deliver persistent transgene expression to a
wide range of cell types, and are nonimmunogenic. P resent disadvantages of rSV40
vectors for gene therapy are a small cloning capacity and the possible risks related to
random integration of the viral genome into the host genome (Vera and Fortes, 2004).
SV40 was used to transduce gene expression in vitro and in vivo. Using cloned SV40
genome, we replaced large T antigen gene (Tag) with a polylinker, and inserted firefly
luciferase, controlled by SV40 early promoter. Transfection into Tag-expressing cells
yielded Tag-deficient virus, SVluc. SVluc was Tag-deficient and therefore replication-

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deficient in cells that did not supply Tag. SVluc transduced functional luciferase
expression in vitro. BALB/c mice were inoculated with SVluc, and their tissues were
assayed 3-21 days post-inoculation (dpi) for luciferase protein production and enzyme
activity. Luciferase protein was detected by immuno histochemistry throughout the
experiment, from 3 to 21 dpi. There was no inflamma tory reaction against SVluc-
infected cells at any time, in any tissue studied. Luciferase activity was first detected by
luminometry 14 dpi, and remained level through day 21. Thus, replication-deficient
recombinant SV40 can mediate gene transfer in vitro and in vivo.

Production of pBSV(∆T′).pBSV(∆T′) was made from pBSV-1, in which the complete SV40
genome was cloned as aBamHI fragment into pBR322. P artial digestion with AvrII and
complete digestion with BclI, removed the first exon and intron, and almost all of the
second exon of the Tag gene. A polylinker with 3 unique restriction sites, BstXI, XhoI,
and XbaI, flanked by Sp6 promoter beyond BstXI and T7 promoter beyond XbaI,
replaced the excised fragment. The polylinker had anAvrII site at the Sp6 end and a BclI
site at the T7 end. The bacteriophage promoters were included to facilitate sequencing
using standard Sp6 and T7 primers.

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VECTORS FOR BOVINE PAPILLOMA VIRUS
Bovine papilloma virus is of interest because its g enome can be maintained
extrachromosomally (at 20-200 copies per cell) in transformed or tumour cells.
Bovine papilloma virus (BPV-1) DNA replicates exclusively as an extrachromosomal
molecule in virally induced tumors as well as in transformed mouse fibroblasts in
culture. The complete viral genome or a 69% HindlII BamHI fragment thereof
have been used as vectors to introduce cloned proka ryotic or eukaryotic genes into
mammalian cells in culture. These recombinant molec ules replicate as multicopy
plasmids in stably transformed cells. This suggests that a broad potential exists for
BPV-1 DNAderived vectors (Matthias et al., 1983).

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UNIT : 3 GENE CLONING STRATEGIES AND CONSTRUCTION O F GENE LIBRARIES
Cloning from mRNA, isolation and purification of RN A, synthesis of cDNA,
Isolation of plasmids, cloning cDNA in plasmid vect ors, cloning cDNA in
bactriaophage vectors. cDNA library.
Cloning of genomic DNA: Isolation and purification of DNA, preparation of DNA,
fragments and cloning. Constriction of genomic libraries ( Using lambda gt 10 and
11 vectors ) In vitro packaging of lambda phage and amplification of libraries
Advanced cloning strategies synthesis and cloning o f cDNA, PCR amplified DNA,
use of adaptors and linkers, homopolymers tailing in cDNA cloning, expression of
cloned DNA molecule.
Selection, screening and analysis of recombinant Ge netic selection, insertional
inactivation, chromogenic substrates, complementati on of defined mutations,
nucleic acid hybridization, screening methods for cloned libraries, PCR screening
protocols, immunological screening, restriction mapping of cloned genes, blotting
techniques, sequencing methods. Purification strate gies of expressed His-tagged
proteins.
CLONING FROM mRNA

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First, eukaryotic cells are lysed and the mRNA is purified. Next, reverse transcriptase
plus primers containing oligo(dT) stretches are added. The oligo(dT) hybridizes to the
adenine in the mRNA poly(A) tail and acts as a primer for reverse transcriptase. This
enzyme makes the complementary DNA strand so formin g an mRNA/cDNA hybrid
molecule. The mRNA strand is digested with ribonuclease H and DNA polymerase I is
added to synthesize the opposite DNA strand, thus creating double-stranded cDNA. S1
nuclease trims off single-stranded ends. Since each mRNA has a different sequence,
linkers must be ligated to the ends of the cDNA to allow convenient insertion into the
cloning vector. After addition, the linkers are digested with the appropriate restriction
enzyme and the cDNA is ligated into the vector. The resulting hybrid DNA molecules are
then transformed into bacteria, so giving the final cDNA library.
ISOLATION AND PURIFICATION OF RNA
Most eukaryotic genes have intervening sequences of non-coding DNA (introns) between
the segments of coding sequence (exons). In higher eukaryotes, the introns are often
longer than the exons and the overall length of the gene is therefore much larger than
the coding sequence. This creates two problems. First, cloning large segments of DNA is
technically difficult; plasmids with large inserts are often unstable and transform
poorly. Secondly, bacteria cannot process RNA to remove the introns and so eukaryotic
genes containing introns cannot be expressed in bacterial cells.
Using a DNA copy of mRNA, known as complementary DNA or cDNA , solves both
problems since the mRNA has already been processed so that all the introns are
removed. To make a cDNA library, the mRNA must be i solated and used as a
template.The library generated therefore reflects only those genes expressed in the
particular tissue under the chosen conditions. First, total RNA is extracted from a
particular cell culture, tissue or specific embryonic stage. The messenger RNA from
eukaryotic cells is normally isolated from the total RNA by taking advantage of its
poly(A) tail. Since adenine base pairs to thymine or uracil, columns containing oligo(U)
or oligo(dT) tracts bound to a solid matrix are used to bind the mRNA by its poly(A) tail.
The mRNA is retained and the other RNA is washed through the column. The mRNA is
then released by eluting with a buffer of high ionic strength that disrupts the H-bonding
of the poly(A) tail to the oligo(dT).

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In order to isolate only messenger RNA from a sampl e of eukaryotic tissue, the
unique features of the mRNA molecule are used. Only mRNA has a poly(A) tail, a
long stretch of adenines following the coding seque nce. The poly(A) tail of the
mRNA will bind by base pairing to an oligonucleotid e consisting of a long stretch
of deoxythymidine residues—oligo(dT). The oligo(dT) is attached to glass or
magnetic beads, which consequently bind mRNA specif ically. Other RNAs will not
bind to the beads, and can be washed from the colum n.
A variation of this approach is the use of magnetic beads with attached oligo(dT) tracts.
After binding the mRNA the beads are separated magnetically.
SYNTHESIS OF cDNA
To generate cDNA the enzyme reverse transcriptase, originally found in retroviruses, is
added to the mRNA. This enzyme will make a compleme ntary DNA (cDNA) strand using
the mRNA as template. Several further steps are required to generate a double-stranded
cDNA copy of the original mRNA.
Ribonuclease H, which only recognizes RNA, is used to remove the mRNA strand of the
mRNA/cDNA hybrid molecule leaving a single-stranded cDNA.DNA polymeraseI is then
used to synthesize the second DNA strand. Any remai ning single-stranded ends are
trimmed off by S1 nuclease, which is an exonuclease specific for singlestranded regions
of DNA. [Such single-stranded ends mostly result from oligo(dT) primers binding in the
middle of the mRNA poly(A) tail.] The resulting doublestranded cDNA molecules can be
isolated and cloned into an appropriate vector, resulting in a cDNA library. Since each
mRNA has a different sequence, convenient restriction sites are generally added at each
end. This is done by attaching linkers— short pieces of DNA that have restriction sites
compatible with those in the multiple cloning site of the vector. Not only is cDNA easier
to handle, because the cloned fragments are much shorter than the original eukaryotic

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genes, but the cDNA versions of eukaryotic genes can often be successfully expressed in
bacteria.
ISOLATION OF PLASMIDS
An obvious prerequisite for cloning in plasmids is the purification of the plasmid DNA.
Although a wide range of plasmid DNAs are now routinely purified, the methods used
are not without their problems. Undoubtedly the trickiest stage is the lysis of the host
cells; both incomplete lysis and total dissolution of the cells result in greatly reduced
recoveries of plasmid DNA.
The ideal situation occurs when each cell is just sufficiently broken to permit the
plasmid DNA to escape without too much contaminatin g chromosomal DNA. Provided
the lysis is done gently, most of the chromosomal D NA released will be of high
molecular weight and can be removed, along with cel l debris, by high-speed
centrifugation to yield a cleared lysate. The production of satisfactory cleared lysates
from bacteria other than E. coli, particularly if large plasmids are to be isolated, is
frequently a combination of skill, luck and patience.
Many methods are available for isolating pure plasmid DNA from cleared lysates but
only two will be described here.
The first of these is the ‘classical’ method and is due to Vinograd (Radloff et al. 1967).
This method involves isopycnic centrifugation of cleared lysates in a solution of CsCl
containing ethidium bromide (EtBr). EtBr binds by intercalating between the DNA base
pairs, and in so doing causes the DNA to unwind. A CCC DNA molecule, such as
a plasmid, has no free ends and can only unwind to a limited extent, thus limiting the
amount of EtBr bound. A linear DNA molecule, such as fragmented chromosomal DNA,
has no such topological constraints and can therefore bind more of the EtBr molecules.
Because the density of the DNA–EtBr complex decreas es as more EtBr is bound, and
because more EtBr can be bound to a linear molecule than to a covalent circle, the
covalent circle has a higher density at saturating concentrations of EtBr.
Thus covalent circles (i.e. plasmids) can be separated from linear chromosomal DNA
Currently the most popular method of extracting and purifying plasmid DNA is that of
Birnboim and Doly (1979). This method makes use of the observation that there is a
narrow range of pH (12.0–12.5) within which denatur ation of linear DNA, but not
covalently closed circular DNA, occurs. Plasmid con taining cells are treated with

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lysozyme to weaken the cell wall and then lysed with sodium hydroxide and sodium
dodecyl sulphate (SDS). Chromosomal DNA remains in a high-molecular-weight form
but is denatured. Upon neutralization with acidic sodium acetate, the chromosomal
DNA renatures and aggregates to form an insoluble network. Simultaneously, the high
concentration of sodium acetate causes precipitation of protein–SDS complexes and of
highmolecular- weight RNA. Provided the pH of the alkaline denaturation step has been
carefully controlled, the CCC plasmid DNA molecules will remain in a native state and
in solution, while the contaminating macromolecules co-precipitate. The precipitate can
be removed by centrifugation and the plasmid concentrated by ethanol precipitation. If
necessary, the plasmid DNA can be purified further by gel filtration.
Recently a number of commercial suppliers of convenience molecular-biology products
have developed kits to improve the yield and purity of plasmid DNA. All of them take
advantage of the benefits of alkaline lysis and have as their starting-point the cleared
lysate. The plasmid DNA is selectively bound to an ion-exchange material, prepacked in
columns or tubes, in the presence of a chaotropic a gent (e.g. guanidinium
hydrochloride). After washing away the contaminants, the purified plasmid is eluted in
a small volume.

Purification of Col E1 KanR plasmid DNA by isopycnic centrifugation in a CsCl –
EtBr gradient.
Factors affecting the yield of plasmids

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The yield of plasmid is affected by a number of factors. The first of these is the actual
copy number inside the cells at the time of harvest. The copynumber control systems
described earlier are not the only factors affecting yield. The copy number is also
affected by the growth medium, the stage of growth and the genotype of the host cell
(Nugent et al. 1983, Seelke et al. 1987, Duttweiler & Gross 1998).
The second and most important factor is the care taken in making the cleared lysate.
Unfortunately, the commercially available kits have not removed the vagaries of this
procedure. Finally, the presence in the host cell of a wild-type endA gene can affect the
recovery of plasmid. The product of the endA gene is endonuclease I, a periplasmic
protein whose substrate is double-stranded DNA. The function of endonuclease I is not
fully understood.
Strains bearing endA mutations have no obvious phenotype other than im proved
stability and yield of plasmid obtained from them. Although most cloning vehicles are of
low molecular weight (see next section), it is sometimes necessary to use the much
larger conjugative plasmids. Although these high-molecular-weight plasmids can be
isolated by the methods just described, the yields are often very low. Either there is
inefficient release of the plasmids from the cells as a consequence of their size or there
is physical destruction caused by shear forces during the various manipulative steps. A
number of alternative procedures have been described (Gowland & Hardmann 1986),
many of which are a variation on that of Eckhardt (1978). Bacteria are suspended in a
mixture of Ficoll and lysozyme and these results in a weakening of the cell walls. The
samples are then placed in the slots of an agarose gel, where the cells are lysed by the
addition of detergent. The plasmids are subsequently extracted from the gel following
electrophoresis. The use of agarose, which melts at low temperature, facilitates
extraction of the plasmid from the gel.
CLONING cDNA IN PLASMID VECTORS
An ideal cloning vehicle would have the following three properties:
•
low molecular weight;
•
ability to confer readily selectable phenotypic traits
• on host cells;
•
Single sites for a large number of restriction endonucleases, preferably in genes with a
readily scorable phenotype.

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The advantages of a low molecular weight are several. First, the plasmid is much easier
to handle, i.e. it is more resistant to damage by shearing, and is readily isolated from
host cells. Secondly, lowmolecular- weight plasmids are usually present as multiple
copies, and this not only facilitates their isolation but leads to gene dosage effects for all
cloned genes. Finally, with a low molecular weight there is less chance that the vector
will have multiple substrate sites for any restriction endonuclease.

After a piece of foreign DNA is inserted into a vector, the resulting chimeric molecules
have to be transformed into a suitable recipient. Since the efficiency of transformation
is so low, it is essential that the chimeras have some readily scorable phenotype.
Usually this result from some gene, e.g. antibiotic resistance, carried on the vector, but
could also be produced by a gene carried on the inserted DNA.
One of the first steps in cloning is to cut the vector DNA and the DNA to be inserted
with either the same endonuclease or ones producing the same ends. If the vector has
more than one site for the endonuclease, more than one fragment will be produced.
When the two samples of cleaved DNA are subsequentl y mixed and ligated, the
resulting chimeras will, in all probability, lack one of the vector fragments. It is
advantageous if insertion of foreign DNA at endonuclease-sensitive sites inactivates a
gene whose phenotype is readily scorable, for in this way it is possible to distinguish
chimeras from cleaved plasmid molecules which have selfannealed. Of course, readily
detectable insertional inactivation is not essential if the vector and insert are to be
joined by the homopolymer tailing method or if the insert confers a new phenotype on
host cells.

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CLONING cDNA IN BACTERIOPHAGE VECTORS
Bacteriophage lambda, which infects E. coli, has been widely used as a cloning vector.
Lambda is a well-characterized virus with both lytic and lysogenic alternatives to its life
cycle. Although lambda DNA circularizes for replication and insertion into the E. coli
chromosome, the DNA inside the phage particle is linear.

In the lambda phage particle, the genome is a linea r DNA molecule with two cos
sequences at each end. After the phage injects its DNA into the bacterial host, the
DNA circularizes. The two cohesive ends base pair a nd are ligated together by
bacterial enzymes so forming a circle
At each end are complementary 12 bp long overhangs known as cos sequences
(cohesive ends). Once inside the E. coli host cell, these pair up and the cohesive ends
are ligated together by host enzymes forming the circular version of the lambda genome.
Only DNA molecules of between 37 and 52 kb can be s tably packaged into the head of
the lambda particle. Small fragments of extra DNA may be inserted into the lambda
genome without preventing packaging. However, to ac commodate longer inserts it is
necessary to remove some of the lambda genome.The l eft hand region has essential
genes for the structural proteins and the right hand region has genes for replication and
lysis.The middle region (~15 kb) of the lambda genome is non-essential and may be
replaced with approximately 23 kb of foreign DNA.

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Since lambda phage is easy to grow and manipulate, the genome has been
modified to accept foreign DNA inserts. The green region of the genome has genes
that are non-essential for lambda growth and packag ing. This region can be
replaced with large inserts of foreign DNA (up to about 23 kb). When used with a
helper phage, such modified lambdas provide useful cloning vectors.
Since the middle region of lambda has the genes for integration and recombination,
such lambda replacement vectors cannot integrate into the host chromosome and form
lysogens by themselves. To generate lysogens it is necessary to use a helper phage to
provide the integration and recombination functions.
If foreign DNA is inserted into the middle of lambda, the result is a linear DNA molecule
with two cohesive ends. To get such constructs into an E. coli host cell efficiently
requires in vitro packaging.
A lambda cloning vector containing cloned DNA must be packaged in a phage head
before it can infect E. coli. Before the DNA can be packaged, the phage head
proteins must be isolated. To do this, a culture of E. coli, is infected with a
mutant lambda which lacks the gene for one of the h ead proteins called E. A
different culture of E. coli is infected with a different lambda mutant, which lacks
phage head protein D. Both E. coli cultures are grown with the mutant lambdas
and the viruses are induced to enter the lytic cycle. Although the E. coli are lysed
by the phage, they cannot form complete heads. Inst ead a soluble mixture of
phage proteins is isolated.

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Each lysate contains phage tails, assembly proteins , and components of the
heads, except either D or E. These two lysates are mixed along with the lambda
vector containing the cloned DNA. Although mixing i s done in vitro, the
components can self-assemble into a functional phag e that can infect E. coli.
In this technique, a mixture of lambda proteins is mixed with the recombinant lambda
DNA in vitro to form phage particles. Infecting two separate E. coli cultures with two
different defective lambda mutants generates the necessary lambda proteins. Each of
the two mutants lacks an essential head protein and cannot form particles containing
its own DNA. A mixture of the two lysates gives a full set of lambda proteins and when
mixed with lambda DNA can generate infectious phage particles.
cDNA LIBRARY.
Under some circumstances, it may be possible to prepare cDNA directly from a purified
mRNA species. Much more commonly, a cDNA library is prepared by reverse-
transcribing a population of mRNAs and then screene d for particular clones. An
important concept is that the cDNA library is representative of the RNA population from

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which it was derived. Thus, whereas genomic librari es are essentially the same,
regardless of the cell type or developmental stage from which the DNA was isolated, the
contents of cDNA libraries will vary widely according to these parameters. A given cDNA
library will also be enriched for abundant mRNAs but may contain only a few clones
representing rare mRNAs. Furthermore, where a gene is differentially spliced, a cDNA
library will contain different clones representing alternative splice variants.

Table shows the abundances of different classes of mRNAs in two representative
tissues. Generally, mRNAs can be described as abundant, moderately abundant or rare.
Notice that, in the chicken oviduct, one mRNA type is superabundant. This encodes
ovalbumin, the major egg-white protein.
Therefore, the starting population is naturally so enriched in ovalbumin mRNA that
isolating the ovalbumin cDNA can be achieved withou t the use of a library. An
appropriate strategy for obtaining such abundant cDNAs is to clone them directly in an
M13 vector, such as M13 mp8. A set of clones can then be sequenced immediately and
identified on the basis of the polypeptide that each encodes. A successful demonstration
of this strategy was reported by Putney et al. (1983), who determined DNA sequences of
178 randomly chosen muscle cDNA clones. Based on th e amino acid sequences
available for 19 abundant muscle-specific proteins, they were able to identify clones
corresponding to 13 of these 19 proteins, including several protein variants.
For the isolation of cDNA clones in the moderateand low-abundance classes, it is
usually necessary to construct a cDNA library. Once again, the high efficiency obtained
by packaging in vitro makes phage- λvectors attractive for obtaining large numbers of
cDNA clones. Phage- λinsertion vectors are particularly well suited for Typically, 10
5
–

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10
6
clones are sufficient for the isolation of low-abundance mRNAs from most cell types,
i.e. those present at 15 molecules per cell or above.
However, some mRNAs are even less abundant than thi s, and may be further diluted if
they are expressed in only a few specific cells in a particular tissue. Under these
circumstances, it may be worth enriching the mRNA p reparation prior to library
construction, e.g. by size fractionation, and testing the fractions for the presence of the
desired molecule. One way in which this can be achieved is to inject mRNA fractions
into Xenopus oocytes and test them for production of the corresponding protein (Melton
1987).
CLONING OF GENOMIC DNA
Most early cDNA libraries were constructed using plasmid vectors, and were difficult to
store and maintain for long periods. They were largely replaced by phage-λ libraries,
which can be stored indefinitely and can also be prepared to much higher titres. λgt10
and λgt11 were the standard vectors for cDNA cloning until about 1990. Both λgt10 and
λgt11 are insertion vectors, and they can accept approximately 7.6 kb and 7.2 kb of
foreign DNA, respectively. In each case, the foreign DNA is introduced at a unique EcoRI
cloning site. λgt10 is used to make libraries that are screened by hybridization. The
EcoRI site interrupts the phage cI gene, allowing selection on the basis of plaque
morphology.
λgt11 contains an E. coli lacZ gene driven by the lac promoter. If inserted in the correct
orientation and reading frame, cDNA sequences cloned in this vector can be expressed
as β-galactosidase fusion proteins, and can be detected by immunological screening or
screening with other ligands. λgt11 libraries can also be screened by hybridization,
although λgt10 is more appropriate for this screening strategy because higher titres are
possible.
λZAP series
While phage-λ vectors generate better libraries, they cannot be manipulated in vitro with
the convenience of plasmid vectors. Therefore, phage clones have to be laboriously
subcloned back into plasmids for further analysis. This limitation of conventional
phage-λ vectors has been addressed by the development of hybrids, sometimes called
phasmids, which possess the most attractive features of both bacteriophage λ and
plasmids. The most popular current vectors for cDNA cloning are undoubtedly those of

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the λZAP series marketed by Stratagene (Short et al. 1988). A map of the original λZAP
vector is shown below.

The advantageous features of this vector are: (i) the high capacity – up to 10 kb of
foreign DNA can be cloned, which is large enough to encompass most cDNAs; (ii) the
presence of a polylinker with six unique restriction sites, which increases cloning
versatility and also allows directional cloning; and (iii) the availability of T3 and T7 RNA
polymerase sites flanking the polylinker, allowing sense and antisense RNA to be
prepared from the insert. Most importantly, all these features are included within a
plasmid vector called pBluescript, which is itself inserted into the phage genome. Thus
the cDNA clone can be recovered from the phage and propagated as a high-copy-
number plasmid without any subcloning, simply by co infecting the bacteria with a
helper f1 phage that nicks the λZAP vector at the flanks of the plasmid and facilitates
excision. Another member of this series, λZAP Expre ss, also includes the human
cytomegalovirus promoter and SV40 terminator, so th at fusion proteins can be
expressed in mammalian cells as well as bacteria. Thus, cDNA libraries can be cloned
in the phage vector in E. coli, rescued as plasmids and then transfected into
mammalian cells for expression cloning.
ISOLATION AND PURIFICATION OF DNA
PREPARATION OF DNA

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The synthesis of double-stranded cDNA suitable for insertion into a cloning vector
involves three major steps: (i) first-strand DNA synthesis on the mRNA template, carried
out with a reverse transcriptase; (ii) removal of the RNA template; and (iii) secondstrand
DNA synthesis using the first DNA strand as a templ ate, carried out with a DNA-
dependent DNA polymerase, such as E. coli DNA polymerase I. All DNA polymerases,
whether they use RNA or DNA as the template, requir e a primer to initiate strand
synthesis.
FRAGMENTS AND CLONING

An early cDNA cloning strategy, involving hairpin-p rimed second-strand DNA
synthesis and homopolymer tailing to insert the cDN A into the vector.

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The first reports of cDNA cloning were published in the mid-1970s and were all based
on the homopolymer tailing technique. Of several alternative methods, the one that
became the most popular was that of Maniatis et al. (1976). This involved the use of an
oligo-dT primer annealing at the polyadenylate tail of the mRNA to prime first-strand
cDNA synthesis, and took advantage of the fact that the first cDNA strand has the
tendency to transiently fold back on itself, forming a hairpin loop, resulting in self-
priming of the second strand (Efstratiadis et al. 1976). After the synthesis of the second
DNA strand, this loop must be cleaved with a single-strand-specific nuclease, e.g. S1
nuclease, to allow insertion into the cloning vector.
A serious disadvantage of the hairpin method is that cleavage with S1 nuclease results
in the loss of a certain amount of sequence at the 5′ end of the clone. This strategy has
therefore been superseded by other methods in which the second strand is primed in a
separate reaction. One of the simplest strategies is shown in Fig.

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Improved method for cDNA cloning.
The first strand is tailed with oligo(dC) allowing the second strand to be initiated
using an oligo(dG) primer.
After first-strand synthesis, which is primed with an oligo-dT primer as usual, the cDNA
is tailed with a string of cytidine residues using the enzyme terminal transferase. This
artificial oligo-dC tail is then used as an annealing site for a synthetic oligo-dG primer,
allowing synthesis of the second strand.
Using this method, Land et al. (1981) were able to isolate a full-length cDNA
corresponding to the chicken lysozyme gene. However, the efficiency can be lower for
other cDNAs (e.g. Cooke et al. 1980). For cDNA expression libraries, it is advantageous
if the cDNA can be inserted into the vector in the correct orientation. With the self-
priming method, this can be achieved by adding a sy nthetic linker to the double-
stranded cDNA molecule before the hairpin loop is cleaved (e.g. Kurtz & Nicodemus
1981; Fig).

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Where the second strand is primed separately, direction cloning can be achieved using
an oligo-dT primer containing a linker sequence (e.g. Coleclough & Erlitz 1985; Fig.).


An alternative is to use primers for cDNA synthesis that are already linked to a plasmid
(Fig).

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This strategy was devised by Okayama and Berg (1982 ) and has two further notable
characteristics. First, full-length cDNAs are preferentially obtained because an RNA–
DNA hybrid molecule, the result of first-strand synthesis, is the substrate for a terminal
transferase reaction.
A cDNA that does not extend to the end of the mRNA will present a shielded 3-hydroxyl
group, which is a poor substrate for tailing. Secondly, the second-strand synthesis step
is primed by nicking the RNA at multiple sites with RNase H. Second-strand synthesis
therefore occurs by a nick-translation type of reaction, which is highly efficient. Simpler
cDNA cloning strategies incorporating replacement synthesis of the second strand is
widely used (e.g. Gubler & Hoffman 1983, Lapeyre & Amalric 1985).
The Gubler–Hoffman reaction, as it is commonly known, is show in Fig.

The Gubler–Hoffman method, a simple and general met hod for non-directional
cDNA cloning. First-strand synthesis is primed usin g an oligo(dT) primer. When
the first strand is complete, the RNA is removed wi th RNase H and the second
strand is random-primed and synthesized with DNA po lymerase I. T4 DNA
polymerase is used to ensure that the molecule is b lunt-ended prior to insertion
into the vector.
Full-length cDNA cloning
Limitations of conventional cloning strategies
Conventional approaches to the production of cDNA l ibraries have two major
drawbacks. First, where oligo-dT primers are used to initiate first-strand synthesis,
there is generally a 3′-end bias (preferential recovery of clones representing the 3′ end of

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cDNA sequences) in the resulting library. This can be addressed through the use of
random oligonucleotide primers, usually hexamers, for both first- and second-strand
cDNA syntheses. However, while this eliminates 3′-end bias in library construction, the
resulting clones are much smaller, such that fulllength cDNAs must be assembled from
several shorter fragments. Secondly, as the size of a cDNA increases, it becomes
progressively more difficult to isolate full-length clones. This is partly due to deficiencies
in the reverse-transcriptase enzymes used for first-strand cDNA synthesis. The enzymes
are usually purified from avian myeloblastosis virus (AMV) or produced from a cloned
Moloney murine leukaemia virus (MMLV) gene in E. coli. Native enzymes have poor
processivity and intrinsic RNase activity, which leads to degradation of the RNA
template (Champoux 1995). Several companies produce engineered murine reverse
transcriptases that lack RNase H activity, and these are more efficient in the production
of full-length cDNAs (Gerard & D’Allesio 1993). An example is the enzyme Super- Script
II, marketed by Life Technologies (Kotewicz et al. 1988). This enzyme can also carry out
reverse transcription at temperatures of up to 50°C. The native enzymes function
optimally at 37°C and therefore tend to stall at sequences that are rich in secondary
structure, as often found in 5′ and 3′ untranslated regions.
Selection of 5′ mRNA ends
Despite improvements in reverse transcriptases, the generation of full-length clones
corresponding to large mRNAs remains a problem. Thi s has been addressed by the
development of cDNA cloning strategies involving the selection of mRNAs with intact 5′
ends. Nearly all eukaryotic mRNAs have a 5′ end cap , a specialized, methylated
guanidine residue that is inverted with respect to the rest of the strand and is
recognized by the ribosome prior to the initiation of protein synthesis. Using a
combination of cap selection and nuclease treatment, it is possible to select for full-
length first-strand cDNAs and thus generate libraries highly enriched in full-length
clones. An example is the method described by Edery et al. (1995).

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The CAPture method of full-length cDNA cloning, usi ng the eukaryotic initiation
factor eIF-4E to select mRNAs with caps protected f rom RNase digestion by a
complementary DNA strand.
In this strategy, first-strand cDNA synthesis is initiated as usual, using an oligo-dT
primer. Following the synthesis reaction, the hybrid molecules are treated with RNase
A, which only digests single-stranded RNA. DNA–RNA hybrids therefore remain intact. If
the first-strand cDNA is full-length, it reaches all the way to the 5′ cap of the mRNA,
which is therefore protected from cleavage by RNase A. However, part-length cDNAs will
leave a stretch of unprotected single-stranded RNA between the end of the double-
stranded region and the cap, which is digested away with the enzyme. In the next stage
of the procedure, the eukaryotic translational initiation factor eIF-4E is used to isolate
full-length molecules by affinity capture. Incomplete cDNAs and cDNAs synthesized on
broken templates will lack the cap and will not be retained. A similar method based on
the biotinylation of mRNA has also been reported (Caminci et al. 1996). Both methods,
however, also co-purify cDNAs resulting from the mispriming of first-strand synthesis,
which can account for up to 10% of the clones in a library.

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An alternative method, oligo-capping, addresses this problem by performing selection at
the RNA stage (Maruyama & Sugano 1994, Suzuki et al. 1997, 2000;).

Oligo-capping, the addition of specific oligonucleotide primers to full-length RNAs
by sequential treatment with alkaline phosphatase a nd acid pyrophosphatase.
Once the oligo cap has annealed to the 5′ end of th e mRNA, it can serve as a
primer binding site for PCR amplification.
The basis of the method is that RNA is sequentially treated with the enzymes alkaline
phosphatase and acid pyrophosphatase. The first enzyme removes phosphate groups
from the 5′ ends of uncapped RNA molecules, but does not affect full-length molecules
with a 5′ cap. The second treatment removes the cap from full-length RNAs, leaving a
5′-terminal residue with a phosphate group. Full-length molecules can be ligated to a
specific oligonucleotide, while broken and degraded molecules cannot. The result is an

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oligo-capped population of full-length mRNAs. This selected population is then reverse-
transcribed using an oligo-dT primer. Second-strand synthesis and cloning is then
carried out by PCR using the oligo-dT primer and a primer annealing to the
oligonucleotide cap. Only full-length cDNAs annealing to both primers will be amplified,
thus eliminating broken or degraded RNAs, incomplete first cDNA strands (which lack a
5′ primer annealing site) and misprimed cDNAs (which lack a 3′ primer annealing site).
CONSTRICTION OF GENOMIC LIBRARIES (USING LAMBDA GT 10 AND 11
VECTORS )
IN VITRO PACKAGING OF LAMBDA PHAGE AND AMPLIFICATIO N OF LIBRARIES
Lambda-sensitive bacteria are grown on agar plates in high density to form a lawn of
confluent growth. The artificially synthesized transducing phage are added in a
concentration resulting in about 100 phage particles per plate, hence producing about
100 plaques of lysed bacterial cells per plate. Each plaque contains phage clones
containing millions of identical phage genomes. Recombinant phage are produced when
foreign DNA (e. g. , mammalian DNA ) is ligated to the manipulated phage (Fig).

The viruses themselves can be cut and inserted into . These constructs are
called insertion vectors and differ from plasmids because they are detected by phage

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plaques in a bacterial lawn rather than antibiotic-resistance colonies. The uptake in
cells is also different for bacteriophage lambda and plasmids. Lambda undergoes its
lytic cycle and phage heads are produced within the bacterial cell it entered. Cos sites
determine where specific cuts are made in the DNA s equence. This can be
accomplished in vitro when DNA, appropriate enzymes , and cell extracts containing
heads and tails are combined. This process is called in vitro packaging.
Packaging limits of lambda particles are procured because the size of the inserted DNA
must fall between 78% and 105% of the wild-type pha ge DNA. If the distance
between cos sites is too small or too large, the resultant phages will not be viable. The
distance between the cos sites can be lengthened by inserting a segment of non-phage
passenger DNA.

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Plasmid ColE1 carries a gene for resistance to rifampicin (rif-r) and the cos sites of
phage lambda, which can be recognized by the cos-site-cutting (Ter) system of E. coli.
Cosmids such as this can function properly, provided that two cos sites are present and
the cos sites are separated by no less than 38 kb and no more than 54 kb. Cleavage of
ColE1 and foreign DNA by the restriction enzyme Hind III can be used to produce
linear, recombinant molecules. Transducing phage particles can be formed if the insert
between the two cos sites is 38–54 kb in length. No particles are produced if no insert is
made or if the insert is larger or smaller than that range. In vitro packaging (adding
heads and tails) forms transducing particles containing cosmids with cohesive termini.
Upon infection of a rifampicin-sensitive (Rif-s) cell with a transducing phage particle,
the linear chimera becomes circularized and replicates using the ColE1 replication
system. Plating cells on medium containing rifampicin selects for those cells containing
the rif-r gene, the ColE1 region, and a foreign insert.
Purification strategies of expressed His-tagged proteins:
Immobilized metal affinity chromatography (IMAC) is most frequently used for the
purification of polyhistidine (His-) tagged proteins. This technique is based on the
interaction between certain exposed protein residues (preferentially histidines) with
transition metal cations (Cu2+, Ni2+, Zn2+, Co2+). The transition metal itself is
immobilized to a cross-linked agarose matrix via a chelating group such as
iminodiacetic acid (IDA) or nitrilotriacetic acid (NTA).
Successful purification experiments of His-tagged proteins strongly depend on the
particular amino acid sequence, the protein conformation and the microenvironment
and location of the His-tag (C- or N-terminal). Furthermore, undesired co-purification of
non-specific host cell proteins is often observed, especially in the case of E.
coli expression systems.
IDA-immobilized Cu2+, Ni2+, Zn2+ and Co2+ions exhib it varying affinities &
specificities toward histidines. While Ni2+ ions show a high affinity but low specificity,
Co2+-ions are more specific but with reduced binding affinity and are therefore, an
option to reduce non-specific protein binding.

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Nickel and Cobalt IDA Agaroses, Fast Flow - Nickel-IDA Agarose, Fast Flow: 6 % Nickel-
IDA (Iminodiacetic acid (IDA)) Agarose is a superior, 6 % cross-linked immobilized metal
affinity chromatography (IMAC) resin that uses nickel ions for purifying recombinant
polyhistidine (His-) tagged proteins. Its unique properties facilitate the rapid and high-
yielding one-step purification from crude lysates both under denaturing and non-
denaturing conditions.
Nickel, Zinc, Cobalt, Copper and Metal Free IDA Aga roses, Gravity Flow - Nickel
Agaroses: His-tagged proteins are efficiently purified by a one-step procedure from
crude lysates both under denaturing and non-denaturing conditions.
ADVANCED CLONING STRATEGIES
Generalized overview of cloning strategies, with favoured routes shown by arrows.
Note that in cell-based cloning strategies, DNA fragments are initially generated
and cloned in a non-specific manner, so that screen ing for the desired clone is
carried out at the end of the process. Conversely, when specific DNA fragments

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are obtained by PCR or direct chemical synthesis, there is no need for subsequent
screening

Any cell-based cloning procedure has four essential parts: (i) a method for generating
the DNA fragment for cloning; (ii) a reaction that inserts that fragment into the chosen
cloning vector; (iii) a means for introducing that recombinant vector into a host cell
wherein it is replicated; and (iv) a method for selecting recipient cells that have acquired
the recombinant.
To simplify the description of such procedures, the assumption is made that we know
exactly what we are cloning. This is indeed the case with simple subcloning strategies,
where a defined restriction fragment is isolated from one cloning vector and inserted
into another. However, we also need to consider wha t happens in cases where the
source of donor DNA is very complex. We may wish, for example, to isolate a single gene
from the human genome, in which case the target seq uence could be diluted over a
millionfold by unwanted genomic DNA. We need to fin d some way of rapidly sifting
through, or screening, large numbers of unwanted sequences to identify our particular
target. There are two major strategies for isolating sequences from complex sources
such as genomic DNA. The first, a cell-based cloning strategy, is to divide the source

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DNA into manageable fragments and clone everything. Such a collection of clones,
representative of the entire starting population, is known as a library. We must then
screen the library to identify our clone of interest, using a procedure that discriminates
between the desired clone and all the others.
The second strategy is to selectively amplify the target sequence directly from the source
DNA using the polymerase chain reaction (PCR), and then clone this individual
fragment. Each strategy has its advantages and disadvantages. Note that, in the library
approach, screening is carried out after the entire source DNA population has been
cloned indiscriminately. Conversely, in the PCR approach, the screening step is built
into the first stage of the procedure, when the fragments are generated, so that only
selected fragments are actually cloned. In this chapter we consider principles for the
construction and screening of genomic and complemen tary DNA (cDNA) libraries, and
compare the library-based route of gene isolation to equivalent PCR-based techniques.
SYNTHESIS AND CLONING OF cDNA
There are two general stages. First DNA from some particular source is cut to liberate a
gene or other fragment of interest. This fragment is then “cloned” by inserting it into
another DNA macromolecule, known as a vector. After cloning, the chimeric DNA is
normally inserted into an appropriate host cell. A chimera is any hybrid molecule of
DNA, such as a vector plus a cloned gene, which has been engineered from two different
sources of DNA. We will first consider the enzymes used to cut and join fragments of
DNA.
Then we will discuss the vectors used in cloning and how DNA fragments are inserted
into them. Ultimately, cloned genes may be used in the manufacture of high levels of
recombinant protein or may be applied in gene therapy to cure inherited defects.
Nucleases Cut Nucleic Acids
Nucleases are enzymes that degrade nucleic acids. Ribonucleases (RNases) attack
RNA and deoxyribonucleases (DNases) attack DNA. Most nucleases are specific,
though the degree of specificity varies greatly. Some nucleases will only attack single
stranded nucleic acids, others will only attack double-stranded nucleic acids and a few
will attack either kind. Exonucleases attack at the end of nucleic acid molecules and
usually remove just a single nucleotide, or sometimes a short oligonucleotide. Any
particular exonuclease attacks either the 3¢-end or the 5¢-end but not both.

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Endonucleases cleave the nucleic acid chain in the middle. Some endonucleases are
non-specific, others, in particular the restriction enzymes, are extremely specific and
will only cut DNA after binding to specific recognition sequences. All these enzymes
have proved extremely useful both in genetic analysis and genetic engineering.
Restriction and Modification of DNA
Methylation of DNA by bacteria is used to distinguish the cell’s own DNA from DNA of
foreign origin. In nature, foreign DNA entering a bacterial cell would most likely be due
to virus infection.When viruses attack bacteria, the virus coat is left outside and only
the virus DNA enters the target cell. The virus DNA will take over the victim’s cellular
machinery and use it to manufacture more virus part icles unless the bacterial cell
fights back. The key is to degrade only foreign DNA without endangering the bacterial
cell’s own DNA. Bacteria make restriction and modification enzymes that respectively
cut and methylate DNA, ensuring than the foreign DN A is recognized and
destroyed.Whenever a bacterial cell makes a restriction enzyme, it also makes the
corresponding modification enzyme that modifies and protects its own DNA.The result
is that the DNA in a bacterial cell is immune to that cell’s own restriction enzymes. In
contrast, incoming, unmodified DNA will be degraded by the restriction enzyme.
Restriction enzymes are endonucleases that cut double-stranded DNA.They bind to
DNA at a specific sequence of bases, called the recognition site and then proceed to cut
the DNA. Modification enzymes bind to the DNA at the same recognition site as the
corresponding restriction enzymes and methylate the DNA. Modification enzymes
usually add the methyl group to adenine or cytosine within the recognition site.

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Restriction enzymes recognize non-methylated double -stranded DNA and cut it at
specific recognition sites. For example, EcoRI recognizes the sequence, 5’-
GAATTC-3’, and cuts after the G. Since this sequenc e is an inverted repeat, the
enzyme also cuts the other strand after the corresponding G, giving a zig-zag cut.
Modification enzymes are paired with restriction enzymes and recognize the same
sequence. Modification enzymes methylate the recogn ition sequence, which
prevents the restriction enzyme from cutting it.
Once methylated, DNA is protected from the restrict ion endonuclease. Only non-
methylated DNA will be cut and destroyed by the restriction enzyme. All the DNA in a
bacterial cell, including the chromosome and any plasmids, is normally protected by
modification.
Recognition of DNA by Restriction Endonucleases

Due to their ability to recognize specific sites in DNA, restriction endonucleases have
become one of the most widely used tools in genetic engineering. Restriction enzyme
recognition sites are usually four, six or eight bases long and the sequence forms an
inverted repeat. Thus the sequence on the top strand of the DNA is the same as the

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sequence of the bottom strand read in the reverse direction, as shown in above. Several
hundred different restriction enzymes are now known and each has its own specific
recognition site. Some recognition sites require a specific base at each position. Others
are less specific and may require only a purine or a pyrimidine at a particular position.
Some examples are shown in Table.
Since any random series of four bases will be found quite frequently, four base
recognizing enzymes cut DNA into many short fragments. Conversely, since any
particular eight-base sequence is less likely, the eight-base-recognizing enzymes cut
DNA only at longer intervals and generate fewer larger pieces.The six-base enzymes are
the most convenient in practice, as they give an intermediate result.
Naming of Restriction Enzymes
Restriction enzymes have names derived from the initials of the bacteria they come
from. The first letter of the genus name is capitalized and followed by the first two
letters of the species name (consequently, these three letters are in italics). The strain is
sometimes represented e.g. the R in EcoRI refers to Escherichia coli strain RY13.
The roman letter indicates the number of restriction enzymes found in the same
species. For example, Moraxella bovis has two different restriction enzymes called MboI
and MboII.
If two restriction enzymes from different species share the same recognition sequence
they are known as isoschizomers. Note that isoschizomers may not always cut in the
same place even though they bind the same base sequence. For example, the sequence
GGCGCC is recognized by four enzymes, each of which cuts in different places: NarI
(GG/CGCC), BbeI (GGCGC/C), EheI (GGC/GCC), and KasI (G/GCGCC).
Cutting of DNA by Restriction Enzymes
It might seem logical for the DNA to be cut at the recognition site where the restriction
enzyme binds.This is often true, but not always.The re are two major classes of
restriction enzyme that differ in where they cut the DNA, relative to the recognition site.
Type I restriction enzymes cut the DNA a thousand or more base pairs away from the
recognition site.This is done by looping the DNA around so that the enzyme binds both
at the recognition site and the cutting site.

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Type I restriction enzymes have three different sub units. The specificity subunit
recognizes a specific sequence in the DNA molecule. The modification subunit
adds a methyl group to the recognition site. If the DNA is nonmethylated, the
restriction subunit cuts the DNA, but at a different site, usually over 1000 base
pairs away. In the EcoK restriction enzyme, the subunits are HsdS, HsdM, and
HsdR
Since the exact length of the loop is not constant, and since the base sequence at the
cut site is not fixed, these enzymes are of little practical use to molecular biologists.
Even more bizarre is that type I restriction enzymes are suicidal. Most enzymes carry
out the same reaction over and over again on a continual stream of target molecules.
Each molecule of a type I restriction enzyme can cut DNA only a single time and then it
is inactivated!
Type I restriction systems consist of a single protein with three different subunits.
One subunit recognizes the DNA, other methylates the recognition sequence and the
third cuts the DNA at a distance from the recognition sequence. In type II restriction

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systems the restriction endonuclease and the methylase are two separate proteins that
operate independently but recognize the same DNA sequence.
Type II restriction enzymes cut the DNA in the middle of the recognition site.
Since the exact position of the cut is known, these are the restriction enzymes that are
normally used in genetic engineering.There are two different ways of cutting the
recognition site in half. One way is to cut both strands of the double stranded DNA at
the same point. This leaves blunt ends as shown.

HpaI is an blunt end restriction enzyme, that is, it cuts both strands of DNA in
exactly the same position. EcoRI is a sticky end restriction enzyme. The enzyme
cuts between the G and A on both strands, which gen erates an a four base pair
overhang on the ends of the DNA. Since these bases are free to base pair with any
complementary sequence, they are considered “sticky ”.
The alternative is to cut the two strands in differ ent places, which generates
overhanging ends. The ends made by such a staggered cut will base pair with each
other and consequently they are known as sticky ends.
Enzymes that generate sticky ends are the most useful. If two different pieces of DNA
are cut with the same restriction enzyme or enzymes that generate the same overhang,
the same sticky ends are generated.This allows fragments of DNA from two different
original DNA molecules to be bound together by matching the sticky ends.
BamHI and BglII generate the same overhanging or sticky ends. BamHI recognizes
the sequence 5’- GGATCC-3’ and cuts after the first 5’G, which generates the 3’-
CTAG- 5’ overhang on the bottom strand. BglII recognizes the sequence 5’-
AGATCT-3’ and cuts after the first 5’ A, which generates a 5’-GATC-3’ overhang on

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the top strand. If these two pieces are allowed to anneal, the complementary
sequences will hydrogen bond together, allowing the nicks to be sealed more
easily by DNA ligase


Such pairing is temporary since the pieces of DNA are only held together by hydrogen
bonding between the base pairs, not by permanent co valent bonds. Nonetheless, this
assists the permanent bonding of the sugar-phosphate backbone by DNA ligase.When
two sticky ends made by the same enzyme are ligated, the junction may be cut apart
later by using the same enzyme again. However, if t wo sticky ends made by two
different enzymes are ligated together, a hybrid site is formed that cannot be cut by
either enzyme.
DNA Fragments are joined by DNA Ligase
The enzyme DNA ligase is used to join DNA fragments covalently. DNA ligase operates
during DNA replication where it joins up the fragments of the lagging strand. If DNA
ligase finds two DNA fragments touching each other end to end, it will ligate them
together.
In practice, segments of DNA with matching sticky ends will tend to stay attached much
of the time and consequently DNA ligase will join them efficiently. Since DNA fragments
with blunt ends have no way to bind each other, they drift apart most of the time.
Ligating blunt ends is very slow and requires a high concentration of DNA ligase, as well
as, a high concentration of DNA. In fact, bacterial ligase cannot join blunt ends at all. In
practice, T4 ligase is normally used in genetic engineering as it can join blunt ends if

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need be. T4 ligase originally came from bacteriophage T4, although nowadays it is
manufactured by expressing the gene that encodes it in E. coli.
T4 DNA ligase connects the sugarphosphate backbone of two pieces of DNA. In the
example, overlapping sticky ends connect a double s tranded piece of DNA, but the
backbone of each strand has not been connected. T4 DNA ligase recognizes these
nicks or breaks in the backbone and uses energy fro m the hydrolysis of ATP to
drive the ligation reaction.


Making a Restriction Map
A diagram that shows the location of restriction enzyme cut sites on a segment of

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DNA is known as a restriction map. The first step in generating such a map is to digest
the DNA with a series of restriction enzymes, one at a time. The fragments of digested
DNA are separated by agarose gel electrophoresis.
Comparison with appropriate standards allows the si zes of the fragments to be
estimated. This reveals how many recognition sites each enzyme has in the DNA and
their distances apart.What remains unknown is the relative order of the fragments.
For example, suppose we start with a 5,000 base pair (bp) piece of DNA that is cut twice
by the restriction enzyme BamHI giving three fragments of 3,000 bp, 1,500 bp, and 500
bp. There are three alternative arrangements for three fragments.
You might think there should be six possible arrang ements, but the other three
theoretical arrangements are merely the first three, turned back to front, they are not
genuinely different physical molecules.
Fig.shows the backward arrangement just for fragment number III. To decide which of
the three possible arrangements are correct, double digests using two restriction
enzymes must be performed.
The DNA is cut with each enzyme alone and with both simultaneously (Fig. 22.07B).
The results of gel electrophoresis for the two single digests and the double digest are
shown. Suppose that the second restriction enzyme is EcoRI and that alone it cuts just
once to give two fragments of 4,000 bp and 1,000 bp. Thus, for EcoRI alone there is only
one possible arrangement. In the double digest, the largest fragment seen in the BamHI
lane has disappeared. This means that there is an EcoRI cut site within this 3,000 bp
BamHI fragment. Since, in this example, there is only one EcoRI cut site, only one of the
BamHI fragments disappears in the double digest.This a llows us to reduce the
possibilities to the restriction maps shown in Figure.
Note that two alternatives remain.To decides between these needs a third enzyme.
Double digests with BamHI plus enzyme III and of Ec oRI plus enzyme III would be
analyzed, as above. Eventually, this approach allows the construction of a complete
restriction map of any segment of DNA. This may then be used as a guide to further
manipulations.

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(A) To determine the location and number of restric tion enzyme sites, a segment
of DNA is digested with a restriction enzyme. In this example, the piece of DNA is
5,000 base pairs in length. Cutting with BamHI gave three fragments: of 3,000 bp,
1500 bp, and 500 bp. The figure shows the three pos sible arrangements of these
three fragments. The fourth arrangement shown is no t really different but is
merely the third possible arrangement drawn in the opposite orientation.
(B) Double digestion is the next step in compiling a restriction map. Cutting the
DNA with EcoRI alone would give two fragments: of 4000 bp and 1 000 bp. When
the DNA is cut with both EcoRI and BamHI simultaneously, four fragments are
resolved by gel electrophoresis. Two of these are identical to the 1500 bp and 500
bp fragments from the BamHI single digest; therefore, no EcoRI sites are present
within these fragments. The remaining two fragments , 2000 bp and 1000 bp, add
up to give the 3000 bp BamHI fragment. Therefore, the single EcoRI site must be
within the 3000 bp BamHI fragment. Of the three possible arrangements sho wn in
part (A), the third arrangement is ruled out (if it was cut with EcoRI alone, it could
not give two fragments of 4000 bp and 1000 bp).

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PCR AMPLIFIED DNA
Cloning Substrate or Diagnostic Tool
PCR is useful because it allows identifying and isolating one or a few copies of a specific
target sequence from a complex background. This per mits rapid cloning of known
sequences without the preparation of a library for gene isolation. PCR is also useful for
screening for the presence of pathogens, even when only a small amount of sequence
information is available for the pathogen. Production of amplified product using
pathogen- specific primers signals the presence of the pathogen in the sample
.
USE OF ADAPTORS AND LINKERS

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It may be the case that the restriction enzyme used to generate the cohesive ends in the
linker will also cut the foreign DNA at internal sites. In this situation, the foreign DNA
will be cloned as two or more subfragments.
One solution to this problem is to choose another restriction enzyme, but there may not
be a suitable choice if the foreign DNA is large and has sites for several restriction
enzymes. Another solution is to methylate internal restriction sites with the appropriate
modification methylase.
An example of this is described in Fig.
Alternatively, a general solution to the problem is provided by chemically synthesized
adaptor molecules which have a preformed cohesive end (Wu et al. 1978). Consider a
blunt-ended foreign DNA containing an internal BamHI site, which is to be cloned in a
BamHl-cut vector. The Bam adaptor molecule has one blunt end bearing a 5′ phosphate
group and a Bam cohesive end which is not phosphorylated. The adaptor can be ligated
to the foreign DNA ends. The foreign DNA plus added adaptors is then phosphory lated
at the 5′ termini and ligated into the BamHI site of the vector. If the foreign DNA were to
be recovered from the recombinant with BamHI, it would be obtained in two fragments.
However, the adaptor is designed to contain two other restriction sites (SmaI, HpaII),
which may enable the foreign DNA to be recovered intact.
Note that the only difference between an adaptor and a linker is that the former has
cohesive ends and the latter has blunt ends. A wide range of adaptors are available
commercially.
Use of a BamHI adaptor molecule.
A synthetic adaptor molecule is ligated to the foreign DNA. The adaptor is used in
the 5′-hydroxyl form to prevent self-polymerization. The foreign DNA plus ligated
adaptors is phosphorylated at the 5′-termini and ligated into the vector previously
cut with BamHI.

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HOMOPOLYMERS TAILING IN cDNA CLONING
A general method for joining DNA molecules makes us e of the annealing of
complementary homopolymer sequences. Thus, by addin g oligo(dA) sequences to the 3′
ends of one population of DNA molecules and oligo(dT) blocks to the 3′ ends of another
population, the two types of molecule can anneal to form mixed dimeric circles.
An enzyme purified from calf thymus, terminal deoxynucleotidyltransferase, provides
the means by which the homopolymeric extensions can be synthesized, for if presented
with a single deoxynucleotide triphosphate it will repeatedly add nucleotides to the 3′
OH termini of a population of DNA molecules (Chang & Bollum 1971). DNA with
exposed 3′ OH groups, such as arise from pretreatment with phage λ exonuclease or
restriction with an enzyme such as PstI, is a very good substrate for the transferase.
However, conditions have been found in which the enzyme will extend even the shielded
3′ OH of 5′ cohesive termini generated by EcoRI (Roychoudhury et al. 1976, Humphries
et al. 1978).

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Use of calf-thymus terminal deoxynucleotidyltransfe rase to add complementary
homopolymer tails to two DNA molecules.
The terminal transferase reactions have been characterized in detail with regard to their
use in gene manipulation (Deng & Wu 1981, Michelson & Orkin 1982). Typically, 10–40
homopolymeric residues are added to each end.
One of the earliest examples of the construction of recombinant molecules, the insertion
of a piece of λ DNA into SV40 viral DNA, made use of homopolymer tailing ( Jackson et
al. 1972). In their experiments, the single-stranded gaps which remained in the two
strands at each join were repaired in vitro with DNA polymerase and DNA ligase so as to
produce covalent-ly closed circular molecules. The recombinants were then transfected
into susceptible mammalian cells.
Subsequently, the homopolymer method, using either dA.dT or dG.dC homopolymers
was used extensively to construct recombinant plasmids for cloning in E. coli. In recent
years, homopolymer tailing has been largely replaced as a result of the availability of a
much wider range of restriction endonucleases and o ther DNA-modifying enzymes.
However, it is still important for cDNA cloning.

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EXPRESSION OF CLONED DNA MOLECULE.
When gene manipulation in fungi first became possible, there were many unsuccessful
attempts to express heterologous genes from bacteri a or higher eukaryotes. This
suggested that fungal promoters have a unique structure, a feature first shown for S.
cerevisiae (Guarente 1987). Four structural elements can be recognized in the average
yeast promoter.
First, several consensus sequences are found at the transcription-initiation site.
Two of these sequences, TC (G/A) A and PuPuPyPuPu, account for more than half of the
known yeast initiation sites (Hahn et al. 1985, Rudolph & Hinnen 1987).
These sequences are not found at transcriptioninitiation sites in higher eukaryotes,
which imply a mechanistic difference in their transcription machinery compared with
yeast.

The second motif in the yeast promoter is the TATA box (Dobson et al. 1982). This is an
AT-rich region with the canonical sequence TATAT/AAT/A, located 60–120 nucleotides
before the initiation site. Functionally, it can be considered equivalent to the Pribnow
box of E. coli promoters. The third and fourth structural elements are upstream
activating sequences (UASs) and upstream repressing sequences (URSs). These are
found in genes whose transcription is regulated. Binding of positive-control proteins to
UASs increases the rate of transcription and deleti on of the UASs abolishes
transcription. An important structural feature of UASs is the presence of one or more
regions of dyad symmetry (Rudolph & Hinnen 1987).
Binding of negative-control proteins to URSs reduces the transcription rate of those
genes that need to be negatively regulated. The level of transcription can be affected by
sequences located within the gene itself and which are referred to as downstream
activating sequences (DASs). Chen et al. (1984) noted that, when using the

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phosphoglycerate kinase (PGK) promoter, several heterologous proteins accumulate to
1–2% of total cell protein, whereas phosphoglycerate kinase itself accumulates to over
50%.
These disappointing amounts of heterologous protein reflect the levels of mRNA which
were due to a lower level of initiation rather than a reduced mRNA half-life (Mellor et al.
1987). Addition of downstream PGK sequences restored the rate of mRNA transcription,
indicating the presence of a DAS. Evidence for these DASs has been found in a number
of other genes
SELECTION, SCREENING AND ANALYSIS OF RECOMBINANT GE NETIC SELECTION
Steps in molecular cloning
The conventional restriction and ligation cloning protocol involves four major steps
namely fragmentation of DNA with restriction endonucleases, ligation of DNA fragments
to a plasmid vector, introduction into bacterial cells by transformation and screening
and selection of recombinants.
Selection and preparation of vector and insert
A cloning vehicle, also termed as a vector, can be classified as a carrier carrying a gene
to be transferred from one organism to another. Other cloning vectors include plasmids,
cosmids, bacteriophage, phagemids and artificial chromosomes. In the early days of
producing proteins in E. coli, limitations to transcription initiation were believed to lead
to lower protein expression levels (Gralla, 1990). This event resulted in efforts put into
construction of expression vectors, which carried strong promoters to enhance Mrna
yield and a stable mRNA eventually. The promoters used included phage promoters like
T7 and T5, the synthetic promoters tac and trc, and the arabinose inducible araBAD
(Trepe, 2006). Additional vectors that were made available included Lambda promoters,
PR and PL, (Elvin et al., 1990), rhamnose promoter (Cardona & Valvano, 2005), Trp-lac
promoter (Chernajovskyi et al., 1983) etc. Certain promoter variants as seen in the
expression vector pAES25 yield the maximum level of soluble active target protein
(Broedel & Papciak , 2007).
Downstream of each specific promoter, there is a multiple cloning site (MCS) for cloning
the gene to be expressed. While the inducible promoters are used to drive the foreign
gene expression, the constitutive promoters (Liang et al,., 1999) are used mainly to
express the antibiotic expression marker genes for plasmid maintenance.

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TA cloning vectors (Zhou & Gomez-Sanchez, 2000; Che n et al., 2009) that takes
advantage of the well-known propensity of non-proofreading DNA polymerases (e.g.,
Taq, Tfl, Tth) to add a single 3´-A to PCR products are also employed for cloning large
PCR fragments. The proof-reading polymerases lack 5'-3' proofreading activity and are
capable of adding adenosine triphosphate residues to the 3' ends of the double stranded
PCR product. Such a PCR amplified product can then be cloned in any linearized vector
with complementary 3' T overhangs.
The GC cloning technology is based on the recent di scovery that the above proof-
reading enzymes similarly add a single 3´-G to DNA molecules, either during PCR or as
a separate G-tailing reaction to any blunt DNA. GC cloning vectors pSMART® GC and
pGC™ Blue (commercialized by Lucigen, USA) contain a single 3´-C overhang, which is
compatible with the single 3´-G overhang on the inserts. Mead and coworkers (Mead et
al., 1991) report cloning of PCR products without a ny restriction digestion taking
advantage of the single 3' deoxyadenylate extension that Thermus aquaticus, Thermus
flavus, and Thermococcus litoralis DNA polymerases add to the termini of amplified
nucleic acid.
Gateway cloning system is a relatively new trend in the field of molecular cloning, where
in the site specific recombination system of lambda phage is used (Katzen 2007). This
system enables the researchers to efficiently transfer DNA fragments between different
vector and expression systems, without changing the orientation of the gene or its
reading frame. The specific sequences are called “Gateway att sites” and recombination
is facilitated by two enzymes “LR clonase” and “BP clonase”. This easy Ligase-free
cloning system is very beneficial for cloning, combining and transferring of DNA
segments between different expression platforms in a high-throughput manner, but
making the gateway entry clone usually involves conventional restriction enzyme based
cloning, and this is a major drawback of this system.
DNA vectors that are used in many molecular biology gene cloning experiments need
not necessarily result in protein expression. Expression vectors are often specifically
designed to contain regulatory sequences that act as enhancer and promoter regions,
and lead to efficient transcription of the gene that is carried on the expression vector.
The regularly used cloning cum expression vectors include pET vectors, pBAD vectors,
pTrc vectors etc wherein the GOI is cloned with a suitable promoter of the vector using

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the start codon of the vector or using a gene of interest with its own start codon into an
apopropriate restriction site in the MCS.
RNA polymerases are enzyme complexes that synthesize RNA molecules using DNA as a
template. The transcription begins when RNA polymerase binds to the DNA double helix
which is at a promoter site just upstream of the gene to be transcribed. While in
prokaryotes, one DNA-dependent RNA polymerase trans cribes all classes of DNA
molecules and the core Escherichia coli enzyme called E. coli RNA polymerase consists of
three types of subunit, α, β, and β′, and has the composition α
2β β′; the holoenzyme
contains an additional σ subunit or sigma factor (A aron, 2001). The phage RNA
polymerase like T7 RNA polymerase found in pET base d expression vectors are much
smaller and simpler than bacterial ones: the polymerases from phage T3 and T7 RNA,
e.g., are single polypeptide chains of <100 kDa.
The DNA fragment to be cloned is first isolated by a number of ways like cDNA
preparation, nuclease fragments of genomic DNA, syn thetic DNA’s, amplified DNA
fragments by means of polymerase chain reaction. After appropriate restriction enzyme
digestion and purification, the purified inserts are ligated to the vector of choice.
INSERTIONAL INACTIVATION
Once a gene or other fragment of DNA has been clone d into a plasmid vector and
transformed into a bacterial cell, we face the problem of detecting its presence. The
plasmid itself may be detected by conferring antibiotic resistance on the host cell, but
this leaves the question of whether the presumed insert is actually there. If the cloned
gene itself codes for a product that is easy to detect, there is no problem. In most cases,
however, the presence of the inserted DNA itself must be directly monitored. The least
sophisticated and most tedious method is to screen a large number of suspects for the
inserted DNA. Many separate bacterial colonies that received the plasmid vector,
hopefully with DNA inserted, are grown in separate vials.
Plasmid DNA is extracted from each of these bacteri al cultures and cut with the
restriction enzyme used in the original cloning experiment. If there is no insert in the
plasmid, this merely converts the plasmid from a circular to a linear molecule of DNA. If
the vector contains inserted DNA, two pieces of DNA are produced, one being the
original plasmid and the other the inserted DNA fragment.To see how many fragments
of DNA are present, and the cut DNA is separated by agarose gel electrophoresis. If

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enough transformed colonies are tested, sooner or later one carrying a plasmid with the
inserted DNA fragment will be found.This approach was of necessity used in the early
days of genetic engineering.
Today, modified vectors are available that facilitate screening by a variety of
approaches. Rather less laborious is to use a plasmid with two antibiotic resistance
genes. One antibiotic resistance gene is used to select for cells, which have received the
plasmid vector itself. The second is used for the insertion and detection of cloned DNA.

A plasmid that has a unique restriction enzyme site within an antibiotic
resistance gene can be used to identify those plasm ids into which a cloned gene
has been inserted successfully. If the gene of inte rest is ligated into this
restriction site, the antibiotic resistance gene wi ll no longer be active. Any
bacteria harboring the plasmid with an insert will no longer be resistant to this
particular antibiotic.
The cut site for the restriction enzyme used must lie within this second antibiotic
resistance gene. When the cloned fragment of DNA is inserted this antibiotic resistance
gene will be disrupted. This is referred to as insertional inactivation.
Consequently, cells that receive a plasmid without an insert will be resistant to both
antibiotics. Those receiving a plasmid with an insert will be resistant to only the first
antibiotic. The most convenient and widely used method to screen for inserts uses color
screening.The most common procedure uses b -galactosidase and X-gal to produce
bacterial colonies that change color when an insert is present within the vector. The
process, called blue/white screening, has a unique vector that carries the 5¢-end of

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the lacZ gene. This truncated gene encodes the alpha fragment of b-galactosidase,
which consists of the N-terminal region or first 146 amino acids.A specialized bacterial
host strain is required whose chromosome carries a lacZ gene missing the front portion
but encodes the rest of the b-galactosidase protein. If the plasmid and chromosomal
gene segments are active they produce two protein fragments that associate to give an
active enzyme. This is referred to as alpha complementation.
CHROMOGENIC SUBSTRATES
Chromogenic substrates have been widely used for ma ny years and offer the simplest
and most cost-effective method of detection. These substrates are divided into two
groups based on the nature of the product of the en zyme-substrate reaction.
When precipitating substrates come in contact with the appropriate enzyme, they are
converted to insoluble products that precipitate onto the sample or membrane. Because
of the insoluble nature of these products, precipitating substrates are commonly used
for IHC and Western blotting. Conversely, soluble s ubstrates form water-soluble
coloured products that dissolve into the test solution and are commonly used in ELISA
assays.
Besides the difference in the nature of their products, these two types of chromogenic
substrates differ in the detection instrumentation used. Precipitating substrates require
no more specialized equipment than a light microscope to detect the presence, intensity
and localization of the insoluble product, while a microplate reader is used to measure
the absorbance and therefore the amount of soluble product in solution. Chromogenic
blotting substrates are available in a wide variety of specifications and formats, and the
appropriate substrate choice depends on the enzyme label, desired sensitivity and the
form of the signal or method of detection needed.
For either type of chromogenic substrate, protein detection is similar to developing film;
the sample is incubated in substrate until the color development is satisfactory, after
which the reaction is halted and/or the amount of product measured. This approach
allows the user to control the resolution of the data. The low sensitivity of chromogenic
substrates, though, makes optimization of detecting low-abundance proteins difficult;
although the reaction can be developed for several hours or even overnight, this also
allows background signal to develop as well. Where chromogenic substrates fail in
terms of sensitivity, they are ideal for applications where protein abundance is high.

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Chromogenic substates are typically used to detect abundant proteins, and reaction
development can be monitored visually; this allows greater flexibilty for optimization
compared to chemiluminescent or fluorescent blotting systems.
Chemiluminescent Substrates:
Chemiluminescent substrates are popular because the y offer several advantages over
other detection methods, including a large linear response range that allows the
detection and quantitation of a wide range of protein concentrations. Luminol is one of
the most commonly used chemiluminescent reagents; luminol is oxidized by peroxide to
form the excited-state 3-aminophthalate, which decays to a lower energy state by
releasing photons of light at a wavelength of 425nm.
Chemiluminescent substrates differ from other substrates in that the light detected is a
transient product of the reaction that is only present while the enzyme-substrate
reaction is occurring. This is in contrast to chromogenic substrates that produce a
stable, coloured product. In a chemiluminescent reaction, the substrate is the limiting
reagent in the reaction; as it is exhausted, light production decreases and eventually
ceases. A well-optimized procedure using the proper enzyme conjugate dilutions will
produce a stable output of light for several hours, allowing consistent and sensitive
detection of proteins.
Chemiluminescent substrates are commonly used for the short-term, rapid detection of
protein detection by Western blot analysis using x-ray film, phosphorimagers or CCD
cameras.
Fluorescent Substrates
While fluorophore-tagged antibodies and other molecules are more commonly used to
detect target proteins, fluorescent substrates are also available for enzymatic detection.
These substrates remain non-fluorescent or emit low fluorescence until metabolized by
the enzyme probe, after which they emit intense fluorescence. These substrates offer
greater sensitivity and the ability for rapid quantitation because of fluorescent
microscopic imaging, fluorescent microplate readers and analytical software.
COMPLEMENTATION OF DEFINED MUTATIONS
Once a gene or other fragment of DNA has been clone d into a plasmid vector and
transformed into a bacterial cell, we face the problem of detecting its presence. The
plasmid itself may be detected by conferring antibiotic resistance on the host cell, but

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this leaves the question of whether the presumed insert is actually there. If the cloned
gene itself codes for a product that is easy to detect, there is no problem. In most cases,
however, the presence of the inserted DNA itself must be directly monitored.
The most convenient and widely used method to screen for inserts uses color screening.
The most common procedure uses b -galactosidase and X-gal to produce bacterial
colonies that change color when an insert is present within the vector. The process,
called blue/white screening, has a unique vector that carries the 5¢-end of the lacZ
gene. This truncated gene encodes the alpha fragment of b-galactosidase, which
consists of the N-terminal region or first 146 amino acids.A specialized bacterial host
strain is required whose chromosome carries a lacZ gene missing the front portion but
encodes the rest of the b-galactosidase protein. If the plasmid and chromosomal gene
segments are active they produce two protein fragments that associate to give an active
enzyme. This is referred to as alpha complementation.

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Eliminating Unwanted Restriction Sites - The restriction site shown in blue is
unwanted. During normal DNA replication, occasional mutations occur.
Consequently a very small percentage of the plasmid s will carry a random
mutation (red) that alters this particular restriction recognition sequence. A
sample of the plasmid DNA is isolated from a bacterial culture. The plasmids are
treated with the appropriate restriction enzyme. All will be cut, except those with
mutant restriction sites. The mixture is then transformed back into bacterial
cells. Bacteria receiving cut, linearized plasmids will degrade them. Only mutant
plasmids which remain circular will survive.
Assembling an active protein from fragments made separately is normally not possible.
Fortunately, b-galactosidase is exceptional in this respect. The reason for splitting lacZ
between plasmid and host is that the lacZ gene is unusually large (approximately 3000
bp—almost as large as many small plasmids) and it greatly helps if cloning plasmids are
small.
In order to utilize this unique protein for cloning, a polylinker is inserted into the lacZa
coding sequence on the plasmid, very close to the front of the gene. Luckily, the very
front most part of the b-galactosidase protein is inessential for enzyme activity. As long
as the polylinker is inserted without disrupting the reading frame, the small addition
does not affect the enzyme. However, if a foreign segment of DNA is inserted into the
polylinker, the alpha fragment of b-galactosidase is disrupted and no active enzyme can
form.
Alpha complementation - The b-galactosidase protein is unique since it can be
expressed as two pieces that come together to form a functional protein. The two
protein fragments can be encoded on two different m olecules of DNA within the
bacterial cell. The alpha fragment can be expressed from a plasmid and the
remainder of the b-galactosidase can be expressed from the chromosome

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NUCLEIC ACID HYBRIDIZATION
Nucleic acid hybridization is the most commonly use d method of library screening
because it is rapid, it can be applied to very large numbers of clones and, in the case of
cDNA libraries, can be used to identify clones that are not full-length (and therefore
cannot be expressed).
Grunstein and Hogness (1975) developed a screening procedure to detect DNA
sequences in transformed colonies by hybridization in situ with radioactive RNA probes.
Their procedure can rapidly determine which colony among thousands contains the
target sequence. A modification of the method allows screening of colonies plated at a
very high density (Hanahan & Meselson 1980). The co lonies to be screened are first
replica-plated on to a nitrocellulose filter disc that has been placed on the surface of an
agar plate prior to inoculation (Fig.).
A reference set of these colonies on the master plate is retained. The filter bearing the
colonies is removed and treated with alkali so that the bacterial colonies are lysed and
the DNA they contain is denatured. The filter is then treated with proteinase K to
remove protein and leave denatured DNA bound to the nitrocellulose, for which it has a
high affinity, in the form of a ‘DNA print’ of the colonies. The DNA is fixed firmly by
baking the filter at 80°C. The defining, labelled RNA is hybridized to this DNA and the
result of this hybridization is monitored by autoradiography. A colony whose DNA print
gives a positive autoradiographic result can then be picked from the reference plate.

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Variations of this procedure can be applied to phage plaques (Jones & Murray 1975,
Kramer et al. 1976). Benton and Davis (1977) devised a method cal led plaque lift, in
which the nitrocellulose filter is applied to the upper surface of agar plates, making
direct contact between plaques and filter. The plaques contain phage particles, as well
as a considerable amount of unpackaged recombinant DNA. Both phage and
unpackaged DNA bind to the filter and can be denatu red, fixed and hybridized. This
method has the advantage that several identical DNA prints can easily be made from a
single-phage plate: this allows the screening to be performed in duplicate, and hence
with increased reliability, and also allows a single set of recombinants to be screened
with two or more probes. The Benton and Davis (1977) procedure is probably the most
widely applied method of library screening, success fully applied in thousands of
laboratories to the isolation of recombinant phage by nucleic acid hybridization (Fig.).

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More recently, however, library presentation and screening have become increasingly
automated. In place of RNA probes, DNA or synthetic oligonucleotide probes can be
used. A number of alternative labelling methods are also available that avoid the use of
radioactivity. These methods involve the incorporation of chemical labels into the probe,
such as digoxigenin or biotin, which can be detected with a specific antibody or the
ligand streptavidin, respectively.
Probe design
A great advantage of hybridization for library screening is that it is extremely versatile.
Conditions can be used in which hybridization is very stringent, so that only sequences
identical to the probe are identified. This is necessary, for example, to identify genomic
clones corresponding to a specific cDNA or to ident ify overlapping clones in a
chromosome walk.

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Alternatively, less stringent conditions can be used to identify both identical and related
sequences. This is appropriate where a probe from one species is being used to isolate a
homologous clone from another species (e.g. see Old et al. 1982). Probes corresponding
to a conserved functional domain of a gene may also cross-hybridize with several
different clones in the same species at lower stringency, and this can be exploited to
identify members of a gene family. The identification of the vertebrate Hox genes
provides an example in which cross-species hybridization was used to identify a family
of related clones (Levine et al. 1984). In this case a DNA sequence was identified that
was conserved between the Drosophila developmental genes fushi tarazu and
Antennapedia. When this sequence, the homoeobox, was used to screen a Southern blot
of DNA from other species, including frogs and humans, several hybridizing bands were
revealed. This led to the isolation of a number of clones from vertebrate cDNA libraries
representing the large family of Hox genes that play a central role in animal
development.
Hybridization thus has the potential to isolate any sequence from any library if a probe
is available. If a suitable DNA or RNA probe cannot be obtained from an existing cloned
DNA, an alternative strategy is to make an oligonucleotide probe by chemical synthesis.
This requires some knowledge of the amino acid sequence of the protein encoded by the
target clone. However, since the genetic code is degenerate (i.e. most amino acids are
specified by more than one codon), degeneracy must be incorporated into probe design
so that a mixture of probes is made, at least one variant of which will specifically match
the target clone. Amino acid sequences known to include methionine and tryptophan
are particularly valuable because these amino acids are each specified by a single
codon, hence reducing the degeneracy of the resulting probe. Thus, for example, the
oligopeptide His-Phe-Pro-Phe-Met may be identified and chosen to provide a probe
sequence, in which case 32 different oligonucleotides would be required:

These 32 different sequences do not have to be synthesized individually because it is
possible to perform a mixed addition reaction for each polymerization step. This mixture
is then end-labelled with a single isotopic or alternatively labelled nucleotide, using an

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exchange reaction. This mixed-probe method was originally devised by Wallace and co-
workers (Suggs et al. 1981). To cover all codon possibilities, degeneracies of 64-fold
(Orkin et al. 1983) or even 256-fold (Bell et al. 1984) have been employed successfully.
What length of oligonucleotide is required for reliable hybridization? Even though 11-
mers can be adequate for Southern blot hybridization (Singer- Sam et al. 1983), longer
probes are necessary for good colony and plaque hyb ridization. Mixed probes of 14
nucleotides have been successful, although 16-mers are typical (Singer-Sam et al.
1983). An alternative strategy is to use a single longer probe of 40–60 nucleotides. Here
the uncertainty at each codon is largely ignored and instead increased probe length
confers specificity. Such probes are usually designed to incorporate the most commonly
used codons in the target species, and they may include the non-standard base inosine
at positions of high uncertainty because this can pair with all four conventional bases.
Such probes are sometimes termed guessmers. Hybridization is carried out under low
stringency to allow for the presence of mismatches. This strategy is examined
theoretically by Lathe (1985) and has been applied to sequences coding for human
coagulation factor VIII (Toole et al. 1984, Wood et al. 1984) and the human insulin
receptor (Ullrich et al. 1985).
Chromosome walking
Chromosome walking utilizes overlapping fragments o f a particular chromosome to
isolate genes upstream and downstream from the original DNA fragment. The first step
is to identify the region of the chromosome to which the probe hybridizes. In this
example, probe #1 hybridizes to the purple region o f the chromosome. When the
chromosome is cut with restriction enzyme #1, fragment 1A will hybridize to probe #1 at
one end. This allows fragment 1A to be isolated and sequenced and its downstream
sequence is used to generate probe #2. To find the next segment of the chromosome, a
different restriction enzyme is used. This time probe #2 will hybridize to fragment 2B.
Once again the probe recognizes only the first half of this fragment. The downstream
sequence of fragment 2B can then be determined, and this information can be used to
make probe #3. Next, the chromosome is cut with res triction enzyme #1 again. Now
probe #3 will hybridize with fragment 1B, whose downstream sequence can therefore be
determined, and another probe, called probe 4 can be made. This procedure can be
continued as far as desired, working in either direction.

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SCREENING METHODS FOR CLONED LIBRARIES
The identification of a specific clone from a DNA library can be carried out by exploiting
either the sequence of the clone or the structure/function of its expressed product. The
former applies to any type of library, genomic or cDNA, and can involve either nucleic
acid hybridization or the PCR. In each case, the design of the probe or primers can be
used to home in on one specific clone or a group of structurally related clones. Note that
PCR screening can also be used to isolate DNA sequences from uncloned genomic DNA
and cDNA. Screening the product of a clone applies only to expression libraries, i.e.
libraries where the DNA fragment is expressed to yield a protein. In this case, the clone
can be identified because its product is recognized by an antibody or a ligand of some
nature, or because the biological activity of the protein is preserved and can be assayed
in an appropriate test system.

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PCR SCREENING PROTOCOLS
The PCR is widely used to isolate specific DNA sequences from uncloned genomic DNA
or cDNA, but it also a useful technique for library screening (Takumi, 1997). As a
screening method, PCR has the same versatility as h ybridization, and the same
limitations. It is possible to identify any clone by PCR but only if there is sufficient
information about its sequence to make suitable primers.* To isolate a specific clone,
PCR is carried out with gene-specific primers that flank a unique sequence in the
target. A typical strategy for library screening by PCR is demonstrated by Takumi and
Lodish (1994). Instead of plating the library out on agar, as would be necessary for
screening by hybridization, pools of clones are maintained in multiwell plates. Each well
is screened by PCR and positive wells are identified. The clones in each positive well are
then diluted into a series in a secondary set of plates and screened again. The process
is repeated until wells carrying homogeneous clones corresponding to the gene of
interest have been identified. There are also several applications where the use of
degenerate primers is favourable. A degenerate primer is a mixture of primers, all of
similar sequence but with variations at one or more positions. This is analogous to the
use of degenerate oligonucleotides as hybridization probes, and the primers are
synthesized in the same way. A common circumstance requiring the use of degenerate
primers is when the primer sequences have to be deduced from amino acid sequences
(Lee et al. 1988). Degenerate primers may also be employed to search for novel members
of a known family of genes (Wilks 1989) or to search for homologous genes between
species (Nunberg et al. 1989). As with oligonucleotide probes, the selection of amino
acids with low codon degeneracy is desirable. However, a 128-fold degeneracy in each
primer can be successful in amplifying a single-copy target from the human genome
(Girgis et al. 1988). Under such circumstances, the concentration of any individual
primer sequence is very low, so mismatching between primer and template must occur
under the annealing conditions chosen. Since mismatching of the 3′-terminal nucleotide
of the primer may prevent efficient extension, degeneracy at this position is to be
avoided.
IMMUNOLOGICAL SCREENING
Immunological screening involves the use of antibod ies that specifically recognize
antigenic determinants on the polypeptide synthesized by a target clone. This is one of

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the most versatile expressioncloning strategies, because it can be applied to any protein
for which an antibody is available. Unlike the screening strategies discussed below,
there is also no need for that protein to be functional. The molecular target for
recognition is generally an epitope, a short sequence of amino acids that folds into a
particular three-dimensional conformation on the surface of the protein. Epitopes can
fold independently of the rest of the protein and therefore often form even when the
polypeptide chain is incomplete or when expressed as a fusion with another protein.
Importantly, many epitopes can form under denaturin g conditions, when the overall
conformation of the protein is abnormal. The first immunological screening techniques
were developed in the late 1970s, when expression libraries were generally constructed
using plasmid vectors.
The method of Broome and Gilbert (1978) was widely used at the time. This method
exploited the fact that antibodies adsorb very strongly to certain types of plastic, such
as polyvinyl, and that IgG antibodies can be readily labelled with
125
I by iodination in
vitro. As usual, transformed cells were plated out on Petri dishes and allowed to form
colonies. In order to release the antigen from positive clones, the colonies were lysed,
e.g. using chloroform vapour or by spraying with an aerosol of virulent phage (a replica
plate is required because this procedure kills the bacteria). A sheet of polyvinyl that had
been coated with the appropriate antibody was then applied to the surface of the plate,
allowing antigen–antibody complexes to form. The sheet was then removed and exposed
to
125
I-labelled IgG specific to a different determinant on the surface of the antigen (i.e. a
determinant not involved in the initial binding of the antigen to the antibody-coated
sheet.

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The sheet was then washed and exposed to X-ray film . The clones identified by this
procedure could then be isolated from the replica p late. Note that this ‘sandwich’
technique is applicable only where two antibodies recognizing different determinants of
the same protein are available. However, if the protein is expressed as a fusion,
antibodies that bind to each component of the fusion can be used, efficiently selecting
for recombinant molecules. While plasmid libraries have been useful for expression
screening (Helfman et al. 1983, Helfman & Hughes 1987), it is much more convenient to
use bacteriophage-λ insertion vectors, because these have a higher capacity and the
efficiency of in vitro packaging allows large numbers of recombinants to be prepared and
screened. Immunological screening with phage-λ cDNA libraries was introduced by
Young and Davies (1983) using the expression vector λgt11, which generates fusion
proteins with β-galactosidase under the control of the lac promoter
In the original technique, screening was carried out using colonies of induced lysogenic
bacteria, which required the production of replica plates, as above. A simplification of
the method is possible by directly screening plaques of recombinant phage. In this
procedure (Fig.), the library is plated out at moderately high density (up to 5 × 10
4

plaques/9 cm
2
plate), with E. coli strain Y1090 as the host. This E. coli strain
overproduces the lac repressor and ensures that no expression of cloned sequences
(which may be deleterious to the host) takes place until the inducer isopropyl-β- D-
thiogalactoside (IPTG) is presented to the infected cells. Y1090 is also deficient in the
lon protease, hence increasing the stability of recombinant fusion proteins.
Fusion proteins expressed in plaques are absorbed on to a nitrocellulose membrane
overlay and this membrane is processed for antibody screening.
When a positive signal is identified on the membrane, the positive plaque can be picked
from the original agar plate (a replica is not necessary) and the recombinant phage can
be isolated. The original detection method using io dinated antibodies has been
superseded by more convenient methods using non-isotopic labels, which are also more
sensitive and have a lower background of nonspecific signal.

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Generally, these involve the use of unlabelled primary antibodies directed against the
polypeptide of interest, which are in turn recognized by secondary antibodies carrying
an enzymatic label. As well as eliminating the need for isotopes, such methods also
incorporate an amplification step, since two or more secondary antibodies bind to the
primary antibody. Typically, the secondary antibody recognizes the species-specific
constant region of the primary antibody and is conj ugated to either horseradish
peroxidase (De Wet et al. 1984) or alkaline phosphatase (Mierendorf et al. 1987), each of
which can in turn be detected using a simple colorimetric assay carried out directly on
the nitrocellulose filter. Polyclonal antibodies, which recognize many different epitopes,
provide a very sensitive probe for immunological screening, although they may also
cross-react to proteins in the expression host. Mon oclonal antibodies and cloned
antibody fragments can also be used, although the s ensitivity of such reagents is
reduced because only a single epitope is recognized.

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RESTRICTION MAPPING OF CLONED GENES

Representational difference analysis is a PCR subtraction technique, i.e. common
sequences between two sources are eliminated prior to amplification. The method was
developed for the comparative analysis of genomes (Lisitsyn et al. 1993) but has been
modified for cloning differentially expressed genes (Hubank & Schatz 1994). Essentially,
the technique involves the same principle as subtraction hybridization, in that a large
excess of a DNA from one source, the driver, is used to make common sequences in the
other source, the tester, unclonable (in this case unamplifiable). The general scheme is
shown in Fig.
cDNA is prepared from two sources, digested with restriction enzymes and amplified.
The amplified products from one source are then ann ealed to specific linkers that
provide annealing sites for a unique pair of PCR primers. These linkers are not added to
the driver cDNA. A large excess of driver cDNA is then added to the tester cDNA and the
populations are mixed. Driver/ driver fragments possess no linkers and cannot be
amplified, while driver/tester fragments possess only one primer annealing site and will
only be amplified in a linear fashion. However, cDNAs that are present only in the tester

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will possess linkers on both strands and will be am plified exponentially and can
therefore be isolated and cloned.
BLOTTING TECHNIQUES
We have seen how fusion proteins expressed in plaques produced by recombinant λgt11
or λZAP vectors may be detected by immunochemical s creening. A closely related
approach has been used for the screening and isolat ion of clones expressing
sequencespecific DNA-binding proteins. As above, a plaque lift is carried out to transfer
a print of the library on to nitrocellulose membranes. However, the screening is carried
out, without using an antibody, by incubating the m embranes with a radiolabelled
doublestranded DNA oligonucleotide probe, containing the recognition sequence for the
target DNA-binding protein. This technique is called south-western screening, because it
combines the principles of Southern and western blots.
It has been particularly successful in the isolation of clones expressing cDNA sequences
corresponding to certain mammalian transcription factors (Singh et al. 1988, Staudt et
al. 1988, Vinson et al. 1988, Katagiri et al. 1989, Williams et al. 1991, Xiao et al. 1991).
A limitation of this technique is that, since individual plaques contain only single cDNA
clones, transcription factors that function only in the form of heterodimers or as part of
a multimeric complex does not recognize the DNA probe and the corresponding cDNAs
cannot be isolated. Clearly the procedure can also be successful only in cases where the
transcription factor remains functional when expressed as a fusion polypeptide. It is
also clear that the affinity of the polypeptide for the specific DNA sequence must be
high, and this has led to the preferential isolation of certain types of transcription
factor. More recently, a similar technique has been used to isolate sequencespecific
RNA-binding proteins, in this case using a single-stranded RNA probe. By analogy to
the above, this is termed north-western screening and has been successful in a number
of cases (e.g. see Qian & Wilusz 1993; reviewed by Bagga & Wilusz 1999).
Both south-western and north-western screenings are most efficient when the
oligonucleotide contains the binding sequence in multimeric form. This may mean that
several fusion polypeptides on the filter bind to each probe, hence greatly increasing the
average dissociation time. To minimize non-specific binding, a large excess of unlabelled
double-stranded DNA (or single-stranded RNA) is mix ed with the specific probe.
However, it is usually necessary to confirm the specificity of binding in a second round

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of screening, using the specific oligonucleotide probe and one or more alternative probes
containing similar sequences that are not expected to be recognized.
SEQUENCING METHODS
The first significant DNA sequence to be obtained was that of the cohesive ends of
phage-λ DNA (Wu & Taylor 1971), which are only 12 bases long. The methodology used
was derived from RNA sequencing and was not applica ble to large-scale DNA
sequencing. An improved method, plus and minus sequencing, was developed and used
to sequence the 5386 bp phage ΦX 174 genome (Sanger et al. 1977a). This method was
superseded in 1977 by two different methods, that of Maxam and Gilbert
(1977) and the chain-termination or dideoxy method (Sanger et al. 1977b). For a while
the Maxam and Gilbert (1977) method, which makes us e of chemical reagents to bring
about base-specific cleavage of DNA, was the favoured procedure. However, refinements
to the chain-termination method meant that by the early 1980s it became the preferred
procedure.
To date, most large sequences have been determined using this technology, with the
notable exception of bacteriophage T7 (Dunn & Studier 1983). For this reason, only the
chain-termination method will be described here.
The chain-terminator or dideoxy procedure for DNA s equencing capitalizes on two
properties of DNA polymerases: (i) their ability to synthesize faithfully a complementary
copy of a single-stranded DNA template; and (ii) th eir ability to use 2′,3′-
dideoxynucleotides as substrates.

Once the analogue is incorporated at the growing point of the DNA chain, the 3′ end
lacks a hydroxyl group and is no longer a substrate for chain elongation. Thus, the
growing DNA chain is terminated, i.e. dideoxynucleotides act as chain terminators. In
practice, the Klenow fragment of DNA polymerase is used because this lacks the 5′ → 3′

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exonuclease activity associated with the intact enzyme. Initiation of DNA synthesis
requires a primer and this is usually a chemically synthesized oligonucleotide which is
annealed close to the sequence being analysed.
DNA synthesis is carried out in the presence of the four deoxynucleoside triphosphates,
one or more of which is labelled with 32P, and in f our separate incubation mixes
containing a low concentration of one each of the four dideoxynucleoside triphosphate
analogues. Therefore, in each reaction there is a population of partially synthesized
radioactive DNA molecules, each having a common 5′ end, but each varying in length to
a base-specific 3′ end.

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After a suitable incubation period, the DNA in each mixture is denatured and
electrophoresed in a sequencing gel. A sequencing gel is a high-resolution gel designed
to fractionate single-stranded (denatured) DNA fragments on the basis of their size and
which is capable of resolving fragments differing in length by a single base pair. They
routinely contain 6–20% polyacrylamide and 7 mol/l urea. The function of the urea is to
minimize DNA secondary structure, which affects electrophoretic mobility. The gel is
run at sufficient power to heat up to about 70°C. This also minimizes DNA secondary
structure. The labelled DNA bands obtained after such electrophoresis are revealed by
autoradiography on large sheets of X-ray film and from these the sequence can be read.


To facilitate the isolation of single strands, the DNA to be sequenced may be cloned into
one of the clustered cloning sites in the lac region of the M13 mp series of vectors.

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A feature of these vectors is that cloning into the same region can be mediated by any
one of a large selection of restriction enzymes but still permits the use of a single
sequencing primer.
Modifications of chain-terminator sequencing
The sharpness of the autoradiographic images can be improved by replacing the
32
P-
radiolabel with the much lower-energy
33
P or
35
S. In the case of
35
S, this is achieved by
including a α-
35
S-deoxynucleoside triphosphate in the sequencing reaction.
This modified nucleotide is accepted by DNA polymerase and incorporated into the
growing DNA chain. Non-isotopic detection methods have also been developed with
chemiluminescent, chromogenic or fluorogenic reporter systems. Although the
sensitivity of these methods is not as great as with radiolabels, it is adequate for many
purposes.


Other technical improvements to Sanger’s original method have been made by replacing
the Klenow fragment of Escherichia coli DNA polymerase I. Natural or modified forms of
the phage T7 DNA polymerase (‘Sequenase’) have foun d favour, as has the DNA
polymerase of the thermophilic bacterium Thermus aquaticus (Taq DNA polymerase).
The T7 DNA polymerase is more processive than Klenow polymerase, i.e. it is capable of
polymerizing a longer run of nucleotides before releasing them from the template. Also,
its incorporation of dideoxynucleotides is less affected by local nucleotide sequences
and so the sequencing ladders comprise a series of bands with more even intensities.
The Taq DNA polymerase can be used in a chain-termi nation reaction carried out at
high temperatures (65–70°C) and this minimizes chain-termination artefacts caused by
secondary structure in the DNA. Tabor and Richardso n (1995) have shown that
replacing a single phenylalanine residue of Taq DNA polymerase with a tyrosine residue

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results in a thermostable sequencing enzyme that no longer discriminates between
dideoxy- and deoxynucleotides.
The combination of chain-terminator sequencing and M13 vectors to produce single-
stranded DNA is very powerful. Very good-quality sequencing is obtainable with this
technique, especially when the improvements given by 35S-labelled precursors and T7
DNA polymerase are exploited.
Further modifications allow sequencing of ‘double-stranded’ DNA, i.e. double-stranded
input DNA is denatured by alkali and neutralized and then one strand is annealed with
a specific primer for the actual chainterminator sequencing reactions. This approach
has gained in popularity as the convenience of having a universal primer has grown less
important with the widespread availability of oligonucleotide synthesizers.
With this development, Sanger sequencing has been liberated from its attachment to
the M13 cloning system; for example, polymerase chain reaction (PCR)-amplified DNA
segments can be sequenced directly. One variant of the doublestranded approach, often
employed in automated sequencing, is ‘cycle sequenc ing’. This involves a linear
amplification of the sequencing reaction, using 25 cycles of denaturation, annealing of a
specific primer to one strand only and extension in the presence of Taq DNA polymerase
plus labeled dideoxynucleotides. Alternatively, labelled primers can be used with
unlabelled dideoxynucleotides.
Automated DNA sequencing
In manual sequencing, the DNA fragments are radiolabelled in four chain-termination
reactions, separated on the sequencing gel in four lanes and detected by
autoradiography. This approach is not well suited to automation. To automate the
process, it is desirable to acquire sequence data in real time by detecting the DNA
bands within the gel during the electrophoretic separation. However, this is not trivial,
as there are only about 10
−15
−10
−16
moles of DNA per band. The solution to the
detection problem is to use fluorescence methods. In practice, the fluorescent tags are
attached to the chain-terminating nucleotides. Each of the four dideoxynucleotides
carries a spectrally different fluorophore. The tag is incorporated into the DNA molecule
by the DNA polymerase and accomplishes two operatio ns in one step: it terminates
synthesis and it attaches the fluorophore to the end of the molecule. Alternatively,
fluorescent primers can be used with nonlabelled dideoxynucleotides. By using four

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different fluorescent dyes, it is possible to electrophorese all four chain-terminating
reactions together in one lane of a sequencing gel. The DNA bands are detected by their
fluorescence as they electrophorese past a detector (Fig. 7.6).

If the detector is made to scan horizontally across the base of a slab gel, many separate
sequences can be scanned, one sequence per lane. Because the different fluorophores
affect the mobility of fragments to different exten ts, sophisticated software is
incorporated into the scanning step to ensure that bands are read in the correct order.
A simpler method is to use only one fluorophore and to run the different chain-
terminating reactions in different lanes. For high-sensitivity DNA detection in four-
colour sequencing and high-accuracy base calling, one would ideally like the following
criteria to be met: each of the four dyes to exhibit strong absorption at a common laser
wavelength; to have an emission maximum at a distinctly different wavelength; and to
introduce the same relative mobility shift of the DNA sequencing fragments. Recently,
dyes with these properties have been identified and successfully applied to automated
sequencing (Glazer & Mathies 1997).
Automated DNA sequencers offer a number of advantag es that are not particularly
obvious. First, manual sequencing can generate excellent data, but even in the best

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sequencing laboratories poor autoradiographs are fr equently produced that make
sequence reading difficult or impossible. Usually the problem is related to the need to
run different termination reactions in different tracks of the gel. Skilled DNA sequencers
ignore bad sequencing tracks, but many laboratories do not. This leads to poor-quality
sequence data. The use of a single-gel track for all four dideoxy reactions means that
this problem is less acute in automated sequencing. Nevertheless, it is desirable to
sequence a piece of DNA several times and on both strands, to eliminate errors caused
by technical problems. It should be noted that long runs of the same nucleotide or a
high G+C content can cause compression of the bands on a gel, necessitating manual
reading of the data, even with an automated system. Note also that multiple, tandem
short repeats, which are common in the DNA of highe r eukaryotes, can reduce the
fidelity of DNA copying, particularly with Taq DNA polymerase.
The second advantage of automated DNA sequencers is that the output from them is in
machine-readable form. This eliminates the errors that arise when DNA sequences are
read and transcribed manually.
A third advantage derives from the new generation o f sequencers that have been
introduced recently. In these sequencers, the slab gel is replaced with 48 or 96
capillaries filled with the gel matrix. The key feature of this system is that the
equipment has been designed for use with robotics, thereby minimizing hands-on time
and increasing throughput. With a 96-capillary sequencer, it is possible to sequence up
to 750 000 nucleotides per day.
PURIFICATION STRATEGIES OF EXPRESSED His-TAGGED PRO TEINS.
Previous studies of proteins generally examined a single protein at a time. With the
recent sequencing of whole genomes, proteome analys is has turned to methods that
allow simultaneous monitoring of multiple proteins. Microarrays have been used for
DNA for some time, but the variable structures and properties of proteins made such an
array approach more difficult. Nonetheless, new technologies have been developed that
allow high-throughput analysis of proteins. As a result, protein microarrays have
recently become available for proteome analysis.
Protein microarrays have been used for the biochemi cal and enzymatic analysis of
proteins as well as to survey protein–protein interactions. So far most proteome arrays
have used the yeast, Saccharomyces cerevisiae, as model organism. A complete

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proteome analysis needs an array of approximately 6,000 proteins in this case. Such
arrays are assembled using proteins that have been tagged with groups allowing
binding to solid supports such as 96-well microtiter dishes or glass microscope slides.
Libraries that include nearly 90% of the yeast prot eins have been fused to the
glutathione-S-transferase (GST) tag, which allows binding to a solid support via
glutathione or to the His tag, which allows binding via nickel. These constructs have
been expressed under control of the GAL1 (galactose inducible) or CUP1 (copper
inducible) promoters. Such protein libraries may be pooled or distributed individually
into the wells of microtiter dishes. Simpler and less expensive screening is usually done
individually, whereas complex or expensive assays are more often run first on pooled
protein samples that are subdivided for further analysis if positive results are found.

To assemble a protein microarray, a library of His- tagged proteins is incubated
with a nickel coated glass slide. The proteins adhere to the slide wherever a nickel
ion is attached.

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The functional assays must be designed so that the arrays can be screened
conveniently, usually for fluorescence, less often for radioactivity. For example, the
yeast proteome has been screened for those proteins that bind calmodulin (a small
calcium binding protein) or phospholipids, in the laboratory of Michael Snyder at Yale
University.
The His-tagged proteins were attached to nickel-coated glass slides. Both calmodulin
and phospholipid were tagged with biotin. After binding of calmodulin or phospholipids
to the proteome array, the biotin was detected by s treptavidin carrying a Cy3
fluorescent label. This revealed 39 calmodulin-binding proteins of which six were
previously known. Some 150 phospholipid-binding proteins were also found.

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UNIT: 4 TRANSFORMATION TECHNIQUES
Purification of vectors DNA, restriction digestion, end modification, cloning of
foreign genes (from mRNA, genomic DNA) transformati on screening, selection,
expression and preservation. Transformation and tra nsfection techniques,
preparation of competent cells of bacteria, chemica l methods-calcium phosphate
precipitation methods, liposome mediated method, ph ysical methods-
Electroporation, gene gun method. Method of DNA tra nsfer to yeast, mammalian
and plant cells, transformation and transfection efficiency.
PURIFICATION OF VECTORS DNA
Introduction
In DNA cloning, a DNA fragment that contains a gene of interest is inserted into the
purified DNA genome of a self-replicating genetic element - generally a virus or a
plasmid. A DNA fragment containing a human gene, for example, can be joined in a test
tube to the chromosome of a bacterial virus, and the new recombinant DNA molecule can
then be introduced into a bacterial cell. Starting with only one such recombinant DNA
molecule that infects a single cell, the normal replication mechanisms of the virus can
produce more than 10
12
identical virus DNA molecules in less than a day, thereby
amplifying the amount of the inserted human DNA fragment by the same factor. A virus
or plasmid used in this way is known as a cloning vector, and the DNA propagated by
insertion into it is said to have been cloned.
A DNA Library Can Be Made Using Either Viral or Plasmid Vectors
In order to clone a specific gene, one begins by co nstructing a DNA librarya
comprehensive collection of cloned DNA fragments, including (one hopes) at least one
fragment that contains the gene of interest. The library can be constructed using either
a virus or a plasmid vector and is generally housed in a population of bacterial cells.
The principles underlying the methods used for cloning genes are the same for either
type of cloning vector, although the details may be different. For simplicity, in this
chapter we ignore these differences and illustrate the methods with reference to plasmid
vectors. The plasmid vectors used for gene cloning are small circular molecules of
double-stranded DNA derived from larger plasmids th at occur naturally in bacterial
cells. They generally account for only a minor fraction of the total host bacterial cell
DNA, but they can easily be separated on the basis of their small size from

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chromosomal DNA molecules, which are large and prec ipitate as a pellet upon
centrifugation. For use as cloning vectors, the purified plasmid DNA circles are first cut
with a restriction nuclease to create linear DNA molecules. The cellular DNA to be used
in constructing the library is cut with the same restriction nuclease, and the resulting
restriction fragments (including those containing the gene to be cloned) are then added
to the cut plasmids and annealed via their cohesive ends to form recombinant DNA
circles. These recombinant molecules containing for eign DNA inserts are then
covalently sealed with the enzyme DNA ligase.
The formation of a recombinant DNA molecule.

The cohesive ends produced by many kinds of restric tion nucleases allow two DNA
fragments to join by complementary base-pairing. DN A fragments joined in this
way can be covalently linked in a highly efficient reaction catalyzed by the
enzyme DNA ligase. In this example a recombinant pl asmid DNA molecule
containing a chromosomal DNA insert is formed.
In the next step in preparing the library, the recombinant DNA circles are introduced
into bacterial cells that have been made transiently permeable to DNA; such cells are
said to be transfected with the plasmids. As these cells grow and divide, doubling in
number every 30 minutes, the recombinant plasmids a lso replicate to produce an
enormous number of copies of DNA circles containing the foreign DNA.
Purification and amplification of a specific DNA se quence by DNA cloning in a
bacterium.

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Each bacterial cell carrying a recombinant plasmid develops into a colony of
identical cells, visible as a spot on the nutrient agar.
By inoculating a single colony of interest into a liquid culture, one can obtain a
large number of identical plasmid DNA molecules, ea ch containing the same DNA
insert.
Many bacterial plasmids carry genes for antibiotic resistance, a property that can be
exploited to select those cells that have been successfully transfected; if the bacteria are
grown in the presence of the antibiotic, only cells containing plasmids will survive. Each
original bacterial cell that was initially transfected will, in general, contain a different
foreign DNA insert; this insert will be inherited by all of the progeny cells of that
bacterium, which together form a small colony in a culture dish.
The mixture of many different surviving bacteria contains the DNA library, composed of
a large number of different DNA inserts. The problem is that only a few of the bacteria
will harbor the particular recombinant plasmids that contain the desired gene. One
needs to be able to identify these rare cells in order to recover the DNA of interest in
pure form and in useful quantities. Before discussing how this is achieved, we need to

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describe a second strategy for generating a DNA library that is commonly used in gene
cloning.
RESTRICTION DIGESTION
Cleaving the entire genome of a cell with a specific restriction nuclease as just described
is sometimes called the "shotgun" approach to gene cloning. It produces a very large
number of DNA fragments - on the order of a million for a mammalian genome - which
will generate millions of different colonies of transfected bacterial cells. Each of these
colonies will be composed of a clone derived from a single ancestor cell and therefore
will harbor a recombinant plasmid with the same ins erted genomic DNA sequence.
Such a plasmid is said to contain a genomic DNA clo ne, and the entire collection of
plasmids is said to constitute a genomic DNA library. But because the genomic DNA is
cut into fragments at random, only some fragments w ill contain genes; many will
contain only a portion of a gene, while most of the genomic DNA clones obtained from
the DNA of a higher eucaryotic cell will contain only noncoding DNA.
An alternative strategy is to begin the cloning process by selecting only those DNA
sequences that are transcribed into RNA and thus ar e presumed to correspond to
genes. This is done by extracting the mRNA (or a purified subfraction of the mRNA) from
cells and then making a complementary DNA (cDNA) co py of each mRNA molecule
present; this reaction is catalyzed by the reverse transcriptase enzyme of retroviruses,
which synthesizes a DNA chain on an RNA template. T he single- stranded DNA
molecules synthesized by the reverse transcriptase are converted into double-stranded
DNA molecules by DNA polymerase, and these molecule s are inserted into a plasmid or
virus vector and cloned.
The synthesis of cDNA. A DNA copy (cDNA) of an mRNA molecule is produced by
the enzyme reverse transcriptase, thereby forming a DNA/RNA hybrid helix.
Treating the DNA/RNA hybrid with alkali selectively degrades the RNA strand into
nucleotides. The remaining single-stranded cDNA is then copied into double-
stranded cDNA by the enzyme DNA polymerase. As indi cated, both reverse
transcriptase and DNA polymerase require a primer t o begin their synthesis.

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For reverse transcriptase a small oligonucleotide is used; in this example oligo(dT)
has been annealed with the long poly-A tract at the 3' end of most mRNAs. Note
that the double-stranded cDNA molecule produced her e lacks cohesive ends; such
blunt-ended DNA molecules
Each clone obtained in this way is called a cDNA clone, and the entire collection of
clones derived from one mRNA preparation constitute s a cDNA library. There are
important differences between genomic DNA clones and cDNA clones
Genomic clones represent a random sample of all of the DNA sequences in an organism
and, with very rare exceptions, will be the same regardless of the cell type used to
prepare them. By contrast, cDNA clones contain only those regions of the genome that
have been transcribed into mRNA; as the cells of different tissues produce distinct sets
of mRNA molecules, a different cDNA library will be obtained for each type of cell used
to prepare the library.
CLONING OF FOREIGN GENES (FROM mRNA, GENOMIC DNA )
The use of a cDNA library for gene cloning has several advantages. First, some proteins
are produced in very large quantities by specialized cells. In this case, the mRNA

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encoding the protein is likely to be produced in such large quantities that a cDNA
library prepared from the cells will be highly enriched for the cDNA molecules encoding
the protein, greatly reducing the problem of identifying the desired clone in the library.
The differences between cDNA clones and genomic DNA clones.

In this example gene A is infrequently transcribed while gene B is frequently
transcribed, and both genes contain introns (green). In the genomic DNA clones
both the introns and the nontranscribed DNA are inc luded, and most clones will
contain only part of the coding sequence of a gene. In the cDNA clones the intron
sequences have been removed by RNA splicing during the formation of the mRNA,
and a continuous coding sequence is therefore present.
Hemoglobin, for example, is made in large amounts b y developing erythrocytes (red
blood cells); for this reason the globin genes were among the first to be cloned. By far
the most important advantage of cDNA clones is that they contain the uninterrupted
coding sequence of a gene. Eucaryotic genes usually consist of short coding sequences
of DNA (exons) separated by longer noncoding sequen ces (introns); the production of
mRNA entails the removal of the noncoding sequences from the initial RNA transcript
and the splicing together of the coding sequences. Neither bacterial nor yeast cells will
make these modifications to the RNA produced from a gene of a higher eucaryotic cell.

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Thus, if the aim of the cloning is either to deduce the amino acid sequence of the
protein from the DNA or to produce the protein in bulk by expressing the cloned gene in
a bacterial or yeast cell, it is much preferable to start with cDNA.
Genomic and cDNA libraries are inexhaustible resources that are widely shared among
investigators. Today, many such libraries are also available from commercial sources.
cDNA Libraries Can Be Prepared from Selected Popula tions of mRNA Molecules
When cDNAs are prepared from cells that express the gene of interest at extremely high
levels, the majority of cDNA clones may contain the gene sequence, which can therefore
be selected with minimal effort. For less abundantly transcribed genes, various methods
can be used to enrich for particular mRNAs before m aking the cDNA library. If an
antibody against the protein is available, for example, it can be used to precipitate
selectively those polyribosomes that have the appropriate growing polypeptide chains
attached to them. Since these polyribosomes will also have attached to them the mRNA
coding for the protein, the precipitate may be enriched in the desired mRNA by as much
as 1000-fold.
Subtractive hybridization
Subtractive hybridization provides a powerful alternative way of enriching for particular
nucleotide sequences prior to cDNA cloning. This selection procedure can be used, for
example, if two closely related cell types are available from the same organism, only one
of which produces the protein or proteins of interest. It was first used to identify cell-
surface receptor proteins present on T lymphocytes but not on B lymphocytes. It can
also be used wherever a cell that expresses the protein has a mutant counterpart that
does not. The first step is to synthesize cDNA molecules using the mRNA from the cell
type that makes the protein of interest. These cDNAs are then hybridized with a large
excess of mRNA molecules from the second cell type.
Those rare cDNA sequences that fail to find a complementary mRNA partner are likely
to represent mRNA sequences present only in the first cell type. Because these cDNAs
remain unpaired after the hybridization, they can be purified by a simple biochemical
procedure (a hydroxyapatite column) that separates single-stranded from double-
stranded nucleic acids.
Besides providing a powerful way to clone genes who se products are known to be
restricted to a specific differentiated cell type, cDNA libraries prepared after subtractive

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hybridization are useful for defining the differences in gene expression between any two
related types of cells.

In this example the technique is used to purify rare cDNA clones corresponding to
mRNA molecules present in T lymphocytes but not in B lymphocytes. Because the
two cell types are very closely related, most of the mRNAs will be common to both
cell types; subtractive hybridization is thus a powerful way to enrich for those
specialized molecules that distinguish the two cells.

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TRANSFORMATION SCREENING
The blue-white screen is a screening technique that allows for the detec tion of
successful ligations in vector-based gene cloning. DNA of interest is ligated into
a vector. The vector is then transformed into competent cell (bacteria). The competent
cells are grown in the presence of X-gal. If the ligation was successful, the bacterial
colony will be white; if not, the colony will be blue. This technique allows for the quick
and easy detection of successful ligation.
Molecular cloning is one of the most commonly used procedures in molecular biology. A
gene of interest may be inserted into a plasmid vector via ligation, and the plasmid is
then transformed into Escherichia coli cells. However, not all the plasmids transformed
into cells may contain the desired gene insert and checking each individual colony for
the presence of the insert is time-consuming, so a method for the detection of the insert
is therefore useful for making this procedure less time and labor intensive. One of the
early methods developed for the detection of insert is blue-white screening which allows
for identification of successful products of cloning reactions through the colour of
thebacterial colony.
The method is based on the principle of α-complementation of the β-galactosidase gene.
This phenomenon of α-complementation was first demo nstrated in work done by Agnes
Ullmann in the laboratory of François Jacob and Jacques Monod.
Molecular mechanism:
β-galactosidase is a protein encoded by the lacZ gene of the lac operon, and it exists as
a homotetramer in its active state. However, a muta nt β-galactosidase strain of E.
coli has its N-terminal residues 11—41 deleted and this mutant, the ω-peptide, is
unable to form a tetramer and is inactive. This mutant form of protein however may
return fully to its active tetrameric state in the presence of an N-terminal fragment of
the protein, the α-peptide. The rescue of function of the mutant β-galactosidase by the
α-peptide is called α-complementation.
In this method of screening, the host E. coli strain carries the lacZ deletion mutant
(lacZ∆M15} which contains the ω-peptide, while the plasmids used carry
the lacZα sequence which encodes the first 59 residues of β- galactosidase, the α-

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peptide. Neither are functional by themselves. However, when the two peptides are
expressed together, as when a plasmid containing the lacZα sequence is transformed
into a lacZ∆M15 cells, they form a functional β-galactosidase enzyme.
The blue/white screening method works by disrupting this α-complementation process.
The plasmid carries within the lacZα sequence an internal multiple cloning site (MCS).
This MCS within the lacZα sequence can be cut by restriction enzymes so that the
foreign DNA may be inserted within the lacZα gene, thereby disrupting the gene and
thus production of α-peptide. Consequently, in cells containing the plasmid with an
insert, no functional β-galactosidase may be formed.

The presence of an active β-galactosidase can be detected by
X-gal, a colourless analog of
lactose that may be cleaved by β-galactosidase to form 5-bromo-4-chloro-indoxyl, which
then spontaneously dimerizes and oxidizes to form a bright blue insoluble pigment 5,5'-
dibromo-4,4'-dichloro-indigo. This results in a characteristic blue colour in cells
containing a functional β-galactosidase. Blue colonies therefore show that they may
contain a vector with an uninterrupted
lacZα ( therefore no insert), while white colonies,
where X-gal is not hydrolyzed, indicate the presenc e of an insert in lacZα which
disrupts the formation of an active β-galactosidase.
SELECTION, EXPRESSION AND PRESERVATION
A number of methods have been devised for screening of desired transformants. These
include the following:

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1. Direct selection for the desired gene
2. Identification of the clone from a gene library
Direct selection for the desired gene
In this method, the cloning experiment is designed in such a way that only the desired
recombinant clones are obtained. Selection here occurs at the plating out stage. An
example of direct selection is cloning of genes that specify antibiotic resistance such as
kanamycin, tetracycline or ampicillin resistance. For example, let us consider selection
of recombinants that contain gene for kanamycin res istance. The transformants are
plated on agar medium containing kanamycin. Only th e cells that contain the cloned
kanamycin resistance gene are able to survive and form colonies. Hence the technique
is called direct selection.
Another situation where direct selection of recombinant clones is done is the marker
rescue technique.

It makes use of auxotrophic mutant strains that have nutritional defects as the hosts
for transformation. For example, suppose an E.coli strain is available which has a
mutation in a gene encoding an enzyme involved in the biosynthesis of amino acid

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leucine. Such a mutant strain will only grow in a medium supplemented with leucine.
By cloning DNA from a normal strain i.e. one that can synthesize its own leucine, in the
mutant strain and selecting those transformants that can grow in absence of leucine it
is possible to isolate the gene of interest.
Marker rescue is applicable for most genes that encode for biosynthetic enzymes as
clones of these genes can be studied on minimal medium as described for leucine.
Auxotrophic strains of yeast and filamentous fungi are also available and marker rescue
has been used to select genes cloned in these organisms. However, the technique has
two limitations: i) a mutant strain must be available for the gene in question ii) a
medium on which only the wild type can survive is needed.
Identification of the clones from a gene library
A library has to be screened in order to find a clone. The identification of a specific clone
from a DNA library can be carried out by exploiting either the sequence of the clone or
the structure/ function of its expressed product. The strategy for screening depends
upon information about the gene of interest, the availability of probe and the cloning
method used.
One of the key elements required to identify a gene during screening is a probe. A probe
is a piece of DNA or RNA that contains a portion of the sequence complementary to the
desired gene for which we are searching. It is used to detect specific nucleic acid
sequences by hybridization (based on complementarit y). The probe can be labeled
radioactively (with P
32
) or non- radioactively (biotin, digoxigenin and fluorescent dyes
etc). Probes can be chemically synthesized based on the amino acid sequence of the
protein coded by the gene. Probes can be homologous or heterologous.
Homologous probe - a probe that is exactly complementary to the nucl eic acid
sequence for which we are searching; e. g. a human cDNA used for searching a human
genomic library.
Heterologous probe - a probe that is similar to, but not exactly complementary to the
nucleic acid sequence for which we are searching; e.g., a mouse cDNA probe used to
search a human genomic library.
There are several methods for screening DNA libraries. Some of the commonly used
methods are described below:

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1. Methods based on nucleic acid hybridization
2. Immunochemical methods
3. Screening DNA libraries using PCR
TRANSFORMATION AND TRANSFECTION TECHNIQUES
PREPARATION OF COMPETENT CELLS OF BACTERIA
1.
Dilute an overnight culture of cells @ 1:100 in 50 mL of LB + chloramphenicol
2.
Incubate shaking at 37˚C until OD600 ≈ 0.5
3.
Spin out cells at 4˚C
4.
Resuspend pellet in 15 mL ice-cold 0.1M MgCl2
5.
Spin out cells at 4˚C
6.
Resuspend pellet in 15 mL ice-cold 0.1M CaCl2
7.
Incubate on ice for 45 min
8.
Spin out cells at 4˚C
9.
Resuspend pellet in 5 mL ice-cold 0.1M CaCl2, 30% glycerol
10.
Aliquot into eppi tubes and snap-freeze
11.
Store a –80˚C
CHEMICAL METHODS-CALCIUM PHOSPHATE PRECIPITATION ME THODS
The ability of mammalian cells to take up exogenously supplied DNA from their culture
medium was first reported by Szybalska and Szybalski (1962). They used total uncloned
genomic DNA to transfect human cells deficient for the enzyme hypoxanthineguanine
phosphoribosyltransferase (HPRT). Rare HPRT-positive cells, which had presumably
taken up fragments of DNA containing the functional gene, were identified by selection
on HAT medium.
At this time, the actual mechanism of DNA uptake was not understood. Much later, it
was appreciated that successful DNA transfer in such experiments was dependent on
the formation of a fine DNA/calcium phosphate coprecipitate, which first settles on to
the cells and is then internalized. The precipitate must be formed freshly at the time of
transfection. It is thought that small granules of calcium phosphate associated with
DNA are taken up by endocytosis and transported to the nucleus, where some DNA
escapes and can be expressed (Orrantia & Chang, 199 0). The technique became
generally accepted after its application, by Graham and Van der Erb (1973), to the

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analysis of the infectivity of adenoviral DNA. It is now established as a general method
for the introduction of DNA into a wide range of cell types in culture.
However, since the precipitate must coat the cells, this method is suitable only for cells
growing in monolayers, not those growing in suspension or as clumps. As originally
described, calcium phosphate transfection was limited by the variable and rather low
proportion of cells that took up DNA (1–2%). Only a small number of these would be
stably transformed.
Improvements to the method have increased the transfection frequency to 20% for some
cell lines (Chu & Sharp 1981). A variant of the technique, using a different buffer
system, allows the precipitate to form slowly over a number of hours, and this can
increase stable transformation efficiency by up to 100-fold when using high-quality
plasmid DNA (Chen & Okayama 1987, 1988).
Other chemical transfection methods
The calcium phosphate method is applicable to many cell types, but some cell lines are
adversely affected by the coprecipitate due to its toxicity and are hence difficult to
transfect. Alternative chemical transfection methods have been developed to address
this problem. One such method utilizes diethylaminoethyl dextran (DEAE-dextran), a
soluble polycationic carbohydrate that promotes interactions between DNA and the
endocytotic machinery of the cell. This technique was initially devised to introduce viral
DNA into cells (McCutchan & Pango 1968) but was lat er adapted as a method for
plasmid DNA transfer (Milman & Herzberg 1981). The efficiency of the original
procedure was improved by Lopata et al. (1984) and Sussman and Milman (1984) by
adding after-treatments, such as osmotic shock or exposure to chloroquine, the latter
having been shown to inhibit the acidification of endosomal vesicles (Luthmann &
Magnusson 1983). Although efficient for the transient transfection of many cell types,
DEAE-dextran cannot be used to generate stably tran sformed cell lines. Another
polycationic chemical, the detergent Polybrene, has been used for the transfection of
Chinese hamster ovary (CHO) cells, which are not am enable to calcium phosphate
transfection (Chaney et al. 1986).
LIPOSOME MEDIATED METHOD
An alternative to chemical transfection procedures is to package DNA inside a
phosopholipid vesicle, which interacts with the target cell membrane and facilitates

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DNA uptake. The first example of this approach was provided by Schaffner (1980), who
used bacterial protoplasts containing plasmids to transfer DNA into mammalian cells.
Briefly, bacterial cells were transformed with a suitable plasmid vector and then treated
with chloramphenicol to amplify the plasmid copy nu mber. Lysozyme was used to
remove the cell walls, and the resulting protoplasts were gently centrifuged on to a
monolayer of mammalian cells and induced to fuse wi th them, using polyethylene
glycol. A similar strategy was employed by Wiberg et al. (1987), who used the
haemoglobin-free ghosts of erythrocytes as delivery vehicles. The procedures are very
efficient in terms of the number of transformants obtained, but they are also labour-
intensive and so have not been widely adopted as a general transfection method.
However, an important advantage is that they are gentle, allowing the transfer of large
DNA fragments without shearing. Yeast cells with the cell wall removed (sphaeroplasts)
have therefore been used to introduce yeast artificial chromosome (YAC) DNA into
mouse ES cells by this method, for the production o f YAC transgenic mice. More
widespread use has been made of artificial phosphol ipid vesicles, which are called
liposomes (Schaefer-Ridder et al. 1982). Initial liposome-based procedures were
hampered by the difficulty encountered in encapsula ting the DNA, and provided
transfection efficiency no better than the calcium phosphate method. However, a
breakthrough came with the discovery that cationic/ neutral lipid mixtures can
spontaneously form stable complexes with DNA that interact productively with the cell
membrane, resulting in DNA uptake by endocytosis (F elgner et al. 1987, 1994). This
low-toxicity transfection method, commonly known as lipofection, is one of the simplest
to perform and is applicable to many cell types that are difficult to transfect by other
means, including cells growing in suspension (e.g. Ruysscharet et al. 1994).
This technique is suitable for transient and stable transformation, and is sufficiently
gentle to be used with YACs and other large DNA fragments. The efficiency is also much
higher than that of chemical transfection methods – up to 90% of cells in a culture dish
can be transfected. A large number of different lipid mixtures is available, varying in
efficiency for different cell lines. A unique benefit of liposome gene-delivery vehicles is
their ability to transform cells in live animals following injection into target tissues or
even the bloodstream.

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Transfection efficiency has been improved and targeting to specific cell types achieved
by combining liposomes with viral proteins that promote cell fusion, nuclear targeting
signals and various molecular conjugates that recognize specific cell surface molecules.
PHYSICAL METHODS - ELECTROPORATION
Electroporation involves the generation of transient, nanometre-sized pores in the cell
membrane, by exposing cells to a brief pulse of electricity. DNA enters the cell through
these pores and is transported to the nucleus. This technique was first applied to
animal cells by Wong and Neumann (1982), who succes sfully introduced plasmid DNA
into mouse fibroblasts. The electroporation technique has been adapted to many other
cell types (Potter et al. 1984). The most critical parameters are the intensity and
duration of the electric pulse, and these must be determined empirically for different
cell types. However, once optimal electroporation parameters have been established, the
method is simple to carry out and highly reproducible. The technique has high input
costs, because a specialized capacitor discharge ma chine is required that can
accurately control pulse length and amplitude (Pott er 1988). Additionally, larger
numbers of cells may be required than for other methods because, in many cases, the
most efficient electroporation occurs when there is up to 50% cell death.
In an alternative method, pores are created using a finely focused laser beam (Kurata et
al. 1986). Although very efficient (up to 0.5% stable transformation), this technique is
applicable to only small numbers of cells and has not gained widespread use.
GENE GUN METHOD
Direct transfer methods
A final group of methods considered in this section encompasses those in which the
DNA is transferred directly into the cell nucleus. One such procedure is microinjection,
a technique that is guaranteed to generate successful hits on target cells but that can
only be applied to a few cells in any one experiment. This technique has been applied to
cultured cells that are recalcitrant to other transfection methods (Capecchi 1980), but
its principal use is to introduce DNA and other molecules into large cells, such as
oocytes, eggs and the cells of early embryos.
Particle bombardment is another direct delivery method, initially developed for the
transformation of plants. This involves coating small metal particles with DNA and then

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accelerating them into target tissues using a powerful force, such as a blast of high-
pressure gas or an electric discharge through a water droplet.
In animals, this technique is most often used to transfect multiple cells in tissue slices
rather than cultured cells (e.g. Arnold et al. 1994, Lo et al. 1994). It has also been used
to transfer DNA into skin cells in vivo (Haynes et al. 1996).
METHOD OF DNA TRANSFER TO YEAST CELLS
Genetic transformation of Saccharomyces cerevisiae, was first reported by Oppenoorth
in 1960; however, other workers were unable to repeat his
results. Khan and Sen carried out an extensive study of DNA transformation of various
genetic markers with a number of different yeast species. Their procedure was to grow
yeast cultures in the presence of DNA extracted from various yeast strains and then to
screen for colonies with a transformed phenotype. They investigated the effects of DNA
concentration and the age of cells for best transformation.
Their studies furnished convincing evidence for the transformation of yeast but failed to
stimulate much interest in the phenomenon. In 1989, Fincham published a
comprehensive review of fungal transformation, whic h discussed transformation
technology and mechanisms of plasmid integration and expression to that date.
The spheroplast method
The removal of the yeast cell wall by enyzmatic treatment to yield protoplasts was first
reported by Eddy and Williamson in 1957, based on the observation by Giaja that the
“the gut juice of the snail Helix pomatia dissolves the cell wall of whole yeasts”.
Protoplasts, produced by the treatment of yeast cells with snail enzyme, regenerated
when embedded in medium containing 30% gelatin or 2 % agar. Hutchison and Hartwell
noted that treatment of yeast cells with Glusulase (commercial snail enzyme) did not
remove the cell wall completely and suggested that cells produced in this way be termed
“spheroplasts”. In 1974, Kao and Michayluk (68) showed that polyethylene glycol (PEG)
stimulated the fusion of plant protoplasts. Van Solingen and van der Plaat used PEG
plus CaCl
2 treatment to fuse yeast spheroplasts derived from strains of the same mating
type but with complementary nutritional requirements. This experimental procedure
resulted in the exchange of genetic information in yeast by a nonsexual process and
could be considered analogous to the soon-to-bediscovered process of transformation.

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In 1977, Ratzkin and Carbon reported the isolation of specific hybrid plasmids,
constructed from an E. coli plasmid and fragments of yeast DNA, which complemented
the hisB and leuB mutations in E. coli. The first successful protocol for the genetic
transformation of S. cerevisiae, developed by Hinnen et al. (58), had its origins in the
investigation of yeast spheroplasts and spheroplast fusion. They used the pYEleu10
plasmid containing the LEU2 gene to transform a yea st leu2 3-112 mutant to
prototrophy. They removed the yeast cell wall enzymatically and stabilized the resulting
spheroplasts with 1.0 M sorbitol before treating them with PEG and plasmid DNA. The
treated yeast cells were then suspended in regeneration agar and plated onto medium
to select for LEU+ cells. More than 100 putative transformants were then analyzed by
Southern blot analysis using the E. coli plasmid sequence as a probe. Three types of
transformants were identified.
In type I transformants, the plasmid DNA was integrated adjacent to the leu2 locus. In
type II transformants, the plasmid had integrated a t other locations in the yeast
genome. Type III transformants did not contain any bacterial DNA sequences. The
authors conclude that the type III transformants were most likely due to replacement of
the leu2 3-112 allele with the LEU2 sequence from t he transforming plasmid. The
transformation efficiency was in the range of 30–50 transformants/mg plasmid DNA.
Because this plasmid did not contain a yeast replicon, integration was required for the
transformants to be stable.
Five months later, Beggs reported the transformation of yeast with an autonomously
replicating plasmid. She constructed chimeric plasmids by inserting the endogenous
autonomously replicating yeast 2-mm circle into the bacterial plasmid pMB9. These
chimeric plasmids were then used to construct a yeast DNA library. Two plasmids from
this library, pJDB248 and pJBD219, proved to complement the leuB mutation in E.coli.
These plasmids were then used to transform a yeast strain containing the leu2-3
mutant allele utilizing essentially the same protocol as that reported by Hinnen et al.
The LEU+ transformants occurred at a frequency of 10
-5
to 10
-3
and an efficiency of 1 ´
104 transformants/mg plasmid DNA. Southern blot ana lysis was used to confirm the
presence of plasmid sequences, verifying that DNA uptake and genetic transformation
had occurred. Finally, the plasmids were recovered from the transformed yeast cells by

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transformation back into E. coli and were shown to have a structure similar to the
original plasmid construct.
Plasmid Vector Development
Three additional types of chimeric plasmid vectors were developed by Struhl et al. (i) YIp
(yeast integrating plasmids), which are unable to replicate and transform by integration
into the genome of the recipient strain; (ii) YEp (yeast episomal plasmids), which carry
the replication origin of the yeast 2-mm circle, an endogenous yeast plasmid, and can
replicate in the recipient cell; and (iii) YRp (yeast replicating plasmids), which can
replicate utilizing yeast autonomous replicating sequences (ARS). These authors showed
that integrating vectors transformed with low efficiencies, 1–10 transformants/ mg.
Plasmids that could replicate in the yeast cell tra nsformed with much higher
efficiencies. The YEp vectors transformed with an e fficiency of 0.5–2.0 X 10
4
transformants/mg input plasmid DNA, and the YRp7 pl asmid produced 0.5–2.0 X 10
3
transformants/mg input plasmid DNA. Struhl et al. demonstrated that plasmids that
require integration into the genome transform less efficiently than those yeast plasmid
vectors that can replicate autonomously in the yeast cell. Since then, two other yeast
plasmid vectors have been developed. Yeast centromere plasmids (YCp) that carry an
ARS and a yeast centromere are more stable than YRp plasmids but are present in only
one copy per cell. Yeast artificial chromosomes (YACs) are propagated as a circular
plasmid with a centromere and an ARS plus two selectable markers, two telomeres, and
a cloning site. The vector is linearized by the removal of a sequence between the
telomeres, and foreign DNA is inserted into the cloning site. The result is a linear
artificial chromosome, 100–1000 kb in length, that can be propagated through mitosis
and meiosis
METHOD OF DNA TRANSFER TO MAMMALIAN CELLS
Gene transfer technology provides the ability to genetically manipulate the cells of
higher animals. During the 1970s it became possible to introduce exogenous DNA
constructs into higher eukaryotic cells in vitro. Mammalian (germline) transgenesis was
first achieved in the early 1980s. The model used in this study was mice. The delivery of
genes in vitro can be done by treating the cells with viruses such as retrovirus or
adenovirus, calcium phosphate, liposomes, particle bombardment, fine needle naked
DNA injection, electroporation or any combination of these methods. These are the

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powerful tools for research and have possible applications in gene therapy. A number of
valuable techniques used to transfer genes in animals and plants cells and their scope
and contributions are explained below.
Electroporation
Electroporation uses electrical pulse to produce tr ansient pores in the plasma
membrane thereby allowing macromolecules into the cells. It is an efficient process to
transfer DNA into cells. Microscopic pores are induced in biological membrane by the
application of electric field. These pores are known as electropores which allow the
molecules, ions and water to pass from one side of the membrane to another. The pores
can be recovered only if a suitable electric pulse is applied. The electropores reseal
spontaneously and the cell can recover. The formation of electropores depends upon the
cells that are used and the amplitude and duration of the electric pulse that is applied
to them. Electric currents can lead to dramatic heating of the cells that can results in
cell death. Heating effects are minimized by using relatively high amplitude, a short
duration pulse or by using two very short duration pulses. Various applications of
electroporation are listed.
In terms of mammalian transgenesis, electroporation is an effective method of
introducing exogenous DNA into embryonic stem (ES) cells. This technique has
recently, been used to transfer genes into cultured mammalian embryos at defined
stages of development. It was reported that there is an increase from 12 to 19% of
transgenic bovine blastocysts when electroporation was included in an otherwise
passive sperm-DNA uptake protocol. Similar findings were reported, again with
transgenic bovine blastocysts. Fish species were al so reported to be genetically
manipulated in this way. Electroporation has been reported to enhance the level of gene
expression and significantly improve immune respons es elicited to DNA vaccines in
both large and small animals.
General applications of electroporation:
Introduction of exogeneous DNA into animal cell lines, plant protoplast, yeast protoplast
and bacterial protoplast.
Electroporation can be used to increase efficiency of transformation or transfection of
bacterial cells.

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Wheat, rice, maize, tobacco have been stably transformed with frequency upto 1% by
this method.
Genes encoding selectable marker may be used to int roduce genes using
electroporation.
To study the transient expression of molecular constructs.
Electroporation of early embryo may result in the production of transgenic animals.
Hepatocytes, epidermal cells, haematopoietic stem cells, fibroblast, mouse T and B
lymphocytes can be transformed by this technique.
Naked DNA may be used for gene therapy by applying electroporation device on animal
cells.
Advantages of electroporation
Method is fast.
Less costly.
Applied for a number of cell types.
Simultaneously a large number of cell can be treated.
High percentage of stable transformants can be produced
Microinjection
In microinjection DNA can be introduced into cells or protoplast with the help of very
fine needles or glass micropipettes having the diameter of 0.5 to 10 micro m. Some of
the DNA injected may be taken up by the nucleus. Co mputerized control of holding
pipette, needle, microscope stage and video technology has improved the efficiency of
this technique. The advantages, limitations and applications are listed.
Advantages of microinjection.
Frequency of stable integration of DNA is far better as compare to other methods.
Method is effective in transforming primary cells as well as cells in established cultures.
The DNA injected in this process is subjected to less extensive modifications.
Mere precise integration of recombinant gene in limited copy number can be obtained.
Limitations of microinjection.
Costly.
Skilled personal required.
More useful for animal cells.
Embryonic cells preferred for manipulation.

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Knowledge of mating timing, oocyte recovery is essential.
Method is useful for protoplasts and not for the walled cells.
Process causes random integration.
Rearrangement or deletion of host. DNA adjacent to site of integration are common.
Applications of microinjection.
Process is applicable for plant cell as well as animal cell but more common for animal
cells.
Technique is ideally useful for producing transgenic animal quickly.
Procedure is important for gene transfer to embryonic cells.
Applied to inject DNA into plant nuclei.
Method has been successfully used with cells and protoplast of tobacco, alfalfa etc.
Microinjection is potentially a useful method for simultaneous introduction of multiple
bioactive compounds such as antibodies, peptides, RNAs, plasmids, diffusion markers,
elicitors, Ca2+ as well as nucleus and artificial micro or
nanoparticles containing those chemicals into the same target single-cells.
Macroinjection
Macroinjection is the method tried for artificial DNA transfer to cereals plants that show
inability to regenerate and develop into whole plants from cultured cells. Needles used
for injecting DNA are with the diameter greater than cell diameter. DNA injected with
conventional syringe into region of plant which will develop into floral tillers. Around 0.3
ml of DNA solution is injected at a point above tiller node until several drops of solution
came out from top of young inflorescence.
Timing of injection is important and should be fourteen days before meiosis. This
method was found to be successful with rye plants. It is also being attempted for other
cereals plants. Advantages and limitation of macroinjections are listed
Advantages and limitations of macroinjection.
This technique does not require protoplast.
Instrument will be simple and cheap.
Methods may prove useful for gene transfer into cereals which do not regenerate from
cultured cell easily.
Technically simple.
Limitations

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Less specific.
Less efficient.
Frequency of transformation is very low.
Biolistics or microprojectiles for DNA transfer
Biolistics or particle bombardment is a physical me thod that uses accelerated
microprojectiles to deliver DNA or other molecules into intact tissues and cells. The
method was developed initially to transfer genes into plants by Sanford. Biolistics
transformation is relatively new and novel method amongst the physical methods for
artificial transfer of exogenous DNA. This method avoids the need of protoplast and is
better in efficiency. This technique can be used for any plant cells, root section,
embryos, seeds and pollen.
The gene gun is a device that literally fires DNA into target cells. The DNA to be
transformed into the cells is coated onto microscopic beads made of either gold or
tungsten. Beads are carefully coated with DNA. The coated beads are then attached to
the end of the plastic bullet and loaded into the firing chamber of the gene gun. An
explosive force fires the bullet down the barrel of the gun towards the target cells that
lie just beyond the end of the barrel. When the bullet reaches the end of the barrel it is
caught and stopped, but the DNA coated beads continue on toward the target cells.
Some of the beads pass through the cell wall into the cytoplasm of the target cells. Here
the bead and the DNA dissociate and the cells becom e transformed. Once inside the
target cells, the DNA is solubilised and may be expressed.
This approach, sometimes known as ‘biolistics’, was originally developed for plant
transgenesis but has been shown to be effective for transferring transgenes into
mammalian cells in vivo. The gene gun is particularly useful for transforming cells that
are difficult to transform by other methods, e.g. plant cells. It is also gaining in use as a
method for transferring DNA construct into whole animals. The general applications of
biolistics are listed
General applications of biolistics.
Biolistics technique has been used successfully to transform soyabean, cotton, spruce,
sugarcane, papaya, corn, sunflower, rice, maize, wheat, tobacco etc.
Genomes of subcellular organelles have been accessible to genetic manipulation by
biolistic method.

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Mitochondria of plants and chloroplast of chlamydomonas have been transformed.
Method can be applied to filamentous fungi and yeast (mitochondria).
The particle gun has also been used with pollen, early stage embryoids, meristems and
somatic embryos.
Advantages and limitations of biolistics.
Requirement of protoplast can be avoided.
Walled intact cells can be penetrated.
Manipulation of genome of subcellular organelles can be achieved.
Limitations
Integration is random.
Requirement of equipments.
Liposome mediated gene transfer
Liposomes are spheres of lipids which can be used to transport molecules into the cells.
These are artificial vesicles that can act as delivery agents for exogenous materials
including transgenes. They are considered as sphere of lipid bilayers surrounding the
molecule to be transported and promote transport after fusing with the cell membrane.
Cationic lipids are those having a positive charge are used for the transfer of nucleic
acid. These liposomes are able to interact with the negatively charged cell membrane
more readily than uncharged liposomes, with the fusion between cationic liposome and
the cell surface resulting in the delivery of the DNA directly across the plasma
membrane. Cationic liposomes can be produced from a number of cationic lipids, e.g.
DOTAP (N-1(- (2,3-dioleoyloxy) propyl)-N,N,N-trimethylammoniumethyl sulphate) and
DOTMA (N-(1-(2,3- dioleoyloxy)propyl)-N,N,N-trimethylammonium chloride) [34]. These
are commercially available lipids that are sold as an in vitro-transfecting agent, as
lipofectin.
Advantages of liposome mediated DNA transfer.
Simplicity.
Long term stability.
Low toxicity.
Protection of nucleic acid from degradation.
Liposomes for use as gene transfer vehicles are prepared by adding an appropriate mix
of bilayer constituents to an aqueous solution of DNA molecules. In this aqueous

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environment, phospholipid hydrophilic heads associate with water while hydrophobic
tails self-associate to exclude water from within the lipid bilayer. This self-organizing
process creates discrete spheres of continuous lipid bilayer membrane enveloping a
small quantity of DNA solution. The liposomes are then ready to be added to target
cells. Germline transgenesis is possible with liposome mediated gene transfer and ES
cells have been successfully transfected by liposomes also.
Calcium phosphate mediated DNA transfer
The process of transfection involves the admixture of isolated DNA (10-100ug) with
solution of calcium chloride and potassium phosphate under condition which allow the
precipitate of calcium phosphate to be formed. Cell s are then incubated with
precipitated DNA either in solution or in tissue culture dish. A fraction of cells will take
up the calcium phosphate DNA precipitate by endocyt osis. Transfection efficiencies
using calcium phosphate can be quite low, in the range of 1-2 %. It can be increased if
very high purity DNA is used and the precipitate allowed to form slowly. Procedures
have been developed where cell taking up exogenous DNA could be upto 20%. This
technique is used for introducing DNA into mammalian cells.
Limitations of calcium phosphate mediated DNA trans fer.
Frequency is very low.
Integrated genes undergo substantial modification.
Many cells do not like having the solid precipitate adhering to them and the surface of
their culture vessel.
Integration with host cell chromosome is random.
Due to above limitations transfection applied to somatic gene therapy is limited.
DNA transfer by DAE-Dextran method
DNA can be transferred with the help of DAE Dextran also. DAE-Dextran may be used
in the transfection medium in which DNA is present. This is polycationic, high
molecular weight substance and convenient for transient assays in cos cells. It does not
appear to be efficient for the production of stable transfectants. If DEAE-Dextran
treatment is coupled with Dimethyl Sulphoxide (DMSO ) shock, then upto 80%
transformed cell can express the transferred gene. It is known that serum inhibits this
transfection so cells are washed nicely to make it serum free.

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Stable expression is very difficult to obtain by th is method. Treatment with
chloroquinine increases transient expression of DNA. The advantage of this method is
that, it is cheap, simple and can be used for transient cells which cannot survive even
short exposure of calcium phosphate.
Transfer of DNA by polycation-DMSO
As calcium phosphate method of DNA transfer is reproducible and efficient but there is
narrow range of optimum conditions. DNA transfer by polycation, polybrene is used to
increase the adsorption of DNA to the cell surface followed by a brief treatment with 25-
30% DMSO to increase membrane permeability and enha nce uptake of DNA. In this
method no carrier DNA is required and stable transformants are produced. This method
works with mouse fibroblast and chick embryo.
Polyethylene glycol mediated transfection
This method is utilized for protoplast only. Polyethylene glycol stimulates endocytosis
and therefore DNA uptake occurs. Protoplasts are ke pt in the solution containing
polyethylene glycol (PEG). The molecular weight of PEG used is 8000 dalton having the
final concentration of 15%. Calcium chloride is added and sucrose and glucose acts as
osmotic buffering agent. To reduce the effects of nuclease present, the carrier DNA from
salmon or herring sperm may also be added. After ex posure of the protoplast to
exogenous DNA in presence of PEG and other chemicals, PEG is allowed to get removed.
Intact surviving protoplasts are then cultured to form cells with walls and colonies in
turn. After several passages in selectable medium f requency of transformation is
calculated. PEGbased vehicles were less toxic and more resistant to nonspecific protein
adsorption making them an attractive alternative for nonviral gene delivery.
Gene transfer through peptide
A variety of peptide sequences are there which are able to bind to, and condense, DNA
to make it more amenable for entry into cells; e.g. the tetrapeptide “Serine - proline-
lysine-lysine” (present on the C- terminus of the histone H1 protein) helps in DNA
transfer. Lysine is a positively charged amino acid. The positively charged lysine amino
acid side chains help to counteract the negatively charged phosphate DNA backbone
and allow the DNA molecules to pack closely to each other. Rational design of peptide
sequences has also been used to develop synthetic DNA binding peptide.

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Tyrosine-lysine-alanine-(Lysine)s-tryptophan-lysine is a peptide which is very effective
to form complexes with DNA. DNA binding peptides that can be coupled to cell specific
ligands can also be synthesized. It allows receptor mediated targeting of the
peptide/DNA complexes to specific cell types.
Gene transfer by retroviruses
The relatively low efficiency of foreign DNA integration into animal cells, combined with
the lack of naturally occurring plasmids, led to the manipulation of viruses as potential
vectors for gene transfer. Retroviruses are found in many species including most
mammals. The genome of retroviruses can be manipula ted to carry exogenous DNA.
The advantages of using retrovirus based vectors ar ise from the stability of the
integration of the viral genome into the host. The integration of a single copy of the viral
DNA at a random location within the host’s genome allows for the long term expression
of the integrated foreign gene. Additionally, retroviruses represent a highly efficient
mechanism for the transfer of DNA into cells. Virus uptake is effective for many somatic
cell lines; however germline cells are infected at low frequency, due to a high level of
mosaicism. Virus can be used for highly developed tissues, such as those of foetuses,
juveniles or adults. This holds great promise in the context of somatic gene therapy.
Retroviral vectors have also been used to introduce transgenes into the ES cell genome.
However, retroviral vectors are limited or problematic in a number of respects. They
exhibit random nature of integration process, which may have deleterious effects on
the host cell, and the general requirement that retrovirus have to infect only dividing
cells.
METHOD OF DNA TRANSFER TO PLANT CELLS
Introduction:
The expression of foreign genes introduced into plants was first achieved in the early
1980s. In the 20 years following these initial successes, we have witnessed a revolution
in plant genetic engineering, with the transformation of well over 100 different plant
species now a routine procedure. Our ability to manipulate the plant genome has come
about through intensive research into vector systems based on the soil bacterium
Agrobacterium tumefaciens and alternative strategies involving direct DNA transfer.
In addition, plant viruses have been developed as versatile episomal vectors for high-
level transient gene expression. This research has immense biotechnological

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implications in the creation of plants with useful genetically engineered characteristics,
such as insect resistance, herbicide tolerance and improved nutritional value.
A fundamental difference between animals and plants is that organized, differentiated
plant tissue shows a high degree of developmental plasticity. An isolated stem segment,
for example, can regenerate into an entire new plan t under appropriate culture
conditions. For most plant species, some form of tissue-culture step is therefore a
prerequisite for the successful production of transgenic plants, i.e. plants carrying the
same foreign DNA sequence (the transgene) in every cell.
Plant callus and cell culture
Callus culture
Tissue culture is the process whereby small pieces of living tissue (explants) are isolated
from an organism and grown aseptically for indefinite periods on a nutrient medium.
For successful plant tissue culture, it is best to start with an explant rich in
undetermined cells, e.g. those from the cortex or meristem, because such cells are
capable of rapid proliferation.
The usual explants are buds, root tips, nodal stem segments or germinating seeds, and
these are placed on suitable culture media where they grow into an undifferentiated
mass known as a callus

Close-up view of a callus culture
Since the nutrient media used for plants can also s upport the growth of
microorganisms, the explants is first washed in a d isinfectant such as sodium
hypochlorite or hydrogen peroxide. Once established, the callus can be propagated
indefinitely by subdivision. For plant cells to develop into a callus, it is essential that
the nutrient medium contains the correct balance of plant hormones (phytohormones),

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i.e. auxins, cytokinins and gibberellins, to maintain the cells in an undifferentiated
state.

The structures of some chemicals which are plant gr owth regulators,
phytohormones
The absolute amounts required vary for different tissue Close-up view of a callus
culture. Explants from different parts of the same plant, and for the same explant from
different species. Thus there is no ideal medium. Most of the media in common use
consist of inorganic salts, trace metals, vitamins, organic nitrogen sources (e.g. glycine),
inositol, sucrose and growth regulators. More complex organic nutrients, such as casein
hydrolysate, coconut water or yeast extract, and a gelling agent are optional extras.
Cell-suspension culture
When callus is transferred into liquid medium and agitated, the cell mass breaks up to
give a suspension of isolated cells, small clusters of cells and much larger aggregates.
Such suspensions can be maintained indefinitely by subculture but, by virtue of the
presence of aggregates, are extremely heterogeneous. Genetic instability adds to this
heterogeneity, so that long-term culture results in the accumulation of mutations
(somaclonal variation), which can adversely affect the vitality and fertility of regenerated
plants. Some species, such as Nicotiana tabacum (tobacco) and Glycine max (soybean),
yield very friable calli, and cell lines obtained from these species are much more
homogeneous, allowing either continuous or batchwise cultivation.
If placed in a suitable medium, isolated single cells from suspension cultures are
capable of division. As with animal cells, conditioned medium may be necessary for
proliferation to occur. Conditioned medium is prepared by culturing high densities of
cells in fresh medium for a few days and then removing the cells by filter sterilization.
Medium conditioned in this way contains essential amino acids, such as glutamine and
serine, as well as growth regulators, such as cytokinins. Provided conditioned medium

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is used, single cells can be plated on to solid media in exactly the same way as
microorganisms, but, instead of forming a colony, plant cells proliferate and form a
callus.
Protoplasts
Protoplasts are cells from which the cellulose walls have been removed. They are very
useful for genetic manipulation because, under certain conditions, protoplasts from
similar or contrasting cell types can be fused to yield somatic hybrids, a process known
as protoplast fusion. Protoplasts can be produced from suspension cultures, callus
tissue or intact tissues, e.g. leaves, by mechanical disruption or, preferably, by
treatment with cellulolytic and pectinolytic enzymes. Pectinase is necessary to break up
cell aggregates into individual cells and the cellulose digests away the cell wall. After
enzyme treatment, protoplast suspensions are collected by centrifugation, washed in
medium without the enzyme, and separated from intact cells and cell debris by flotation
on a cushion of sucrose. When plated on to nutrient medium, protoplasts will
synthesize new cell walls within 5–10 days and then initiate cell division.

Photomicrograph of tobacco protoplasts
Regeneration of fertile plants
The developmental plasticity of plant cells means that whole fertile plants can often be
regenerated from tissue explants, callus, cell suspensions or protoplasts by placing
them on appropriate media. As discussed above, the maintenance of cells in an
undifferentiated state requires the correct balance of phytohormones. However, only

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cytokinin is required for shoot culture and only au xin for root culture, therefore
increasing the level of cytokinins available to the callus induces shoot formation and
increasing the auxin level promotes root formation. Ultimately, plantlets arise through
the development of adventitious roots on shoot buds or through the development of
shoot buds from tissues formed by proliferation at the base of rootlets. The formation of
roots and shoots on callus tissue is known as organogenesis.

Summary of the different cultural manipulations pos sible with plant cells, tissues
and organs.
The culture conditions required to achieve organogenesis vary from species to species,
and have not been determined for every type of callus. The adventitious organogenesis
of shoots and roots can also occur directly from organized plant tissues, such as stem
segments, without first passing through a callus stage.
Under certain conditions, cell suspensions or callus tissue of some plant species can be
induced to undergo a different development process known as somatic embryogenesis.
In this process, the cells undergo a pattern of differentiation similar to that seen in
zygotes after fertilization, to produce embryoids. These structures are embryo-like but

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differ from normal embryos in being produced from s omatic cells and not from the
fusion of two germ cells. The embryoids can develop into fertile plants without the need
to induce root and shoot formation on artificial media. Immature pollen or microspores
can also be induced to form vegetative cells, producing haploid callus or embryoids.
Such cells can be persuaded to undergo diploidizati on by treatment with mitotic
inhibitors. The ease with which plant material is manipulated and interconverted in
culture provides many opportunities for the development of techniques for gene transfer
and the recovery of transgenic plants.
DNA can be introduced into most types of plant mate rial – protoplasts, cell
suspensions, callus, tissue explants, gametes, seeds, zygotes, embryos, organs and
whole plants – so the ability to recover fertile plants from such material is often the
limiting step in plant genetic engineering rather than the DNA transfer process itself. It
is also possible to maintain transformed plant cell lines or tissues (e.g. root cultures)
producing recombinant proteins or metabolites, in the same way that cultured animal
cells can be used as bioreactors for valuable products.
Overview of gene-transfer strategies
Gene transfer to plants is achieved using three different methods. The first exploits the
natural ability of certain bacteria of the genus Agrobacterium to naturally transfer DNA
to the genomes of infected plant cells. This genera lly results in the stable
transformation of the infected cell, and the transferred DNA behaves as a new genetic
locus. Initial limitations with respect to the host range of Agrobacterium prompted
research into alternative methods based on direct DNA transfer. These include the
chemically assisted transformation of protoplasts and the bombardment of plant
material with DNA-coated microprojectiles. Such str ategies can be used for both
transient and stable transformation. Finally, plant viruses can be used as vectors for
gene delivery.
The viruses of plants never integrate into the genome and are not transmitted through
seeds, so stable transformation cannot be achieved. However, plant viruses often cause
systemic infections, resulting in the rapid production of high levels of recombinant
protein throughout the plant, and they can be transmitted through normal infection
routes or by grafting infected scions on to virus-free hosts.
Direct DNA transfer to plants

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Protoplast transformation
Until comparatively recently, the limited host range of A. tumefaciens precluded its use
for the genetic manipulation of a large number of p lant species, including most
monocots. At first, the only alternative to Agrobacterium-mediated transformation was
the introduction of DNA into protoplasts. This process has much in common with the
transfection of animal cells. The protoplasts must initially be persuaded to take up DNA
from their surroundings, after which the DNA integrates stably into the genome in a
proportion of these transfected cells. Gene transfer across the protoplast membrane is
promoted by a number of chemicals, of which polyethylene glycol has become the most
widely used, due to the availability of simple transformation protocols (Negrutiu et al.
1987). Alternatively, DNA uptake may be induced by electroporation, which has also
become a favoured technique (Shillito et al. 1985). As with animal cells, the introduction
of a selectable marker gene along with the transgene of interest is required for the
identification of stable transformants. This can be achieved using plasmid vectors
carrying both the marker and the transgene of interest, but the use of separate vectors
also results in a high frequency of cotransformation (Schocher et al. 1986). Putative
transformants are transferred to selective medium, where surviving protoplasts
regenerate their cell walls and commence cell division, producing a callus. Subsequent
manipulation of the culture conditions then makes it possible to induce shoot and root
development, culminating in the recovery of fertile transgenic plants.
The major limitation of protoplast transformation is not the gene-transfer process itself,
but the ability of the host species to regenerate from protoplasts. A general observation
is that dicots are more amenable than monocots to t his process. In species where
regeneration is possible, an advantage of the technique is that protoplasts can be
cryopreserved and retain their regenerative potential (DiMaio & Shillito 1989).
The first transformation experiments concentrated on species such as tobacco and
petunia in which protoplast-to-plant regeneration is well documented. An early example
is provided by Meyer et al. (1987), who constructed a plasmid vector containing the nptII
marker gene, and a maize complementary DNA (cDNA) e ncoding the enzyme
dihydroquercetin 4- reductase, which is involved in anthocyanin pigment biosynthesis.
The transgene was driven by the strong and constitu tive cauliflower mosaic virus
(CaMV) 35S promoter. Protoplasts of a mutant, white -coloured petunia strain were

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transformed with the recombinant plasmid by electro poration and then selected on
kanamycin-supplemented medium. After a few days, su rviving protoplasts had given
rise to microcalli, which could be induced to regenerate into whole plants. The flowers
produced by these plants were brick-red instead of white, showing that the maize cDNA
had integrated into the genome and was expressed. After successful experiments using
model dicots, protoplast transformation was attempted in monocots, for which no
alternative gene-transfer system was then available. In the first such experiments,
involving wheat (Lorz et al. 1985) and the Italian ryegrass Lolium multiflorum (Potrykus
et al. 1985b), protoplast transformation was achieved and transgenic callus obtained,
but it was not possible to recover transgenic plants. The inability of most monocots to
regenerate from protoplasts may reflect the loss of competence to respond to tissue-
culture conditions as the cells differentiate. In cereals and grasses, this has been
addressed to a certain extent by using embryogenic suspension cultures as a source of
protoplasts. Additionally, since many monocot speci es are naturally tolerant to
kanamycin, the nptII marker used in the initial experiments was replaced with
alternative markers conferring resistance to hygromycin or phosphinothricin. With
these modifications, it has been possible to regenerate transgenic plants representing
certain varieties of rice and maize with reasonable efficiency (Shimamoto et al. 1988,
Datta et al. 1990, Omirulleh et al. 1993). However, the extended tissue-culture step is
unfavourable, often resulting in sterility and other phenotypic abnormalities in the
regenerated plants. The transformation of protoplasts derived from stomatal guard cells
has recently been identified as an efficient and genotype-independent method for the
production of transgenic sugar beet (Hall et al. 1996).
Particle bombardment
An alternative procedure for plant transformation was introduced in 1987, involving the
use of a modified shotgun to accelerate small (1–4 µm) metal particles into plant cells at
a velocity sufficient to penetrate the cell wall (~250 m/s). In the initial test system,
intact onion epidermis was bombarded with tungsten particles coated in tobacco mosaic
virus (TMV) RNA. Three days after bombardment, approximately 40% of the onion cells
that contained particles also showed evidence of TMV replication (Sanford et al. 1987). A
plasmid containing the cat reporter gene driven by the CaMV 35S promoter was t hen
tested to determine whether DNA could be delivered by the same method. Analysis of

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the epidermal tissue 3 days after bombardment revea led high levels of transient
chloramphenicol transacetylase (CAT) activity (Klein et al. 1987).
The stable transformation of explants from several plant species was achieved soon
after these initial experiments. Early reports included the transformation of soybean
(Christou et al. 1988), tobacco (Klein et al. 1988b) and maize (Klein et al. 1988a). In
each case, the nptII gene was used as a selectable marker and transforma tion was
confirmed by the survival of callus tissue on kanamycin-supplemented medium. The
ability to stably transform plant cells by this method offered the exciting possibility of
generating transgenic plants representing species that were, at the time, intractable to
other transformation procedures. In the first such report, transgenic soybean plants
were produced from meristem tissue isolated from im mature seeds (McCabe et al.
1988). In this experiment, the screenable marker gene gusA was introduced by particle
bombardment and transgenic plants were recovered in the absence of selection by
screening for β-glucuronidase (GUS) activity.
Other early successes included cotton, papaya, maize and tobacco (Finer & McMullen
1990, Fitch et al. 1990, Fromm et al. 1990, Gordon- Kamm et al. 1990, Tomes et al.
1990). There appears to be no intrinsic limitation to the scope of this procedure, since
DNA delivery is governed entirely by physical parameters. Many different types of plant
material have been used as transformation targets, including callus, cell suspension
cultures and organized tissues, such as immature embryos, meristems and leaves. The
number of species in which transgenic plants can be produced using variants of particle
bombardment has therefore increased dramatically ov er the last 10 years. Notable
successes include almost all of the commercially important cereals, i.e. rice (Christou et
al. 1991), wheat (Vasil et al. 1992), oat (Somers et al. 1992, Torbert et al. 1995), sugar
cane (Bower & Birch 1992) and barley (Wan & Lemaux 1994, Hagio et al. 1995).
The original gunpowder-driven device has been impro ved and modified, resulting in
greater control over particle velocity and hence greater reproducibility of transformation
conditions. An apparatus based on electric discharge (McCabe & Christou 1993) has
been particularly useful for the development of var iety-independent gene-transfer
methods for the more recalcitrant cereals and legumes.
Several instruments have been developed where particle acceleration is controlled by
pressurized gas. These include a pneumatic apparatu s (Iida et al. 1990), a ‘particle

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inflow gun’ using flowing helium (Finer et al. 1992, Takeuchi et al. 1992) and a device
utilizing compressed helium (Sanford et al. 1991).
Physical parameters, such as particle size and acceleration (which affect the depth of
penetration and the amount of tissue damage), as well as the amount and conformation
of the DNA used to coat the particles, must be optimized for each species and type of
explant. However, the nature of the transformation target is probably the most
important single variable in the success of gene transfer. The pretreatment of explants
with an osmoticum has often been shown to improve t ransformation efficiency,
probably by preventing the deflection of particles by films or droplets of water. Factors
influencing the success of gene transfer by particle bombardment have been extensively
reviewed (Sanford et al. 1993, Birch & Bower 1994).
Other direct DNA-transfer methods
There is a great diversity of approaches for gene transfer to animal cells and many of
the same methods have been attempted in plants. Ele ctroporation has been used to
transform not only protoplasts (see above) but also walled plant cells, either growing in
suspension or as part of intact tissues. In many cases, the target cells have been
wounded or pretreated with enzymes in order to facilitate gene transfer (e.g. D’Halluin et
al. 1992, Laursen et al. 1994). It has been shown; however, that immature rice, wheat
and maize embryos can be transformed using electrop oration without any form of
pretreatment (Kloti et al. 1993, Xu & Li 1994).
Other transformation methods also involve perforation of the cell, including the use of
silicon carbide whiskers (Thompson et al. 1995, Nagatani et al. 1997), ultrasound
(Zhang et al. 1991) or a finely focused laser beam (Hoffman 1996). In most of these
cases, only transient expression of the introduced DNA has been achieved, although
transgenic maize plants have been recovered followi ng whisker-mediated
transformation.
Finally, microinjection has been used to introduce DNA directly into the fertilized eggs
of many animals. In plants, microinjection of DNA into zygotes may also be the most
direct way to produce transgenics, but so far the technique is inefficient and not widely
used (Leduc et al. 1996, Holm et al. 2000).
In planta transformation

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Until recently, gene transfer to plants involved the use of cells or explants as
transformation targets and an obligatory tissue-culture step was needed for the
regeneration of whole fertile plants. Experiments using the model dicot Arabidopsis
thaliana have led the way in the development of so-called in planta transformation
techniques, where the need for tissue culture is minimized or eliminated altogether.
Such methods involve the introduction of DNA, either by Agrobacterium or by direct
transfer, into intact plants. The procedure is carried out at an appropriate time in the
plant’s life cycle, so that the DNA becomes incorporated into cells that will contribute to
the germ line, directly into the germ cells themselves (often at around the time of
fertilization) or into the very early plant embryo. Generally, in planta transformation
methods have a very low efficiency, so the small size of Arabidopsis and its ability to
produce over 10 000 seeds per plant is advantageous . This limitation has so far
prevented in planta techniques from being widely adopted for other plant species.
The first in planta transformation system involved imbibing Arabidopsis seeds overnight
in an Agrobacterium culture, followed by germination (Feldmann & Marks 1987). A large
number of transgenic plants containing T-DNA insertions were recovered, but in general
this technique has a low reproducibility. Bechtold et al. (1993) has described a more
reliable method, in which the bacteria are vacuuminfiltrated into Arabidopsis flowers.
An even simpler technique called floral dip has become widely used (Clough & Bent
1998). This involves simply dipping
Arabidopsis flowers into a bacterial suspension at the time of fertilization. In both these
methods, the transformed plants are chimeric, but g ive rise to a small number of
transgenic progeny (typically about 10 per plant). Similar approaches using direct
DNA transfers have been tried in other species, but germ-line transformation has not
been reproducible.
For example, naked DNA has been injected into the floral tillers of rye plants (De La
Pena et al. 1987) and post-fertilization cotton flowers (Zhou et al. 1983), resulting in the
recovery of some transgenic plants. Transgenic tobacco has been produced following
particle bombardment of pollen (Touraev et al. 1997).
An alternative to the direct transformation of germline tissue is the introduction of DNA
into meristems in planta, followed by the growth of transgenic shoots. In Arabidopsis,
this has been achieved simply by severing apical shoots at their bases and inoculating

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the cut tissue with Agrobacterium suspension (Chang et al. 1994). Using this procedure,
transgenic plants were recovered from the transformed shoots at a frequency of about
5%. In rice, explanted meristem tissue has been transformed using Agrobacterium and
particle bombardment, resulting in the proliferation of shoots that can be regenerated
into transgenic plants (Park et al. 1996). Such procedures require only a limited amount
of tissue culture.
Chloroplast transformation
So far, we have exclusively considered DNA transfer to the plant’s nuclear genome.
However, the chloroplast is also a useful target for genetic manipulation, because
thousands of chloroplasts may be present in photosynthetic cells and this can result in
levels of transgene expression up to 50 times highe r than possible using nuclear
transformation. Furthermore, transgenes integrated into chloroplast DNA do not appear
to undergo silencing or suffer from position effects that can influence the expression
levels of transgenes in the nuclear DNA.
Chloroplast transformation also provides a natural containment method for transgenic
plants, since the transgene cannot be transmitted through pollen (reviewed by Maliga
1993).
The first reports of chloroplast transformation were serendipitous, and the integration
events were found to be unstable. For example, an early experiment in which tobacco
protoplasts were cocultivated with Agrobacterium resulted in the recovery of one
transgenic plant line, in which the transgene was transmitted maternally. Southern-blot
analysis of chloroplast DNA showed directly that th e foreign DNA had become
integrated into the chloroplast genome (De Block et al. 1985). However, Agrobacterium
does not appear to be an optimal system for chlorop last transformation, probably
because the T-DNA complex is targeted to the nucleus. Therefore, direct DNA transfer
has been explored as an alternative strategy. Stable chloroplast transformation was first
achieved in the alga Chlamydomonas reinhardtii, which has a single large chloroplast
occupying most of the volume of the cell (Boynton et al. 1988).
Particle bombardment was used in this experiment. T he principles established using
this simple organism was extended to tobacco, allow ing the recovery of stable
transplastomic tobacco plants (Svab et al. 1990b). These principles included the use of
vectors containing chloroplast homology regions, allowing targeted integration into the

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chloroplast genome, and the use of the selectable marker gene aad (aminoglycoside
adenyltransferase), which confers resistance to str eptomycin and spectinomycin
(Zoubenko et al. 1994). Efficient chloroplast transformation has been achieved both
through particle bombardment (e.g. Staub & Maliga 1 992) and polyethylene glycol
(PEG)-mediated transformation (Golds et al. 1993, Koop et al. 1996). The use of a
combined selectable– screenable marker (aad linked to the gene for green fluorescent
protein) allows the tracking of transplastomic sectors of plant tissue prior to chlorophyll
synthesis, so that transformed plants can be rapidly identified (Khan & Maliga 1999).
A procedure used to make a transgenic plant.

(A) Outline of the process. A disc is cut out of a leaf and incubated in culture with
Agrobacteria that carry a recombinant plasmid with both a selectable marker and a
desired transgene. The wounded cells at the edge of the disc release substances that
attract the Agrobacteria and cause them to inject DNA into these cells. Only those plant
cells that take up the appropriate DNA and express the selectable marker gene survive
to proliferate and form a callus. The manipulation of growth factors supplied to the
callus induces it to form shoots that subsequently root and grow into adult plants
carrying the transgene.

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(B) The preparation of the recombinant plasmid and its transfer to plant cells. An
Agrobacterium plasmid that normally carries the T-DNA sequence is modified by
substituting a selectable marker (such as the kanamycin-resistance gene) and a desired
transgene between the 25-nucleotide-pair T-DNA repe ats. When the Agrobacterium
recognizes a plant cell, it efficiently passes a DNA strand that carries these sequences
into the plant cell, using the special machinery that normally transfers the plasmid's T-
DNA sequence
TRANSFORMATION EFFICIENCY.
The transformation efficiency, which gives you an indication of how effective you were in
getting DNA molecules into bacterial cells. Transformation efficiency is a number. It
represents the total number of bacterial cells that express the green protein, divided by
the amount of DNA used in the experiment.
It gives us the total number of bacterial cells transformed by one microgram of DNA.)
The transformation efficiency is calculated using the following formula:

TRANSFECTION EFFICIENCY
The following procedure may be used to determine th e percentage of stable
transfectants obtained.
Note: The stained cells will not be viable after this procedure.

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Materials Required:
Methylene blue
Methanol
Cold deionized water
Light microscope
1.
After approximately 14 days of selection in the appropriate drug, monitor the cultures
microscopically for the presence of viable cell clones. When distinct “islands” of
surviving cells are visible and nontransfected cells have died out, proceed with
2.
Prepare stain containing 2% methylene blue in 50–70% methanol.
3.
Remove the growth medium from cells by aspiration.
4.
Add to cells sufficient stain to cover the bottom of the dish.
5.
Incubate for 5 minutes.
6.
Remove the stain, and rinse gently under deionized cold water. Shake off excess
moisture.
7.
Allow the plates to air-dry. The plates can be stored at room temperature.
8.
Count the number of colonies, and calculate the percent of transfectants based on the
cell dilution and original cell number.

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UNIT : 5 LABELLING AND DETECTION TECHNIQUES
Labeling of DNA, RNA and proteins by radioactive is otopes, non-radioactive
labelling, in vivo labeling, autoradiography and autofluorography. DNA sequencing
by enzymatic and chemical methods, Agarose gel elec trophoresis, PAGE, PFGE.
Methods of nucleic acid hybridization; southern, no rthern and western blotting
techniques.
LABELING OF DNA
In order to visualize where the DNA hybrids are forming on the blots, the probe DNA
must be labelled using either radioactively labelled (
32
P ) or chemically substituted
nucleotides. When radioactivity is used, autoradiography using X-ray film is employed
to visualize the hybrid positions. When chemically labelled probes are used, colorimetric
reactions are most often used, some relying on antibodies or other chemicals attached
to enzymes that can cause a coloured precipitate to form from an appropriate substrate.
In vivo labelling: Although, in principle, DNA and RNA can be labelled in vivo, by
supplying labelled deoxynucleotides to tissue culture cells, this procedure is of limited
general use; it has been restricted largely to prepare labelled viral DNA from virus-
infected cells, and studying RNA processing events.

In vitro labelling: A much more versatile method involves in vitro labelling: the purified
DNA, RNA or oligonucleotide is labelled in vitro by using a suitable enzyme to
incorporate labelled nucleotides. DNA and RNA can conveniently be labelled in vitro by
incorporation of nucleotides (or nucleotide components) containing a labelled atom or
chemical group.
There are four common ways to label DNA:
1.
End-labeling, either at the 3'ends with DNA polyme rase or at the 5' end using T4
polynucleotide kinase;
2.
Nick translation, using DNA polymerase and a low concentration of DNase I to form the
nicks that are filled in by the polymerase);
3.
Random primer labeling, where DNA polymerase is use d in conjunction with random
hexanucleotides which prime the polymerization reactions; and
4.
PCR labeling.

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1. End-labelling: It does not result in highly labelled probes but is still used for certain
procedures. "End labelling" in which the end of a DNA (or RNA) molecule is specifically
labelled. There are methods for labelling either the 5' end or 3' end specifically.
What is most common is 5' end labelling with P
32
γ ATP, and the enzyme polynucleotide
kinase. The terminal phosphate (the "hot" one) is transferred to the 5' end of the
molecule. Note that only one marked residue is incorporated by this method, so the
specific activity of the label (radioactive counts per minute per microgram of DNA) is
lower than in the aforementioned two methods.

It is less widely used, but is useful for a number of procedures, including labelling of
single-stranded oligonucleotides, and restriction mapping. Inevitably, because only one
or a very few labelled groups are incorporated, the specific activity (the amount of
radioactivity incorporated divided by the total mass) of the labelled DNA is much less
than that for probes in which there has been incorp oration of several labelled
nucleotides along the length of the DNA.

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Larger DNA fragments can be end-labelled by various alternative methods. Fill-in end-
labelling (Figure B) is one popular approach, and uses the Klenow subunit of E.
coli DNA polymerase. The DNA of interest is cleaved with a suitable restriction nuclease
to generate 5 overhangs. The overhangs act as a primer for Klenow DNA polymerase to
incorporate labelled nucleotides complementary to the overhang.
2.
Nick translation: The nick-translation procedure involves introducing single-strand
breaks (nicks) in the DNA, leaving exposed 3
hydroxyl termini and 5 phosphate termini.
The nicking can be achieved by adding a suitable en donuclease such as pancreatic
deoxyribonuclease I (DNase I).
The exposed nick can then serve as a start point for introducing new nucleotides at the
3hydroxyl side of the nick using the DNA polymerase activity of E. coli DNA polymerase
I at the same time as existing nucleotides are removed from the other side of the nick by
the 5 3 exonuclease activity of the same enzyme. As a result, the nick will be moved
progressively along the DNA ('translated') in the 5 3 direction. If the reaction is
carried out at a relatively low temperature (about 15° C), the reaction proceeds no
further than one complete renewal of the existing nucleotide sequence. Although there
is no net DNA synthesis at these temperatures, the synthesis reaction allows the
incorporation of labelled nucleotides in place of the previously existing unlabeled ones.
Nick translation of DNA results in highly labelled probes, but the DNase I
concentrations are critical to the proper labelling and utilization of the probe.

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3. Random primed DNA labelling:


The random primed DNA labelling method (sometimes k nown as oligolabelling) is based
on hybridization of a mixture of all possible hexanucleotides: the starting DNA is
denatured and then cooled slowly so that the individual hexanucleotides can bind to
suitably complementary sequences within the DNA strands.
Synthesis of new complementary DNA strands is primed by the bound hexanucleotides
and is catalyzed by the Klenow subunit of DNA polym erase I (which contains the
polymerase activity in the absence of associated exonuclease activities). DNA synthesis
occurs in the presence of the four dNTPs, at least one of which has a labelled group.
This method produces labelled DNAs of high specific activity. Because all sequence
combinations are represented in the hexanucleotide mixture, binding of primer to
template DNA occurs in a random manner, and labelling is uniform across the length of
the DNA.
4.
PCR Labelling:
Two single stranded DNA primers (18-30 bp long), one forward and one reverse are
synthesized. The primers usually match a known DNA sequence and are used to
amplify the fragment in-between.
After adding the primers, the Taq polymerase (or other thermostable polymerase), the

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buffer and the DNA template, the reaction mix is denatured by heating 30
0
-60
0
seconds
at 94
0
C (denaturing step). Then, the temperature is dropped to 50
0
-60
0
C for 30-60
seconds, to allow the primers to anneal to their target sequences (annealing step).
Then the temperature is raised to 68
0
-72
0
C (optimal temperature for Taq polymerase)
for 0.5-4 minutes, to allow the enzyme to synthesize the new DNA strands (extension
step). These temperature steps are repeated again (usua lly 30 cycles), allowing
exponential amplification of the DNA molecule between the two primers. If part of the
dTTP in the reaction is replaced by labelled fluorescent dyes or haptenes, PCR can be
used to label the newly synthesized DNA molecules with fluorescent dyes or haptenes.
LABELING OF RNA
The preparation of labeled RNA probes (riboprobes) is most easily achieved by in
vitro transcription of insert DNA cloned in a suitable plasmid vector. The vector is
designed so that adjacent to the multiple cloning site is a phage promoter sequence,
which can be recognized by the corresponding phage RNA polymerase. For example, the
plasmid vector pSP64 contains the bacteriophage SP6 promoter sequence immediately
adjacent to a multiple cloning site. The SP6 RNA po lymerase can then be used to
initiate transcription from a specific start point in the SP6 promoter sequence,
transcribing through any DNA sequence that has been inserted into the multiple
cloning site.
By using a mix of NTPs, at least one of which is la beled, high specific activity
radiolabeled transcripts can be generated. Bacteriophage T3 and T7 promoter/RNA
polymerase systems are also used commonly for generating riboprobes. Labeled sense
and antisense riboprobes can be generated from any gene cloned in such vectors (the
gene can be cloned in either of the two orientations) and are widely used in tissue in
situ hybridization.
Nucleic acids may be modified with tags that enable detection or purification. The
resulting nucleic acid probes can be used to identify or recover other interacting
molecules. Common labels used to generate nucleic a cid probes include radioactive
phosphates, biotin, fluorophores and enzymes. In addition, the bioconjugation methods
used for nucleic acid probe generation may be adapted for attaching nucleic acids to
other molecules or surfaces to facilitate targeted delivery or immobilization, respectively.
Nucleic acid probes can be labeled with tags or other modifications during synthesis.

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Enzymatic methods for nucleic acid labeling
Terminal Deoxynucleotidyl Transferase
Terminal deoxynucleotidyl transferase (TdT) is a DNA polymerase enzyme expressed in
certain populations of lymphoid cells. TdT typically adds numerous deoxynucleotides to
the 3′ terminus of a DNA strand. TdT is template-independent and not significantly
affected by DNA sequence, but DNA structure is important. TdT has the highest activity
towards the 3′ end of single-stranded DNA but can a lso modify the 3′ overhang of
double-stranded DNA with lower efficiency. TdT has poor activity towards double-
stranded DNA with blunt ends or 5′ overhangs. TdT is often used to label DNA probes
for RACE (Rapid Amplification of cDNA Ends), TUNEL (Terminal deoxynucleotidyl
transferase dUTP Nick-End Labeling) assays and as a method for adding 3′ overhangs to
DNA fragments to facilitate cloning.
TdT can also be used to label the 3′ end of DNA pro bes with radioactive and
nonradioactive tags for a variety of detection and affinity applications.
T4 RNA ligase

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T4 RNA ligase is an enzyme coded for in the genome of the T4 bacteriophage. T4 RNA
ligase catalyzes the attachment of a terminal 5′-phosphate to a terminal 3′-hydroxyl
group on RNA. T4 RNA ligase is template-independent but requires single-stranded RNA
and ATP.
T4 RNA ligase is used for labeling the 3′ end of RNA with [5′ ³²P]pCp (cytidine-3',5'-bis-
phosphate), modifying mRNA for cDNA library generation and performing 5′-RACE.
T4 RNA ligase can also be used to 3′ end-label RNA with nonradioactive tags using an
appropriately modified nucleoside 3',5'-bisphosphate.
T4 polynucleotide kinase (PNK)
T4 Polynucleotide Kinase (T4 PNK) is an enzyme code d for in the genome of the T4
bacteriophage. T4 PNK transfers an organic phosphate from the gamma position on ATP
to the 5′-hydroxyl group of DNA and RNA. The wild-t ype enzyme also has 3′-
phosphatase activity. T4 PNK is used primarily for labeling the 5′ ends of
polynucleotides with radioactive phosphate from isotope-modified ATP. PNK is more
efficient at modifying short overhangs and blunt-end fragments than TdT or T4 RNA
ligase. While it is possible to perform phosphate-exchange reactions, PNK labeling is
most efficient when the 5′ end of the target molecule has been dephosphorylated.
DNA polymerase
DNA polymerases are a family of enzymes that create deoxyribonucleic acid polymers by
catalyzing the joining of the 5′-phosphorylated end of a deoxyribonucleotide (monomer)
to the 3′-hydroxyl end of an existing DNA strand (DNA elongation) or primer (primer
extension). DNA polymerases are template-dependent but not sequence-dependent. To
synthesize DNA, the 3′-OH end of an existing DNA st rand must be annealed to a
complementary strand of DNA. The DNA polymerase wil l synthesize a new DNA strand
through elongation of the existing 3′-OH end, addin g individual nucleotides
complementary to the template strand being read.
Probes generated with the aid of DNA polymerase are most commonly made by the
random incorporation of modified nucleotides during the DNA replication process,
which can be done by PCR or simple primer extension reactions. Probes generated in

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this manner have high specific activity and allow the detection of small quantities of
target. Traditionally, this method required radioactive nucleotides to generate probes;
however, biotin, fluorophores and other nonradioact ive tags can be used if the
modification does not interfere with the polymerase elongation reaction (except for
terminator sequencing applications).
RNA polymerase
RNA polymerases are a family of enzymes that create ribonucleic acid polymers by
catalyzing the joining of the 5′ phosphorylated-end of a ribonucleotide (monomer) to the
3′-hydroxyl end of a previously incorporated ribonucleotide. RNA polymerases are
template- and sequence-dependent, requiring a promoter sequence within the template
DNA in order to initiate binding of the enzyme and, depending on the host system;
various cofactors are required for RNA transcription to proceed on the single-stranded
template.
RNA polymerases are used for a variety of lab purposes from the in vitro synthesis of
mRNA to the generation of probes for hybridization and binding assays. Probes
generated with the aid of RNA polymerase are made b y the random incorporation of
modified nucleotides during the transcription process. RNA probes generated in this
manner have high specific activity and are most com monly made with radioactive
nucleotides, although biotin and other tags are also available.
Chemical methods for nucleic acid labeling
Periodate oxidation of RNA
Meta- and ortho-periodates (IO4 and IO6, respectively) are anions formed from iodine
and oxygen and commonly found as salts with potassi um (e.g. KIO4) or sodium (e.g.
NaIO
4).

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EDC activation of 5′ phosphate
In solution, periodate cleaves the bonds between ad jacent carbon atoms having
hydroxyl groups (vicinal diols or cis-glycols), creating two aldehydes groups. The
resulting aldehyde groups are spontaneously reactive toward primary amine-containing
molecules and surfaces. Aldehydes can be used in two types of coupling reactions with
either primary amine- or hydrazide-activated tags. Primary amines react with aldehydes
to form Schiff bases, which readily hydrolyze and must be stabilized through their
reduction to secondary amine bonds with sodium cyanoborohydride (NaBH
4).
Hydrazide-modified molecules also spontaneously react with aldehydes but form fairly
stable hydrazone linkages, making the reaction much more efficient.
Carbodiimides are functional groups (RN=C=NR) that are typically used in organic
synthesis to activate the formation of amide or phosphoramidate linkages between
primary amines (RNH
2) and carboxylate (RCOOR′)- or phosphate (R-PO 4)-containing
molecules, respectively.

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Unlike most crosslinkers, carbodiimides do not beco me part of the final crosslink
between the molecules and thus do not add any addit ional chemical structure to the
resulting products.
Because the 5′ phosphate is required, synthetic oligonucleotides must first be treated
with a kinase. With that minor exception, EDC-media ted conjugations are an
economical means for coupling both RNA and DNA to n early any other primary amine-
containing molecule or surface.

Chemical random-labeling
Random chemical labeling of nucleic acids can be ac complished by various means.
Because these methods label at random sites along t he length of a DNA or RNA
molecule, they allow a higher degree of labeling to be achieved than end-labeling
techniques. However, one disadvantage of these methods is that the nucleotide bases
are directly modified, which will reduce or prevent base-pairing between complementary
strands during hybridization experiments. Therefore, it may be necessary to balance the
degree of labeling with the hybridization efficiency of the probes in certain experiments.
Two types of photoreactive label reagents are used for nucleic acids: phenylazide- and
psoralen-based. When the phenylazide functional group is exposed to UV light, it forms

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a labile nitrene that can insert nonspecifically into double bonds and C-H and N-H sites
via addition reactions, provided that more reactive nucleophilic (e.g., primary amines)
are not present.

Molecules containing the psoralen functional group can be used to label double-
stranded DNA or RNA. The psoralen ring structure ef fectively intercalates into the
double-stranded portions, and exposure to UV light causes a cyclo-addition product to
be formed with the 5,6-double bond in thymine residues.
Interaction of psoralen with DNA to form 2 types of monoadducts (A+B) or a diadduct
(C). The formation of the crosslink requires UV absorption events at each reaction.

PROTEINS BY RADIOACTIVE ISOTOPES, NON-RADIOACTIVE L ABELLING
Protein Labelling:
Biological research often requires the use of molecular labels that are covalently
attached to a protein of interest to facilitate detection or purification of the labelled

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protein and/or its binding partners. Labelling stra tegies result in the covalent
attachment of different molecules, including biotin, reporter enzymes, fluorophores and
radioactive isotopes, to the target protein or nucleotide sequence.
Protein labelling is performed using many different methods and can be used for in-
vivo, in-situ, in-vitro and ex-vivo sample analysis.
Protein labels:
1. Biotin labelling:
Biotin is a useful label for protein detection, purification and immobilization because of
its extraordinarily strong binding to avidin, streptavidin or NeutrAvidin Protein. Indeed,
this interaction is one of the strongest non-covalent interactions between a protein and
ligand. Additionally, biotin is considerably smaller than enzyme labels (244.3 Da) and is
therefore less likely to interfere with normal protein function.
Biotinylation is the process of labeling proteins or nucleotides with biotin molecules and
can be performed by enzymatic and chemical means. Chemical methods of biotinylation
are most commonly used. They are composed of the biotinyl group, a spacer arm and a
reactive group that is responsible for attachment to target functional groups on
proteins.
The biotin-avidin interaction is commonly exploited to detect and/or purify proteins
because of the high specificity that these two mole cules have for each other.
Biotinylation reagents are available for targeting specific functional groups or residues,
including primary amines, sulfhydryls, carboxyls and carbohydrates.
Avidin-Biotin Interaction:
The interaction between biotin (vitamin H) and avidin is a useful tool in nonradioactive
methods of purification, detection, immobilization, labelling, viral vector-targeting and
drug targeting systems. The extraordinary affinity of avidin for biotin is one the
strongest known non-covalent interactions of a protein and ligand (Ka=10
15
M
-1
) and
allows biotin-containing molecules in a complex mixture to be discretely bound with

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avidin conjugates. The bond formation between biotin and avidin is very rapid, and
once formed, it is unaffected by extremes in pH, temperature, organic solvents and
other denaturing agents.

The interaction of biotin and avidin or streptavidin has been exploited for use in many
protein and nucleic acid detection and purification methods. Because the biotin label is
stable and small, it rarely interferes with the function of labelled molecules enabling the
avidin-biotin interaction to be used for the development of robust and highly sensitive
assays.

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Proteins that are biotin-labelled (i.e., biotinylated) are routinely detected or purified with
avidin conjugates in many protein research applications, including the enzyme-linked
immunosorbant assay (ELISA), Western blot analysis, immunohistochemistry (IHC),
immunoprecipitation (IP) and other methods of affin ity purification, cell surface
labelling and flow cytometry/fluorescence-activated cell sorting (FACS).
Besides a strong affinity for avidin, biotin exhibits two characteristics that make the
molecule ideal for labelling proteins and macromolecules.
First, biotin is comparatively smaller than globular proteins, which minimizes any
significant interference in many proteins and allows multiple biotin molecules to be
conjugated to a single protein for maximum detection by avidin. Second, as shown in
the diagram below, biotin has a valeric acid side chain that is easily derivatized and
conjugated to reactive moieties and chemical structures without affecting its avidin-
binding function. This feature allows many useful biotinylation reagents to be created.
Biocytin is a derivative of biotin found in serum and urine that has an added lysine
group coupled at the ε-amino acid side chain to the valeric acid side chain. As shown
below, biocytin is longer than biotin, which makes the molecule useful in making long-
chain biotinylation reagents. Biocytin can also be used to make trifunctional
crosslinking reagents because of the free carboxylate group and α-amine.

2.
Enzymes:
Certain enzymes have properties that enable them to function as highly sensitive probes
with a long shelf life and versatility for the detection of proteins in tissues, whole cells or
lysates. Enzyme labels are considerably larger than biotin and require the addition of a

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substrate to generate a chromogenic, chemiluminescent or fluorescent signal that can
be detected by different approaches. Enzyme labels are widely used because of their
multiple types of signal output, signal amplification and the wide selection of enzyme-
labelled products, especially antibodies.
Enzymes commonly used as labels include horseradish peroxidase (HRP), alkaline
phosphatase(AP), glucose oxidase and β-galactosidase, and specific substrates are
available for each enzyme. Indeed, multiple commercial substrates are available for HRP
and AP that generate colorimetric, chemiluminescent or fluorescent signal outputs.
Enzyme probes can be conjugated to antibodies, streptavidin or other target proteins by
multiple mechanisms, including glutaraldehyde, redu ctive amination following
periodate oxidation of sugars to reactive aldehydes.

The benefits of using enzyme probes (also called reporters) to detect a target protein are
three-fold:
a.
High sensitivity – The signal output can be easily detected, and th erefore low
concentrations of target proteins can be identified. Additionally, methods of signal
amplification are available that signficantly increase the number of enzyme molecules to
the site of the target protein. Finally, enzyme reporters exhibit rapid turnover, which
increases the amount of substrate that a single enzyme converts during a given unit of
time.

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b. Long shelf life – The enzymes are quite stable when stored properly, and while the
enzyme substrate is light-sensitive, the enzyme itself is not sensitive to degradation by
ambient light.
c.
Output versatility – Substrates that yield either chromogenic, chemiluminescent or
fluorescent output are available for the most common enzyme probes.
Limitations of enzyme probes:
a.
Large size – Enzyme reporters are considerably larger than or ganic fluorescent
compounds (e.g., FITC, TRITC, AMCA) and therefore m ay interfere with the biological
function of proteins to which they are conjugated.
b.
Substrate requirement – Enzyme probes require the addition of a substrate for protein
detection, and depending on the substrate used, thi s reaction can be sensitive to
environmental conditions (e.g., light, temperature) and ambient light.
c.
Endogenous interference – The enzymes used to detect target proteins in a sample are
often expressed in the experimental system used (e.g., tissues, cells), which will also
process the substrate and yield nonspecific background signal unless inhibited.
3.
Fluorescent Probes
Fluorescent molecules, also called fluorophores or simply fluors, respond directly and
distinctly to light and produce a detectable signal. Unlike enzymes or biotin, fluorescent
labels do not require additional reagents for detection. This feature makes fluorophores
extremely versatile and the new standard in detecting protein location and activation,
identifying protein complex formation and conformational changes, and monitoring
biological processes in vivo.
Fluorophores can be divided into three general groups, and each group of probes has
distinct characteristics. These groups are as follows:
Organic dyes – FITC, TRITC, DyLight Fluors
Biological fluorophores – Green fluorescent protein (GFP), R-Phycoerythrin
Quantum dots

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Detection of fluorescent probes requires specialized equipment, including an excitation
light source, filter set and a detector, that are found in fluorescence microscopes,
fluorescence plate-readers, flow cytometers and cell sorters. This equipment enables the
absolute quantitation of proteins based on fluorescence, which is a significant benefit to
using fluorescent probes over other types of probes.
Each fluorophore has distinct characteristics, which should be considered when
deciding which fluorophore to use for a given application or experimental system.
Organic dyes
Synthetic organic dyes, such as fluorescein, were the first fluorescent compounds used
in biological research. Derivatives of these original compounds have been produced to
improve their photostability and solubility. These dyes are also derivitized to use in
bioconjugation, especially fluorescein isothiocyanate (FITC), rhodamine (tetramethyl
rhodamine isothiocyanate, TRITC) and commercial variants with greater performance.
The small size of these fluors is a benefit over biological fluorophores for bioconjugation
strategies because they can be crosslinked to macromolecules, such as antibodies,
biotin or avidin, without interfering with proper b iological function. Biological
fluorophores
While bioluminescence has been known for millenia, the first use of a biological
fluorophore for research applications occurred in the 1990s, when green fluorescent
protein (GFP) was cloned from the jellyfish Aequorea victoria and used as a gene
expression reporter. Since that time, derivatives of the original GFP, phycobiliproteins
(allophycocyanin, phycocyanin, phycoerythrin and phycoerythrocyanin) and many other
proteins have been designed for use in biological expression systems, and their use is
now commonplace in biological research.
The benefit of these types of fluorophores is that expression plasmids can be introduced
into either bacteria, cells, organs or whole organisms, to drive expression of that
fluorophore either alone or fused to a protein of interest in the context of the biological
processes studied.

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The use of fluorescent proteins can be time consuming, and expressing large amounts
of light-producing proteins can cause reactive oxygen species and induce artifactual
responses or toxicity. Additionally, the size of the fluorescent protein can change the
normal biological function of the cellular protein to which the fluorophore is fused, and
biological fluorophores do not typically provide the level of photostability and sensitivity
offered by synthetic fluorescent dyes.
Quantum dots
Quantum dots are nanocrystals with unique chemical properties that provide tight
control over the spectral characteristics of the fluor. Quantum dots were developed in
the 1980s and since the 1990s have been increasingly used in fluorescence applications
in biological research. Quantum dots are nanoscale-sized (2-50nm) semiconductors
that, when excited, emit fluorescence at a wavelength based on the size of the particle;
smaller quantum dots emit higher energy than large quantum dots, and therefore the
emitted light shifts from blue to red as the size of the nanocrystal increases. And
because quantum dot size can be tightly controlled, there is greater specificity for
distinct excitation and emission wavelengths than other fluors.
Quantum dots have also been reported to be more photostable than other fluorophores,
as one report showed that quantum dots remained flu orescent for 4 months in an in
vivo imaging study. Additionally, quantum dots can be coated for use in different
biological applications such as protein labeling. While the use of quantum dots in
biological applications is increasing, there are reports of cell toxicity in response to the
breakdown of the particles and their use can be cost-prohibitive.
Fluorescence Detection strategies
1.
Fluorescent microscopes – detect localized fluors in samples in both two and three
dimensions
2.
Fluorescence scanners, such as microarray readers – detect localized fluors in
samples in two dimensions
3.
Spectrofluorometers and microplate readers – record the average fluorescence in
samples
4.
Flow cytometers – analyze the fluorescence of individual cells in a sample population

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Fluorescent Labeling
Fluorescent labeling is the process of covalently attaching a fluorophore to another
molecule, such as a protein or nucleic acid. This is generally accomplished using a
reactive derivative of the fluorophore that selectively binds to a functional group present
in the target molecule. The most commonly labeled molecules are antibodies, which are
then used as specific probes for the detection of a particular target. Fluorescent labeling
can be applied to a wide variety of detection syste ms and allows sensitive and
quantitative measurement.
A chemically reactive derivative of a fluorophore is required for labeling molecules.
Common reactive groups include amine-reactive isoth iocyanate derivatives including
FITC, amine-reactive succinimidyl esters such as NHS-fluorescein or NHS-rhodamine,
and sulfhydryl-reactive maleimide-activated fluors such as fluorescein-5-maleimide.
Reaction of any of these reactive dyes with another molecule results in a stable covalent
bond formed between the fluorophore and the labeled molecule.
Isothiocyanates have long been used as a primary re active chemistry for attaching
fluorescent dyes to proteins through the primary amines of lysine side chains. NHS-
ester chemistry is now the preferred labelling method, because it has greater specificity
towards primary amines and produces a more stable l inkage following the labelling
procedure. Sulfhydryl-reactive chemistries occupy a smaller niche in protein labelling
where lysine residues must be preserved or where a cysteine residue is specifically
targeted to localize the fluorescent dye on the labelled protein.
Following a fluorescent labeling reaction, it is often necessary to remove any nonreacted
fluorophore from the labeled target molecule. This is usually accomplished by size
exclusion chromatography, taking advantage of the s ize difference between the
fluorophore and labeled protein, nucleic acid, etc.
Applications of radioisotopes in Food and Agriculture
Agricultural Tracers

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Tracers like those used in medicine are also used in agriculture to study plants and
their intake of fertilisers. The usage of tracers allows scientists and farmers to optimise
the use of fertilising and weed killing chemicals. Optimisation of these chemicals is
desirable because it saves money, and reduces chemical pollution.
When fertilisers are used in overly excessive amounts, the excess will run off and
pollute rivers nearby, as well as possibly seeping through to the water table
underground and polluting the water supply. To prevent this, studies are conducted to
find out the optimal amount of chemical required, with fertilisers and weed killers often
tagged by nitrogen-15 or phosphorus-32 radioisotope s. These radioisotopes are
analysed in the crops to see how much of the original chemical was actually consumed
by the plants, compared to how much was given. Using N-15 also enables assessment
of how much nitrogen is fixed from the air by soil and by root bacteria in legumes.
The ionising radiation from radioisotopes is also used to produce crops that are more
drought and disease resistant, as well as crops with increased yield or shorter growing
time. This practice has been in place for several decades, and has helped feed some
third-world countries. The collection of crops that have been modified with radiation
include wheat, sorghum, bananas and beans.
Insect control:
About 10% of the world's crops are destroyed by insects. In efforts to control insect
plagues, authorities often release sterile laboratory-raised insects into the wild. These
insects are made sterile using ionising radiation - they are irradiated with this radiation
before they hatch. Female insects that mate with sterile male insects do not reproduce,
and the population of the insect pests can be quickly curbed as a consequence. This
technique of releasing sterile insects into the wild, called the sterile insect technique
(SIT), is commonly used in protecting agricultural industries in many countries around
the world.
The technique is considered to be safer and better than conventional chemical
insecticides. Insects can develop resistance against these chemicals, and there are
health concerns about crops treated with them.

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The largest application of this technique so far was conducted in Mexico against
Mediterranean fruit-fly and screwworm in 1981.
In meats and other foods of animal origin, irradiation destroys the bacteria that cause
spoilage as well as diseases and illneses such as salmonella poisoning. This allows for a
safer food supply, and meats that can be stored for longer before spoilage. Additionally,
irradiation also inhibits tubers that cause fruits and vegetables to ripen. The result is
fresh fruits and vegetables that can be stored for longer before ripening.
Irradiation of food is carried out using accelerated electrons (beta radiation), and
ionising radiation from sources such as the radioisotopes cobalt-60 and cesium-137. X-
rays are also sometimes used.
Food irradiation is a well-tested process. Scientists have performed numerous decades
of research, and it has been shown that irradiation will not cause significant chemical
changes in foods that may affect human health, nor will it cause losses that may affect
the nutritional content of food. (Chemical residues left behind by irradiation are in
concentrations equivilant to about 3 drops in a swi mming pool. Chemical-based
preservatives and treatments usually leave more residues.) Taste is usually unaffected.
The World Health Organisation and food safety autho rities in many countries have
approved irradiation as a safe method of food treatment and preservation.
Irradiation poses less of a risk to human health than many chemical treatments that
are used today, which include the addition of chemi cal preservatives. The use of
radiation is sometimes favoured to using chemical preservatives, because no allergic
side-effect results. It is also better than heat-sterilisation because irradiation does not
destroy nutrients and vitamins, whereas heat treatment does.
Irradiation is inexpensive - typical costs are about 1-20 cents per kilogram of food
irradiated.
Food irradiation applications

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Low dose Inhibition of sprouting Potatoes, onions, garlic, ginger,
yam
Insect and parasite
disinfestation
Cereals, fresh fruit, dried foods
Delay ripening Fresh fruit, vegetables
Medium dose

Extend shelf life Fish, strawberries, mushrooms
Halt spoilage, kill pathogens Seafood, poultry, meat
High dose Industrial sterilisation Meat, poultry, seafood, prepared
foods
Decontamination Spices, etc

Quarantine and Exportation
Ionising radiation is used to rid goods of parasites and bugs before they are exported
out of a country. The radiation kills these parasites that may be quarantine hazards in
other countries. The technique is used in Australia to clear primary produce materials
such as raw wool and wood for export. It is also used worldwide in transporting archival
and historical documents. This is beneficial in that any microorganisms existing in the
paper that cause paper deterioration are destroyed.
Increasing Genetic Variability: Ionising radiation to induce mutations in plant
breeding has been used for several decades, and some 1800 crop varieties have been
developed in this way. Gamma or neutron irradiation is often used in conjunction with
other techniques, to produce new genetic lines of root and tuber crops, cereals and oil
seed crops.

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New kinds of sorghum, garlic, wheat, bananas, beans and peppers are more resistant to
pests and more adaptable to harsh climatic conditions. In Mali, irradiation of sorghum
and rice seeds has produced more productive and marketable varieties.
Radioisotopes in Industry
Modern industry uses radioisotopes in a variety of ways to improve productivity and, in
some cases, to gain information that cannot be obtained in any other way. Sealed
radioactive sources are used in industrial radiography, gauging applications and
mineral analysis. Short-lived radioactive material is used in flow tracing and mixing
measurements. Nuclear techniques are increasingly u sed in industry and
environmental management.
Neutron Techniques for Analysis
Neutrons can interact with atoms in a sample causin g the emission of gamma rays
which, when analysed for characteristic energies and intensity, will identify the types
and quantities of elements present. The two main te chniques are Thermal Neutron
Capture (TNC) and Neutron Inelastic Scattering (NIS). TNC occurs immediately after a
low-energy neutron is absorbed by a nucleus, NIS takes place instantly when a fast
neutron collides with a nucleus.
A particular application of NIS is where a probe containing a neutron source can be
lowered into a bore hole where the radiation is scattered by collisions with surrounding
soil. Since hydrogen (the major component of water) is by far the best scattering atom,
the number of neutrons returning to a detector in the probe is a function of the density
of the water in the soil. To measure soil density and water content, a portable device
with an americium-241-beryllium combination generat es gamma rays and neutrons
which pass through a sample of soil to a detector. (The neutrons arise from alpha
particles interacting with Be-9.) A more sophisticated application of this is in borehole
logging.

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Gamma & X-ray Techniques in Analysis
Gamma ray transmission or scattering can be used to determine the ash content of coal
on line on a conveyor belt. The gamma ray interactions are atomic number dependant,
and the ash is higher in atomic number than the coal combustible matter.
This technique is used to determine element concent rations in process streams of
mineral concentrators. Probes containing radioisotopes and a detector are immersed
directly into slurry streams. Signals from the prob e are processed to give the
concentration of the elements being monitored, and can give a measure of the slurry
density. Elements detected this way include iron, nickel, copper, zinc, tin and lead.
X-ray Diffraction (XRD) is a further technique for on-line analysis but does not use
radioisotopes.
Gamma Radiography: Gamma Radiography works in much the same way as x-r ays
screen luggage at airports. Instead of the bulky machine needed to produce x-rays, all
that is needed to produce effective gamma rays is a small pellet of radioactive material
in a sealed titanium capsule. The capsule is placed on one side of the object being
screened, and some photographic film is placed on the other side. The gamma rays, like
x-rays, pass through the object and create an image on the film. Just as x-rays show a
break in a bone, gamma rays show flaws in metal cas tings or welded joints. The
technique allows critical components to be inspecte d for internal defects without
damage.
Gamma sources are normally more portable than x-ray equipment so have a clear
advantage in certain applications, such as in remote areas.
Gauging
The radiation that comes from a radioisotope has it s intensity reduced by matter
between the radioactive source and a detector. Detectors are used to measure this
reduction. This principle can be used to gauge the presence or the absence, or even to
measure the quantity or density, of material between the source and the detector. The

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advantage in using this form of gauging or measurement is that there is no contact with
the material being gauged.
Many process industries utilise fixed gauges to mon itor and control the flow of
materials in pipes, distillation columns, etc, usually with gamma rays. The height of the
coal in a hopper can be determined by placing high energy gamma sources at various
heights along one side with focusing collimators directing beams across the load.
Detectors placed opposite the sources register the breaking of the beam and hence the
level of coal in the hopper. Such level gauges are among the most common industrial
uses of radioisotopes.
Some machines which manufacture plastic film use ra dioisotope gauging with beta
particles to measure the thickness of the plastic film. The film runs at high speed
between a radioactive source and a detector. The detector signal strength is used to
control the plastic film thickness.
In paper manufacturing, beta gauges are used to monitor the thickness of the paper at
speeds of up to 400 m/s.
Gamma Sterilisation: Gamma irradiation is widely used for sterilising me dical
products, for other products such as wool, and for food. Cobalt-60 is the main isotope
used, since it is an energetic gamma emitter. It is produced in nuclear reactors,
sometimes as a by-product of power generation. Smaller gamma irradiators are used for
treating blood for transfusions and for other medical applications.
Scientific Uses: Using tracing techniques, research is conducted wit h various
radioisotopes which occur broadly in the environment, to examine the impact of human
activities. The age of water obtained from underground bores can be estimated from the
level of naturally occurring radioisotopes in the water. This information can indicate if
groundwater is being used faster than the rate of r eplenishment. Trace levels of
radioactive fallout from nuclear weapons testing in the 1950s and 60s is now being
used to measure soil movement and degradation. This is assuming greater importance
in environmental studies of the impact of agriculture.

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Tracing/Mixing Uses
Even very small quantities of radioactive material can be detected easily. This property
can be used to trace the progress of some radioactive material through a complex path,
or through events which greatly dilute the original material. In all these tracing
investigations, the half-life of the tracer radioisotope is chosen to be just long enough to
obtain the information required. No long-term residual radioactivity remains after the
process.
Sewage from ocean outfalls can be traced in order to study its dispersion. Small leaks
can be detected in complex systems such as power station heat exchangers. Flow rates
of liquids and gasses in pipelines can be measured accurately, as can the flow rates of
large rivers.
Mixing efficiency of industrial blenders can be measured and the internal flow of
materials in a blast furnace examined. The extent of termite infestation in a structure
can be found by feeding the insects radioactive wood substitute, then measuring the
extent of the radioactivity spread by the insects. This measurement can be made
without damaging any structure as the radiation is easily detected through building
materials.
Industrial Radioisotopes
Naturally-occurring radioisotopes:
Carbon-14: Used to measure the age of water (up to 50,000 years)
Chlorine-36: Used to measure sources of chloride and the age of water (up to 2 million
years)
Lead-210: Used to date layers of sand and soil up to 80 years
Tritium (H-3): Used to measure 'young' groundwater (up to 30 years)
Artificially-produced radioisotopes:

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Americium-241: Used in backscatter gauges, smoke detectors, fill height detectors and
in measuring ash content of coal.
Chromium 57: Used to label sand to study coastal erosion.
Gold-198 & Technetium-99m: Used to study sewage and liquid waste movements, as
well as tracing factory waste causing ocean pollution, and to trace sand movement in
river beds and ocean floors.
Gold-198: Used to label sand to study coastal erosion.
Hydrogen-3 (Tritiated Water): Used as a tracer to study sewage and liquid wastes
Iridium-192: Used in gamma radiography to locate flaws in metal components.
IN VIVO LABELING
Metabolic labeling refers to methods in which chemical detection- or affinity-tags can be
added to biomolecules in vivo using the endogenous synthesis and modification
machinery of living cells. When analogs of molecular building blocks (e.g., amino acids)
can be designed to contain specifically targetable tags that do not interfere with the
metabolic machinery of the cell, they provide a mechanism for metabolic labeling. The
strategy makes possible a number of powerful experi mental approaches for the
investigation of cellular pathways.
Methods of invivo labelling:
1.
Metabolic Labeling and Staudinger Ligation
The Staudinger ligation (azide-phosphine) chemistry is one of several crosslinking
techniques that are amenable to in vivo metabolic labeling applications. Because the
azide component of the chemoselective reaction pair is so small, it can be supplied to
living cells in the form of bioorthogonal molecules that substitute for the building blocks
cells use to synthesize proteins or other macromolecules.

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Overview of Staudinger ligation (azide-phosphine co njugation) as a strategy for
metabolic labeling.
Left.Phosphine activation of proteins is easily accomplished with reactive, phosphine-
containing chemical modification reagents; alternatively, fluorescent dyes and affinity
tags such as biotin are available ready-made in phosphine-activated form.
Center. The tiny azide tag can be added to biomolecules by in vivo incorporation of
azide-containing derivatives of metabolic building blocks (amino acids, sugars, etc.);
alternatively, proteins or other molecules can be modified in vitro with reactive azide-
containing reagents.
Right. When combined, phosphine-activated compounds conj ugate with high specificity
to azide-tagged molecules, resulting in stable covalent attachment of "A" and "B"
molecules.
2.
Bioorthogonal Labeling Compounds
In practical terms, whether or not a particular labeling chemistry can be used in
metabolic labeling depends upon its chemoselectivity (reaction specificity) and metabolic
compatibility (i.e., production of bioorthogonal derivatives through metabolism).
In this chemoselective ligation strategy, one component of the reaction pair is supplied
as a substitute (an analog) of a naturally occurring molecule that is required for
catabolism of the target macromolecules. This is the meaning of "bioorthogonal" – that
the biological function of the molecule is unaffected by the reactive group it contains. In
other words, the reactive group is "invisible" to the biological system.

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The azide group in the Staudinger reaction pair has this bioorthogonal property. When
supplied to cells, synthetic azide-containing analogs of amino acids or sugars can be
incorporated during protein synthesis or post-translational glycosylation using cellular
metabolic or regulatory machinery. Thus, the relevant chemoselective reactive group is
added by in vivo by metabolic labeling. Alternatively, bioorthogonal derivatives can be
incorporated into specific non-protein targets using in vitro enzymatic reactions. Once
target molecules are labeled (tagged) with the bioorthogonal group (azide), they can be
chemoselectively conjugated or tagged by reaction with the desired phosphine-activated
reagent (biotin, fluor, etc.) using the Staudinger reaction. In this way, chemoselective
ligation using bioorthogonal derivatives blends the simplicity of metabolically encoded
tags with specific labeling and the versatility of small-molecule probes.



When used in combination with specially formulated limiting media that is devoid of
leucine and methionine, the photo-activatable derivatives are treated like naturally
occurring amino acids by the protein synthesis machinery within the cell. As a result,
they can be substituted for leucine or methionine in the primary sequence of proteins
during catabolism and growth. Photo-Leucine and Photo-Methionine derivatives contain
diazirine rings that activate when exposed to UV light to become reactive intermediates
that form covalent bonds with nearby protein side chains and backbones. Naturally
associating binding partners within the cell can be instantly trapped by photoactivation

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of the diazirine-containing proteins in the cultured cells. Crosslinked protein complexes
can be detected by decreased mobility on SDS-PAGE followed by Western blot detection,
size exclusion chromatography, sucrose density grad ient sedimentation or mass
spectrometry.
3.
Glycosylation Labeling Reagents
Metabolic labeling with bioorthogonal monosaccharides (sugars) that are used by cells
to glycosylate proteins and other cell constituents provides for a variety of experimental
approaches. The effects of drugs or other treatment conditions on total or sugar-specific
glycosylation can be measured. When combination with protein-specific purification or
antibody-detection, the glycosylation patterns for specific glycoproteins can be
investigated.
We offer three azido sugar reagents for metabolic labeling with the Staudinger ligation
(azide-phosphine) chemistry. As illustrated in the figure above, the azide tag of
incorporated sugars can be chemoselectively targete d with phosphine-activated
compounds for a variety of purposes (e.g., detection or affinity purification).

AUTORADIOGRAPHY

Radiography is the visualisation of the pattern of distribution of radiation. In general,
the radiation consists of X-rays, gamma or beta rays, and the recording medium is a
photographic film. For classical X-rays, the specimen to be examined is placed between
the source of radiation and the film, and the absorption and scattering of radiation by
the specimen produces its image on the film. In con trast, in autoradiography the
specimen itself is the source of the radiation, which originates from radioactive material

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incorporated into it. The recording medium which makes visible the resultant image is
usually, though not always, photographic emulsion.
Autoradiography is the use of X-ray (or occasionally photographic) film to detect
radioactive materials. It produces a permanent record of the positions and relative
intensities of radiolabeled bands in a gel or blot. Typically, biomolecules are labeled
with 32P or 35S, and detected by overnight film exposure.
History
The first autoradiography was obtained accidently around 1867 when a blackening was
produced on emulsions of silver chloride and iodide by uranium salts. Such studies and
the work of the Curies in 1898 demonstrated autoradiography before, and contributed
directly to, the discovery of radioactivity.
The development of autoradiography as a biological technique really started to happen
after World War II with the development of photographic emulsions and then stripping
film made of silver halide.
Radioactive Exposure of Film
Beta particles emitted by radionuclides penetrate film emulsions to a depth proportional
to their energy. As these particles pass through the film, they activate the silver halide
crystals in the emulsion. Activated crystals are detected by reduction to black silver
grains in the development step. The activated silver halide crystals are not stable,
although they can be stabilized by further exposure to b-particles, or to light. On
average, a crystal will require 5 "hits" from either radioactive emissions or light to be
stably activated, and thus detected.
Autoradiography is the exposure of x-ray film (or a photographic emulsion) to a
radioactive sample. The radioactivity behaves as light and results in the exposure of the
photographic emulsion on the film. Radiation activa tes the silver halide in the
photographic emulsion. During development the activated silver halide is converted to
metallic silver. The metallic silver will appear as 'grains' on the film in positions where
radioactivity is located. In other words the exposed film will be darkening in proportion
to the amount of radioactivity it was exposed to.

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Autoradiography Method
• Living cells are briefly exposed to a ‘pulse’ of a specific radioactive compound.
• The tissue is left for a variable time.
• Samples are taken, fixed, and processed for light or electron microscopy.
• Sections are cut and overlaid with a thin film of photographic emulsion.
• Left in the dark for days or weeks (while the radioisotope decays). This exposure time
depends on the activity of the isotope, the temperature and the background radiation
(this will produce with time a contaminating increase in ‘background’ silver grains in
the film).
• The photographic emulsion is developed (as for conventional photography).
• Counterstaining e.g. with toluidine blue, shows the histological details of the tissue. The
staining must be able to penetrate, but not have an adverse affect on the emulsion.
• Alternatively, pre-staining of the entire block of tissue can be done (e.g. with Osmium
on plastic sections coated with stripping film [or dipping emulsion] as in papers by
McGeachie and Grounds) before exposure to the photographic emulsion. This avoids the
need for individually (post-) staining each slide.
• It is not necessary to coverslip these slides
• The position of the silver grains in the sample is observed by light or electron
microscopy
• These autoradiographs provide a permanent record.
Types of photograhic detection systems
Stripping film consists of an even layer of photographic emulsion on a supporting
gelatin membrane (e.g. Kodak AR10), it is floated on water and then wrapped around
the slide and forms very close contact as it dries. This was once widely used but is now
no longer made. It has the major advantage of uniform thickness but the disadvantage
that the supporting membrane prevents counterstaining of the section and therefore the
tissue block must be pre-stained before sections are coated.
Liquid photographic emulsion . This is the method routinely used today. It is simpler
and much quicker to do, but the layer of liquid emu lsion (e.g. Kodak NB2) can

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be slightly uneven in thickness as it flows down to the bottom of the slide as it is
withdrawn: for most purposes this slight variation is not important, unless the number
of grains are being strictly counted and compared across one slide.
The two most common uses of autoradiography is in c onjunction with gel
electrophoresis or in conjunction with microscopy.
Autoradiography of gels allows the identification and quantitation of specific proteins or
nucleic acids. It is also possible to localize radioactive markers to particular cells in
tissue sections or subcellular structures within cells by autoradiography.
Pulsed field gel electrophoresis:
PFGE allows investigators to separate much larger pieces of DNA than conventional
agarose gel electrophoresis. In conventional gels, the current is applied in a single
direction (from top to bottom). But in PFGE, the direction of the current is altered at a
regular interval as shown in the animated gif below.
In 1984, Schwartz and Cantor described pulsed field gel electrophoresis (PFGE),
introducing a new way to separate DNA. In particular, PFGE resolved extremely large
DNA for the first time, raising the upper size limit of DNA separation in agarose from
30-50 kb to well over 10 Mb (10,000 kb).
Applications
These include cloning large plant DNA using yeast artificial chromosomes (YAC's).
Identifying restriction fragment length polymorphisms (RFLP's) and construction of
physical maps;
Detecting in vivo chromosome breakage and degradation (Elia, et al., 1991);
Determining the number and size of chromosomes ("el ectrophoretic karyotype") from
yeasts, fungi, and parasites such as Leishmania, Plasmodium, and Trypanosoma.
Theory

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Although the theory of pulsed field electrophoresis is a matter of debate, qualitative
statements can be made about the movement of DNA in agarose gels during PFGE.
During continuous field electrophoresis, DNA above 30-50 kb migrates with the same
mobility regardless of size. This is seen in a gel as a single large diffuse band. If,
however, the DNA is forced to change direction during electrophoresis, different sized
fragments within this diffuse band begin to separate from each other.
With each reorientation of the electric field relative to the gel, smaller sized DNA will
begin moving in the new direction more quickly than the larger DNA. Thus, the larger
DNA lags behind, providing a separation from the smaller DNA.
Field Strength
The field strength has a profound effect on pulsed field separations and is a
compromise between separation time and resolution of a particular size class. Four to
six volts/cm is generally required for resolving DN A up to 2000 kb (e.g., S.
cerevisiae chromosomes) in a reasonable period of time (e.g., 1-2 days). However, these
field strengths trap and immobilize even bigger DNA in the agarose matrix, and DNA >
3000 kb requires 2 V/cm or less for separation.
Pulse Time
Pulse time primarily changes the size range of separation. Longer pulse times lead to
separation of larger DNA. For example, at 5.4 V/cm, the 1.6 Mb and 2.2 Mb
chromosomes from S. cerevisiae separate as a single band with 90-second pulse length.
Increasing the pulse length to 120 seconds resolves these into two bands (Gemmill,
1991).
Reorientation Angle
Any angle between 96 and 165 produces roughly equivalent separation. The smaller the
angle, however, the faster the DNA mobility. And for separating extremely large DNA, 96
to 105 is almost a requirement to get a good separation in the shortest possible time.
Buffers

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Two buffers are commonly employed for PFGE--TAE and TBE (1x TAE is 40 mM Tris
acetate, 1 mM EDTA, pH 8.0; 1x TBE is 89 mM Tris, 89 mM boric acid, 2 mM EDTA, pH
8.0). Both are used at a relatively low ionic strength to prevent heating and carry the
designations of either 0.25 and 0.5x to indicate the dilution relative to the standard
concentration.
Agarose
The type of agarose also affects DNA separation, with the fastest mobilities and best
resolution achieved in gels made of low electroendosmosis (EEO) agarose. Although
most standard electrophoresis grades of agarose are suitable for PFGE agarose with
minimal EEO will provide a faster separation.

The gray box is the gel; the six sets of 4 black lines represent the 3 pairs of electrodes.
Initially, the gel is empty but soon Whole chromosomes mixed with blue loading dye will
be placed into the wells. Then the current will be turned on and the direction of the
current will change in a regular pattern. This is repeated until the loading dye reaches
near the end of the gel then the gel is soaked in a solution containing ethidium bromide
which fluoresces orange when bound to DNA.
AUTOFLUOROGRAPHY
The localization and recording of a radiolabel within a solid specimen is known as
autoradiography and involves the production of an image in a photographic emulsion.
Such emulsions consist of silver halide crystals suspended in a clear phase composed
mainly of gelatin. When a β-particle or γ-ray from a radionuclide passes through the
emulsion, the silver ions are converted to silver atoms.

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This results in a latent image being produced, which is converted to a visible image
when the image is developed. Development is a system of amplification in which the
silver atoms cause the entire silver halide crystal to be reduced to metallic silver.
Unexposed crystals are removed by dissolution in fixer, giving an autoradiographic
image which represents the distribution of radiolabel in the original sample.
In direct autoradiography, the sample is placed in intimate contact with the film and
the radioactive emissions produce black areas on the developed autoradiograph. It is
best suited to detection of weak- to medium-strength β-emitting radionuclides (
3
H,
14
C,
35
S).
Direct autoradiography is not suited to the detection of highly energetic β-particles,
such as those from
32
P, or for γ-rays emitted from isotopes like
125
I. These emissions
pass through and beyond the film, with the majority of the energy being wasted. Both
32P and
125I are best detected by indirect autoradiography.
Indirect autoradiography describes the technique by which emitted energy is converted
to light by means of a scintillator, using fluorography or intensifying screens. In
fluorography the sample is impregnated with a liqui d scintillator. The radioactive
emissions transfer their energy to the scintillator molecules, which then emit photons
which expose the photographic emulsion.
Fluorography is mostly used to improve the detection of weak β-emitters (Fig.).

Autoradiographs showing the detection of
35
S- and
3
H-labelled proteins in
acrylamide gels with (+) and without (−) fluorography.

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Intensifying screens are sheets of a solid inorganic scintillator which are placed behind
the film. Any emissions passing through the photographic emulsion are absorbed by the
screen and converted to light, effectively superimposing a photographic image upon the
direct autoradiographic image.
The gain in sensitivity which is achieved by use of indirect autoradiography is offset by
non-linearity of film response. A single hit by a β-particle or γ-ray can produce
hundreds of silver atoms, but a single hit by a photon of light produces only a single
silver atom. Although two or more silver atoms in a silver halide crystal are stable, a
single silver atom is unstable and reverts to a silver ion very rapidly. This means that
the probability of a second photon being captured b efore the first silver atom has
reverted is greater for large amounts of radioactivity than for small amounts. Hence
small amounts of radioactivity are under-represented with the use of fluorography and
intensifying screens. This problem can be overcome by a combination of pre-exposing a
film to an instantaneous flash of light (pre-flashing) and exposing the autoradiograph at
-70°C.
Pre-flashing provides many of the silver halide crystals of the film with a stable pair of
silver atoms. Lowering the temperature to -70°C increases the stability of a single silver
atom, increasing the time available to capture a second photon (Fig.).

The improvement in sensitivity of detection of
125
I-labelled IgG by
autoradiography obtained by using an intensifying s creen and pre-flashed film. A,
no screen and no pre-flashing; B, screen present but film not pre-flashed;

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C, use of screen and pre-flashed film.
DNA SEQUENCING BY ENZYMATIC AND CHEMICAL METHODS
The term DNA sequencing refers to sequencing methods for determining the order of the
nucleotide bases—adenine, guanine, cytosine, and thymine—in a molecule of DNA.
Two DNA sequencing methods were developed independe ntly in the 1970s by Maxam
and Gilbert and bySanger.
Sanger Method
Frederick Sanger described this method of DNA seque ncing which relies on DNA
polymerase’s use 2',3'-dideoxynucleoside triphosphates [ddNTPs] as substrates" . At the
3' end of ddNTPs, there is a hydrogen instead of the hydroxyl group necessary for DNA
chain elongation. This altered structure causes termination of the growing DNA chain
and is the basis behind Sanger’s method, which is also called the dideoxy method.

The DNA sequencing reaction begins with a single stranded DNA molecule for which an
oligonucleotide primer has been designed. The primer anneals to the corresponding
sequence on the single stranded DNA and DNA polymer ase directs the synthesis of a
complementary copy of the template from the 3' end of the primer. Several different
DNA polymerases are appropriate to use for this tec hnique, including T7 DNA
polymerase, Bst DNA polymerase I, Pfu exo
-
, and Taq.

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The reaction is carried out in four different tubes, one for each nucleotide. In each tube
the template DNA, DNA polymerase, and all four nucl eotides are added. The end
products must be labelled in order to be detected, and this is done by attaching
radioactive labels to one of the dNTPs used in the reaction. The DNA sample is divided
into four separate sequencing reactions, containing all four of the standard
deoxynucleotides (dATP, dGTP, dCTP and dTTP) and th e DNA polymerase. To each
reaction is added only one of the four dideoxynucleotides (ddATP, ddGTP, ddCTP, or
ddTTP) which are the chain-terminating nucleotides, lacking a 3'-OH group required for
the formation of a phosphodiester bond between two nucleotides, thus terminating DNA
strand extension and resulting in DNA fragments of varying length.
Several radioactive labels can be used, including ["-
32
P], ["-
33
P], and ["-
35
S]. Usually, ["-
33
P] is used because it gives better resolution than ["-
35
S] and is safer to use than ["-
32
P].
A relatively small amount of one type of ddNTP is also added to the appropriate tube (eg.
ddATP to dATP tube, ddGTP to dGTP tube, etc.).
During the reaction in, for example, dATP, each time the DNA polymerase reaches a T
position on the template, there is a small chance that ddATP will be incorporated,
terminating the chain elongation. In most cases, the DNA polymerase will add dATP and
the DNA synthesis will continue until another T position is reached, where there will be
another small chance for chain termination to take place. This occurs in each tube until
the end of the reaction. Every possible fragment length will be synthesized, with the
shortest chain being the length of the primer plus one nucleotide.
The reaction products are separated, based on size, using a thin polyacrylamide gel,
which usually contains 7M urea. The urea "acts as a denaturant to reduce the effects of
DNA secondary structure". The high temperature that the gel is run at also contributes
to the denaturation of the DNA.
Four lanes are used, one for each tube used in the reaction. Once the gel has been run,
it is exposed to X-ray film, resulting in an autoradiogram which can be read from the
bottom up, giving the 5' to 3' sequence of the DNA that was generated in the reaction.
Up to 400 or 500 bases can be sequenced using this manual method.

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Maxam & Gilbert's method (chemical cleavage)
In 1976–1977, Allan Maxam and Walter Gilbert develo ped a DNA sequencing method
based on chemical modification of DNA and subsequent cleavage at specific bases. Also
sometimes known as "chemical sequencing", this meth od originated in the study of

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DNA-protein interactions (footprinting), nucleic ac id structure and epigenetic
modifications to DNA, and within these it still has important applications.
The DNA fragment to be sequenced is end labelled by the addition of P-dATP either at
the 5 ends (Polynucleotide kinase) or at the 3 ends (deoxynucleotidyl transferase). The
end labelled fragment is now digested with RE which cleaves it into two fragments of
unequal length. As a result only one end of each of the two fragments thus produced
will be labelled. The 2 fragments are separated through gel electrophoresis and they are
sequenced separately.
Alternatively, the end labelled fragment is denatured and its 2 complementary strands
are separated through gel electrophoresis. Each strand will be labelled at one end (5’ or
3’) only.
The single end labelled double or single stranded DNA samples thus produced are
subjected to base specific chemical cleavage so that in a reaction mixture cleavage
occurs only at one of the following four sites; G, C, G+A, or C+T. Each DNA sample is
partially digested in 4 separate reaction mixtures ( one each for specific cleavage at the
sites of G, C, G+A, or C+T). In these reaction mixtures each DNA fragment/strand is
expected to be cleaved, on an average only once at any one of the sites having the
particular base for which the reaction mixture is specific.
The base specific cleavage of DNA fragments involves the following steps: modification of
the concerned base (2) removal of the modified base from the DNA strand, and (3)
induction of strand break (break in the sugar phosphate backbone) in the position from
which the modified base has been removed.
Four identical samples of the prepared fragment are subjected to four different sets of
chemical reactions that selectively cut the DNA backbone at G, G + A, C + T, or C
residues. The reactions are controlled so that each labeled chain is likely to be broken
only once. The labeled subfragments created by all four reactions have the label at one
end and the chemical cleavage point at the other.
Gel electrophoresis and autoradiography of each separate mixture yield one radioactive
band for each nucleotide in the original fragment, each separated according to their

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length. Bands appearing in the G and C lanes can be read directly. Bands in the A + G
lane that are not duplicated in the G lane are read as A. Bands in the T + C lane that
are not duplicated in the C lane are read as T. The sequence is read from the bottom of
the gel up.

AGAROSE GEL ELECTROPHORESIS
The progress of the first experiments on cutting and joining of DNA molecules was
monitored by velocity sedimentation in sucrose grad ients. However, this has been
entirely superseded by gel electrophoresis.
Gel electrophoresis is not only used as an analytical method, it is routinely used
preparatively for the purification of specific DNA fragments. The gel is composed of
polyacrylamide or agarose. Agarose is convenient for separating DNA fragments ranging
in size from a few hundred base pairs to about 20 kb (Fig.).

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Electrophoresis of DNA in agarose gels. The direction of migration is indicated by
the arrow. DNA bands have been visualized by soakin g the gel in a solution of
ethidium bromide, which complexes with DNA by inter calating between stacked
base-pairs, and photographing the orange fluorescen ce which results upon
ultraviolet irradiation.
Polyacrylamide is preferred for smaller DNA fragments. The mechanism responsible for
the separation of DNA molecules by molecular weight during gel electrophoresis is not
well understood. The migration of the DNA molecules through the pores of the matrix
must play an important role in molecular-weight separations since the electrophoretic
mobility of DNA in free solution is independent of molecular weight. An agarose gel is a
complex network of polymeric molecules whose average pore size depends on the buffer
composition and the type and concentration of agarose used. DNA movement through
the gel was originally thought to resemble the motion of a snake (reptation). However,
real-time fluorescence microscopy of stained molecules undergoing electrophoresis has
revealed more subtle dynamics.
DNA molecules display elastic behaviour by stretching in the direction of the applied
field and then contracting into dense balls. The larger the pore sizes of the gel, the

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greater the ball of DNA which can pass through and hence the larger the molecules
which can be separated. Once the globular volume of the DNA molecule exceeds the
pore size, the DNA molecule can only pass through b y reptation. This occurs with
molecules about 20 kb in size and it is difficult to separate molecules larger than this
without recourse to pulsed electrical fields.
PAGE
Polyacrylamide gel electrophoresis (PAGE) has replaced starch gel electrophoresis for
the separation of proteins, small RNA fragments and very small DNA fragments.
Polyacrylamide is more versatile than starch, because the molecular sieving effect can
be controlled to a much greater extent, and because the adsorption of proteins to the
gel is negligible.
Polyacrylamide gels are prepared by the reaction of acrylamide (monomer) with N,N0-
methylenebis(acrylamide) (cross-linker) in the presence of a catalyst and initiator, as
shown in Eq.

Initiators include ammonium persulfate and potassium persulfate, where the S
2O2
_8
dianion decomposes into two SO
__4
radicals, while the commonly used catalyst is
tetramethylethylenediamine [TEMED, (CH
3)2N(CH2)2N(CH3)2], which reacts with the
sulfate radical anion to produce a longer lived radical species.
The concentration of TEMED used during gel casting determines the length of the
polyacrylamide chains formed, and therefore the mechanical stability of the gel.
Lower monomer concentrations require higher TEMED c oncentrations, but excess
catalyst should be minimized, because it may interact with proteins or alter the Ph of
the running buffer. Because the polymerization that occurs during the casting of a
polyacrylamide gel is a random process, a distribution of pore sizes will occur. It has
been shown that the average pore size depends on the total amount of acrylamide used
and the degree of crosslinking. The total monomer concentration is generally between 5
and 20% by weight.

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Experimentally, PAGE may be employed in a column, for preparative separations, or in
a slab, where several samples may be separated under identical conditions.
These configurations are shown in Figure.

The preparation of a PAGE column begins with a tube of ~5–10 mm inner diameter, 10
cm or more in length. During gel polymerization, the gel mixture is gently overlayered
with water to give a flat upper surface. Following casting, the water is removed, and the
sample is applied. Samples are suspended in a conce ntrated sucrose solution (high
density) and layered over the gel, to prevent mixing with the buffer in the upper (usually
cathodic) electrode chamber. The run is begun at low current (1 mA for ~30 mins) to
allow the entire sample to enter the gel.
The current is then increased to 2–5 mA during the run. A tracking dye such as
bromophenol blue (anionic) or methylene blue (cationic) is added to the sample to
monitor the progress of the separation. Buffers are often cooled and stirred to minimize
i2R heating and local pH changes that may occur near the gel–buffer interface. When
the tracking dye has migrated to the bottom of the column, the gel is removed using
pressure, and proteins are fixed in place by precipitation [this occurs during a soaking
step in a solution of trichloroacetic acid (TCA)] and are then stained for detection.
Slab PAGE allows simultaneous electrophoresis of a number of samples to be performed
under identical conditions. Slab PAGE has a higher resolving power than column PAGE,
and is often used for molecular weight and purity determinations.
Because the slab is relatively thin, heat dissipation is more efficient in a slab than in a
column. The slab can be used vertically or horizontally, but in practice, the horizontal
slab method is used only when the monomer plus cros s-linker concentrations are low

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and the gel is soft. A slab gel is prepared in the same way as a column gel, except that a
comb, or slot-former, is inserted before polymerization.
Removal of the comb after the gel has set leaves sample wells that are separated from
each other by continuous strips of gel.
PFGE
In pulsed-field gel electrophoresis (PFGE) molecules as large as 10 Mb can be separated
in agarose gels. This is achieved by causing the DNA to periodically alter its direction of
migration by regular changes in the orientation of the electric field with respect to the
gel. With each change in the electric-field orientation, the DNA must realign its axis
prior to migrating in the new direction. Electric-field parameters, such as the direction,
intensity and duration of the electric field, are set independently for each of the different
fields and are chosen so that the net migration of the DNA is down the gel.

Schematic representation of CHEF (contour-clamped h omogeneous electrical
field) pulsed-field gel electrophoresis.
The difference between the directions of migration induced by each of the electric fields
is the reorientation angle and corresponds to the angle that the DNA must turn as it
changes its direction of migration each time the fields are switched.
A major disadvantage of PFGE, as originally described, is that the samples do not run
in straight lines. This makes subsequent analysis difficult. This problem has been
overcome by the development of improved methods for alternating the electrical field.
The most popular of these is contour-clamped homoge neous electrical-field
electrophoresis (CHEF).
In early CHEF-type systems (Fig.) the reorientation angle was fixed at 120°.
However, in newer systems, the reorientation angle can be varied and it has been found
that for whole-yeast chromosomes the migration rate is much faster with an angle of
106°. Fragments of DNA as large as 200–300 kb are r outinely handled in genomics

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work and these can be separated in a matter of hour s using CHEF systems with a
reorientation angle of 90° or less. Aaij and Borst (1972) showed that the migration rates
of the DNA molecules were inversely proportional to the logarithms of the molecular
weights. Subsequently, Southern (1979a,b) showed th at plotting fragment length or
molecular weight against the reciprocal of mobility gives a straight line over a wider
range than the semilogarithmic plot. In any event, gel electrophoresis is frequently
performed with marker DNA fragments of known size, which allow accurate size
determination of an unknown DNA molecule by interpolation. A particular advantage of
gel electrophoresis is that the DNA bands can be readily detected at high sensitivity.

The bands of DNA in the gel are stained with the intercalating dye ethidium bromide
(Fig.), and as little as 0.05 µg of DNA in one band can be detected as visible
fluorescence when the gel is illuminated with ultraviolet light.
In addition to resolving DNA fragments of different lengths, gel electrophoresis can be
used to separate different molecular configurations of a DNA molecule. Gel
electrophoresis can also be used for investigating protein–nucleic acid interactions in
the so-called gel retardation or band shift assay. It is based on the observation that
binding of a protein to DNA fragments usually leads to a reduction in electrophoretic
mobility. The assay typically involves the addition of protein to linear double-stranded
DNA fragments, separation of complex and naked DNA by gel electrophoresis and
visualization. A review of the physical basis of electrophoretic mobility shifts and their
application is provided by Lane et al. (1992).
METHODS OF NUCLEIC ACID HYBRIDIZATION
Nucleic acid labelling and hybridization on membranes have formed the basis for a
range of experimental techniques central to recent advances in our understanding of
the organization and expression of the genetic material. These techniques may be
applied in the isolation and quantification of specific nucleic acid sequences and in the

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study of their organization, intracellular localization, expression and regulation. A
variety of specific applications includes the diagnosis of infectious and inherited
disease.

An overview of the steps involved in nucleic acid blotting and membrane hybridization
procedures is shown in Fig.
Blotting describes the immobilization of sample nucleic acids on to a solid support,
generally nylon or nitrocellulose membranes. The blotted nucleic acids are then used as
‘targets’ in subsequent hybridization experiments. The main blotting procedures are:
• Blotting of nucleic acids from gels;
• Dot and slot blotting;
• Colony and plaque blotting.
Hybridization of nucleic acids on membranes
The hybridization of nucleic acids on membranes is a widely used technique in gene
manipulation and analysis. Unlike solution hybridizations, membrane hybridizations
tend not to proceed to completion. One reason for this is that some of the bound nucleic
acid is embedded in the membrane and is inaccessibl e to the probe. Prolonged
incubations may not generate any significant increase in detection sensitivity.
The composition of the hybridization buffer can greatly affect the speed of the reaction
and the sensitivity of detection. The key components of these buffers are shown below:
Rate enhancers: Dextran sulphate and other polymers act as volume e xcluders to
increase both the rate and the extent of hybridization

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Detergents and blocking agents: Dried milk, heparin and detergents such as sodium
dodecyl sulphate (SDS) have been used to depress non-specific binding of the probe to
the membrane. Denhardt’s solution (Denhardt 1966) u ses Ficoll, polyvinylpyrrolidone
and bovine serum albumin.
Denaturants: Urea or formamide can be used to depress the melting temperature of the
hybrid so that reduced temperatures of hybridization can be used
Heterologous DNA: This can reduce non-specific binding of probes to non-homologous
DNA on the blot
Stringency control
Stringency can be regarded as the specificity with which a particular target sequence is
detected by hybridization to a probe. Thus, at high stringency, only completely
complementary sequences will be bound, whereas low- stringency conditions will allow
hybridization to partially matched sequences. Stringency is most commonly controlled
by the temperature and salt concentration in the posthybridization washes, although
these parameters can also be utilized in the hybridization step.
In practice, the stringency washes are performed under successively more stringent
conditions (lower salt or higher temperature) until the desired result are obtained.
The melting temperature (Tm) of a probe–target hybrid can be calculated to provide a
starting-point for the determination of correct stringency. The Tm is the temperature at
which the probe and target are 50% dissociated. For probes longer than
100 base pairs:
Tm = 81.5°C + 16.6 log M + 0.41 (% G + C)
where M = ionic strength of buffer in moles/litre.With long probes, the hybridization is
usually carried out at Tm − 25°C. When the probe is used to detect partially matched
sequences, the hybridization temperature is reduced by 1°C for every 1% sequence
divergence between probe and target.
Oligonucleotides can give a more rapid hybridization rate than long probes as they can
be used at a higher molarity. Also, in situations where target is in excess to the probe,
for example dot blots, the hybridization rate is diffusion-limited and longer probes
diffuse more slowly than oligonucleotides. It is standard practice to use oligonucleotides
to analyse putative mutants following a site-directed mutagenesis experiment where the
difference between parental and mutant progeny is often only a single base-pair change.

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The availability of the exact sequence of oligonucl eotides allows conditions for
hybridization and stringency washing to be tightly controlled so that the probe will only
remain hybridized when it is 100% homologous to the target. Stringency is commonly
controlled by adjusting the temperature of the wash buffer. The ‘Wallace rule’ is used to
determine the appropriate stringency wash temperature:
Tm = 4 × (number of GC base pairs) + 2 × (number of AT base pairs)
In filter hybridizations with oligonucleotide probes, the hybridization step is usually
performed at 5°C below Tm for perfectly matched sequences. For every mismat ched
base pair, a further 5°C reduction is necessary to maintain hybrid stability.
The design of oligonucleotides for hybridization experiments is critical to maximize
hybridization specificity.
Consideration should be given to:
• Probe length – the longer the oligonucleotide, the less chance there is of it binding to
sequences other than the desired target sequence under conditions of high stringency;
• Oligonucleotide composition – the GC content will influence the stability of the resultant
hybrid and hence the determination of the appropriate stringency washing conditions.
Also the presence of any non-complementary bases wi ll have an effect on the
hybridization conditions.
SOUTHERN BLOTTING TECHNIQUES
The original method of blotting was developed by Southern (1975, 1979b) for detecting
fragments in an agarose gel that are complementary to a given RNA or DNA sequence.
In this procedure, referred to as Southern blotting, the agarose gel is mounted on a
filter-paper wick which dips into a reservoir containing transfer buffer (Fig.).
The hybridization membrane is sandwiched between the gel and a stack of paper towels
(or other absorbent material), which serves to draw the transfer buffer through the gel
by capillary action. The DNA molecules are carried out of the gel by the buffer flow and
immobilized on the membrane. Initially, the membran e material used was
nitrocellulose.
The main drawback with this membrane is its fragile nature. Supported nylon
membranes have since been developed which have grea ter binding capacity for nucleic
acids in addition to high tensile strength.

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For efficient Southern blotting, gel pretreatment is important. Large DNA fragments (>
10 kb) require a longer transfer time than short fragments. To allow uniform transfer of
a wide range of DNA fragment sizes, the electrophoresed DNA is exposed to a short
depurination treatment (0.25 mol/l HCl) followed by alkali. This shortens the DNA
fragments by alkaline hydrolysis at depurinated sites. It also denatures the fragments
prior to transfer, ensuring that they are in the single-stranded state and accessible for
probing.

Finally, the gel is equilibrated in neutralizing solution prior to blotting. An alternative
method uses positively charged nylon membranes, which remove the need for extended
gel pretreatment. With them the DNA is transferred in native (non-denatured) form and
then alkali-denatured in situ on the membrane.
After transfer, the nucleic acid needs to be fixed to the membrane and a number of
methods are available.
Oven baking at 80°C is the recommended method for n itrocellulose membranes and
this can also be used with nylon membranes. Due to the flammable nature of
nitrocellulose, it is important that it is baked in a vacuum oven. An alternative fixation
method utilizes ultraviolet cross-linking. It is based on the formation of cross-links
between small fractions of the thymine residues in the DNA and positively charged
amino groups on the surface of nylon membranes. A c alibration experiment must be
performed to determine the optimal fixation period.
Following the fixation step, the membrane is placed in a solution of labelled (radioactive
or non-radioactive) RNA, single-stranded DNA or oli godeoxynucleotide which is
complementary in sequence to the blottransferred DNA band or bands to be detected.

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Conditions are chosen so that the labelled nucleic acid hybridizes with the DNA on the
membrane. Since this labelled nucleic acid is used to detect and locate the
complementary sequence, it is called the probe. Conditions are chosen which maximize
the rate of hybridization, compatible with a low background of non-specific binding on
the membrane. After the hybridization reaction has been carried out, the membrane is
washed to remove unbound radioactivity and regions of hybridization are detected
autoradiographically by placing the membrane in contact with X-ray film.
A common approach is to carry out the hybridization under conditions of relatively low
stringency which permit a high rate of hybridization, followed by a series of post-
hybridization washes of increasing stringency (i.e. higher temperature or, more
commonly, lower ionic strength).
Autoradiography following each washing stage will reveal any DNA bands that are
related to, but not perfectly complementary with, the probe and will also permit an
estimate of the degree of mismatching to be made.


Mapping restriction sites around a hypothetical gene sequence in total genomic DNA by
the Southern blot method. Genomic DNA is cleaved with a restriction endonuclease into
hundreds of thousands of fragments of various sizes. The fragments are separated
according to size by gel electrophoresis and blot-transferred on to nitrocellulose paper.
Highly radioactive RNA or denatured DNA complementary in sequence to gene X is
applied to the nitrocellulose paper bearing the blotted DNA. The radiolabelled RNA or

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DNA will hybridize with gene X sequences and can be detected subsequently by
autoradiography, so enabling the sizes of restriction fragments containing gene X
sequences to be estimated from their electrophoretic mobility. By using several
restriction endonucleases singly and in combination, a map of restriction sites in and
around gene X can be built up.
The Southern blotting methodology can be extremely sensitive. It can be applied to
mapping restriction sites around a single-copy gene sequence in a complex genome
such as that of humans (Fig.), and when a ‘mini-satellite’ probe is used it can be applied
forensically to minute amounts of DNA
NORTHERN BLOTTING TECHNIQUES
Southern’s technique has been of enormous value, but it was thought that it could not
be applied directly to the blot-transfer of RNAs separated by gel electrophoresis, since
RNA was found not to bind to nitrocellulose. Alwine et al. (1979) therefore devised a
procedure in which RNA bands are blot-transferred f rom the gel on to chemically
reactive paper, where they are bound covalently. The reactive paper is prepared by
diazotization of aminobenzyloxymethyl paper (creating diazobenzyloxymethyl (DBM)
paper), which itself can be prepared from Whatman 5 40 paper by a series of
uncomplicated reactions. Once covalently bound, the RNA is available for hybridization
with radiolabelled DNA probes. As before, hybridizi ng bands are located by
autoradiography.
Alwine et al.’s method thus extends that of Southern and for this reason it has acquired
the jargon term northern blotting.
Subsequently it was found that RNA bands can indeed be blotted on to nitrocellulose
membranes under appropriate conditions (Thomas 1980 ) and suitable nylon
membranes have been developed.
Because of the convenience of these more recent methods, which do not require freshly
activated paper, the use of DBM paper has been superseded.
WESTERN BLOTTING TECHNIQUES
The term ‘western’ blotting (Burnette 1981) refers to a procedure which does not directly
involve nucleic acids, but which is of importance in gene manipulation. It involves the
transfer of electrophoresed protein bands from a polyacrylamide gel on to a membrane
of nitrocellulose or nylon, to which they bind strongly (Gershoni & Palade 1982, Renart

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& Sandoval 1984). The bound proteins are then availing able for analysis by a variety of
specific protein–ligand interactions. Most commonly, antibodies are used to detect
specific antigens. Lectins have been used to identify glycoproteins. In these cases the
probe may itself be labelled with radioactivity, or some other ‘tag’ may be employed.
Often, however, the probe is unlabelled and is itself detected in a ‘sandwich’ reaction,
using a second molecule which is labelled, for instance a species-specific second
antibody, or protein A of Staphylococcus aureus (which binds to certain subclasses of
IgG antibodies), or streptavidin (which binds to antibody probes that have been
biotinylated). These second molecules may be labell ed in a variety of ways with
radioactive, enzyme or fluorescent tags.
An advantage of the sandwich approach is that a single preparation of labelled second
molecule can be employed as a general detector for different probes. For example, an
antiserum may be raised in rabbits which react with a range of mouse immunoglobins.
Such a rabbit anti-mouse (RAM) antiserum may be radiolabelled and used in a number
of different applications to identify polypeptide bands probed with different, specific,
monoclonal antibodies, each monoclonal antibody being of mouse origin. The sandwich
method may also give a substantial increase in sensitivity, owing to the multivalent
binding of antibody molecules.
Alternative blotting techniques
The original blotting technique employed capillary blotting but nowadays the blotting is
usually accomplished by electrophoretic transfer of polypeptides from an SDS-
polyacrylamide gel on to the membrane (Towbin et al. 1979). Electrophoretic transfer is
also the method of choice for transferring DNA or R NA from low-pore-size
polyacrylamide gels. It can also be used with agarose gels. However, in this case, the
rapid electrophoretic transfer process requires high currents, which can lead to
extensive heating effects, resulting in distortion of agarose gels. The use of an external
cooling system is necessary to prevent this. Another alternative to capillary blotting is
vacuumdriven blotting (Olszewska & Jones 1988), for which several devices are
commercially available. Vacuum blotting has several advantages over capillary or
electrophoretic transfer methods: transfer is very rapid and gel treatment can be
performed in situ on the vacuum apparatus. This ensures minimal gel h andling and,
together with the rapid transfer, prevents significant DNA diffusion.

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UNIT : 6 CHEMICAL SYNTHESIS OF GENES AND PCR
Phosphodiester, phosphotriester and phophite ester methods, principles and
strategies. Oligonucleotide synthesis and application, synthesis of complete gene.
PCR, methodology, essential feature of PCR, primers , Taq polymerases, reverse
transcriptase-PCR, types of PCR-Nested, inverse, RA PD-PCR, RT-PCR ( real time
PCR ), Application of PCR.


PHOSPHODIESTER METHODS PRINCIPLES AND STRATEGIES
The basic strategy of oligonucleotide synthesis is analogous to that of polypeptide
synthesis: A suitably protected nucleotide is coupled to the g rowing end of the
oligonucleotide chain, the protecting group is removed, and the process is repeated until
the desired oligonucleotide has been synthesized. The first practical technique for DNA
synthesis, the phosphodiester method, which was developed by H. Gobind Khorana in
the 1960s, is a laborious process in which all reactions are carried out in solution and
the products must be isolated at each stage of the multistep synthesis. Khorana
nevertheless used this method, in combination with enzymatic techniques, to
synthesize a 126-nucleotide tRNA gene, a project that required several years of intense
effort by numerous skilled chemists.
PHOSPHOTRIESTER METHODS PRINCIPLES AND STRATEGIES
By the early 1980s, these difficult and time-consuming processes had been supplanted
by much faster solid phase methodologies that permitted oligonucleotide synthesis to be
automated. The presently most widely used chemistry, which was formulated by Robert
Letsinger and further developed by Marvin Caruthers , is known as the
phosphoramidite method. This series of nonaqueous reactions adds a single
nucleotide to a growing oligonucleotide chain as follows (Fig.):
1.
The dimethoxytrityl (DMTr) protecting group at the 5’ end of the growing oligonucleotide
chain (which is anchored via a linking group at its 3’ end to a solid support, S) is
removed by treatment with an acid such as trichloroacetic acid (Cl
3CCOOH).
2.
The newly liberated 5’ end of the oligonucleotide is coupled to the 3’phosphoramidite
derivative of the next deoxynucleoside to be added to the chain. The coupling agent in
this reaction is tetrazole, which protonates the incoming nucleotide’s diisopropylamine
moiety so that it becomes a good leaving group. Modified nucleosides (e.g. containing a

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fluorescent label) can be incorporated into the growing oligonucleotide at this stage.
Likewise, a mixture of oligonucleotides containing different bases at this position can be
synthesized by adding the corresponding mixture of nucleosides.

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3. Any unreacted 5’ end (the coupling reaction has a yield of over 99%) is capped by
acetylation so as to block its extension in subsequent coupling reactions. This prevents
the extension of erroneous oligonucleotides (failure sequences).
4.
The phosphite triester group resulting from the coupling step is oxidized with to the
more stable phosphotriester, thereby yielding a chain that has been lengthened by one
nucleotide. This reaction sequence, in commercially available automated synthesizers,
can be repeated up to ~250 times with cycle times o f 20 to 30 min. Once an
oligonucleotide of desired sequence has been synthe sized, it is treated with
concentrated NH
4OH to release it from its support and remove its various blocking
groups, including those protecting the exocyclic amines on the bases.
5.
The product can then be separated from the failure sequences and protecting groups by
reverse-phase HPLC and/or gel electrophoresis. The largest DNA molecule yet
synthesized is the entire 582,970-bp genome of Mycoplasma genitalium (among the
smallest bacterial genomes). Initially, overlapping 5- to 7- kb “cassettes” were
constructed by sequentially ligating chemically syn thesized double-stranded
oligonucleotides of ~50 bp. The resulting 101 cassettes were then joined in stages by
utilizing their overlapping ends. Sequencing the final product confirmed that it had the
correct sequence.
OLIGONUCLEOTIDE SYNTHESIS AND APPLICATION - SYNTHES ISOF COMPLETE
GENE.
The sequencing of the human genome is really only t he means to a highly complex
end.The questions of real biochemical significance are these: What are the functions of
its ~23,000 genes? In which cells, under what circumstances, and to what extent are
each of them expressed? How do their gene products interact to yield a functional
organism? And what are the medical consequences of variant genes? The traditional
method of addressing such questions, the one-gene-a t-a-time approach, is simply
incapable of acquiring the vast amounts of data nec essary to answer these
questions.What is required are methods that can globally analyze biological processes,
that is, techniques that can simultaneously monitor all the components of a biological
system.
A technology that is capable of making such global assessments involves the use of
DNA microarrays also called DNA chips. These are arrays of different DNA molecules

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anchored to the surface of a glass, silicon, or nylon wafer in a ~1-cm-wide square grid.
In one of several methodologies that are presently used to manufacture DNA
microarrays, large numbers (up to ~1 million) of di fferent oligonucleotides are
simultaneously synthesized via a combination of photolithography (the process used to
abricate electronic chips) and solid phase DNA synthesis.
In this process (Fig.), which was developed by Stephen Fodor, the nucleotides from
which the oligonucleotides are synthesized each have a photochemically removable
protective group at their 5’ end that has the same function as the DMTr group in
conventional solid phase DNA synthesis. At a given stage in the synthetic procedure,
oligonucleotides that, for example, require a T at their next position are deprotected by
shining light on them through a mask that blocks th e light from hitting those grid
positions that require a different base at this nex t position (in an alternative
methodology called maskless array synthesis, an array of individually programmable
micromirrors directs light to the desired locations). The chip is then incubated with a
solution of activated thymidine nucleotide, which couples only to the deprotected
oligonucleotides.
After washing away the unreacted thymidine nucleotide, the process is repeated with
different masks (light patterns for maskless array synthesis) for each of the remaining
three bases. By repeating these four steps N times, an array of all 4N possible N-residue
sequences can be synthesized simultaneously in 4N coupling cycles, where N ~ 30 (up
to 100 for maskless array synthesis).
A DNA microarray is shown in Fig. In one application of DNA microrrays, L-residue
oligonucleotides (the probes) are arranged in an array of L columns by 4 rows for a total
of 4L sequences.The probe in the array’s Mth column has the “standard” sequence with
the exception of the probe’s Mth position, where it has a different base, A, C, G, or T, in
each row. Thus, one of the four probe DNAs in every column will have the standard
sequence, whereas the other three will differ from the standard DNA by only one base
change. The probe array is then hybridized with the complementary DNA or RNA, whose
variation relative to the standard DNA is to be determined, and the unhybridized DNA
or RNA is washed away. This “target” DNA or RNA is fluorescently labeled so that, when
interrogated by a laser, the positions on the probe array to which it binds are revealed
by fluorescent spots. Since hybridization conditions can be adjusted so that a single

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base mismatch will significantly reduce the level of binding, a target DNA or RNA that
varies from the complement of the standard DNA by a single base change at its Mth
position, say C to A, would be readily detected by an increased fluorescence at the row
corresponding to A in the Mth column relative to that at other positions [a target DNA or
RNA that was exactly complementary to the standard DNA would exhibit high
fluorescence in each of its columns at the base position (row) corresponding to the
standard sequence]. The intensity of the fluorescence at every position in the array, and
hence the sequence variation from the standard DNA, is rapidly determined with a
computerized fluorescence scanning device. In this manner, single nucleotide
polymorphisms (SNPs) can be automatically detected. It is becoming increasingly
apparent that genetic variations, and in particular SNPs, are largely responsible for an
individual’s susceptibility to many diseases as well as to adverse reactions to drugs.
In an alternative DNA microarray methodology, different DNAs are robotically spotted
(deposited) to precise locations on a coated glass surface. These DNAs most often
consist of PCR-amplified inserts from cDNA clones or expressed sequence tags (ESTs),
which are usually robotically synthesized.The DNAs are spotted in nanoliter-sized
droplets that rapidly evaporate leaving the DNA attached to the glass substrate. Up to
30,000 DNAs representing all of the genes in an organism can be spotted onto a single
glass “chip.” Such DNA microarrays, many of which a re commercially available, are
used to monitor the level of expression of the genes in a tissue of interest by the degree
of hybridization of its fluorescently labeled mRNA or Cdna population. They can
therefore be used to determine the pattern of gene expression (the expression profile)
in different tissues of the same organism and how specific diseases and drugs (or drug
candidates) affect gene expression. Thus, DNA microarrays are the major tool for the
study of a cell’s transcriptome (which, in analogy with the word “genome,” is the entire
collection of RNAs that the cell transcribes). Moreover, DNA microarrays are used to
specifically identify infectious agents by detecting unique segments of their DNA.
Consequently, DNA microarrays hold enormous promise for understanding the
interplay of genes during cell growth and changes i n the environment, for the
characterization and diagnosis of both infectious and noninfectious diseases (e.g.,
cancers), for identifying genetic risk factors and designing specific treatments, and as
tools in drug development.

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The photolithographic synthesis of a DNA microarray
In Step 1 of the process, oligonucleotides that are anchored to a glass or silicon surface,
and each having a photosensitive protecting group (filled red square) at their 5’ ends,
are exposed to light through a mask that only permi ts the illumination of the
oligonucleotides destined to be coupled, for example, to a T residue.The light deprotects
these oligonucleotides so that only they react with the activated T nucleotide that is
incubated with the chip in Step 2.The entire process is repeated in Steps 3 and 4 with a
different mask for G residues, and in subsequent reaction cycles for A and C residues,
thereby extending all of the oligonucleotides by one residue.This quadruple cycle is then
repeated for as many nucleotides as are to be added to form the final set of
oligonucleotides. Each “feature” (position) on the microarray contains at least 1 million
identical oligonucleotides.

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PCR
The impact of the PCR upon molecular biology has been profound. The reaction is easily
performed, and leads to the amplification of specific DNA sequences by an enormous
factor. From a simple basic principle, many variations have been developed with
applications throughout gene technology (Erlich 198 9, Innis et al. 1990). Very
importantly, the PCR has revolutionized prenatal diagnosis by allowing tests to be
performed using small samples of fetal tissue.
In forensic science, the enormous sensitivity of PCR-based procedures is exploited in
DNA profiling; following the publicity surrounding Jurassic Park, virtually everyone is
aware of potential applications in palaeontology and archaeology. Many other processes
have been described which should produce equivalent results to a PCR (for review, see
Landegran 1996) but as yet none has found widespread use.
In many applications of the PCR to gene manipulation, the enormous amplification is
secondary to the aim of altering the amplified sequ ence. This often involves
incorporating extra sequences at the ends of the amplified DNA.
METHODOLOGY
First we need to consider the basic PCR. The principle is illustrated in Fig.
The PCR involves two oligonucleotide primers, 17–30 nucleotides in length, which flank
the DNA sequence that is to be amplified. The primers hybridize to opposite strands of
the DNA after it has been denatured, and are orientated so that DNA synthesis by the
polymerase proceeds through the region between the two primers. The extension
reactions create two doublestranded target regions, each of which can again be
denatured ready for a second cycle of hybridization and extension. The third cycle
produces two doublestranded molecules that comprise precisely the target region in
double-stranded form. By repeated cycles of heat denaturation, primer hybridization
and extension, there follows a rapid exponential accumulation of the specific target
fragment of DNA. After 22 cycles, an amplification of about 106- fold is expected (Fig.),
and amplifications of this order are actually attained in practice.

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In the original description of the PCR method (Mullis & Faloona 1987, Saiki et al. 1988,
Mullis 1990), Klenow DNA polymerase was used and, because of the heat-denaturation
step, fresh enzyme had to be added during each cycle.
A breakthrough came with the introduction of Taq DNA polymerase (Lawyer et al. 1989)
from the thermophilic bacterium Thermus aquaticus. The Taq DNA polymerase is
resistant to high temperatures and so does not need to be replenished during the PCR
(Erlich et al. 1988, Sakai et al. 1988). Furthermore, by enabling the extension reaction
to be performed at higher temperatures, the specificity of the primer annealing is not
compromised. As a consequence of employing the heat-resistant enzyme, the PCR could
be automated very simply by placing the assembled reaction in a heating block with a
suitable thermal cycling programme.
The polymerase chain reaction.
In cycle 1 two primers anneal to denatured DNA at opposite sides of the target region,
and are extended by DNA polymerase to give new strands of variable length.
In cycle 2, the original strands and the new strands from cycle 1 are separated, yielding
a total of four primer sites with which primers anneal. The primers that are hybridized
to the new strands from cycle 1 are extended by polymerase as far as the end of the
template, leading to a precise copy of the target region.
In cycle 3, double-stranded DNA molecules are produced (highlighted in colour) that are
precisely identical to the target region.
Further cycles lead to exponential doubling of the target region.

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The original DNA strands and the variably extended strands become negligible after the
exponential increase of target fragments.

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ESSENTIAL FEATURE OF PCR, PRIMERS
The specificity of the PCR depends crucially upon the primers. The following factors are
important in choosing effective primers.
•
Primers should be 17 to 30 nucleotides in length.
•
A GC content of about 50% is ideal. For primers with a low GC content, it is desirable to
choose a long primer so as to avoid a low melting temperature.
•
Sequences with long runs (i.e. more than three or four) of a single nucleotide should be
avoided.
•
Primers with significant secondary structure are undesirable.
•
There should be no complementarity between the two primers.
The great majority of primers which conform to these guidelines can be made to work,
although not all comparable primer sets are equally effective even under optimized
conditions.
In carrying out a PCR it is usual to employ a hot-start protocol. This entails adding the
DNA polymerase after the heat-denaturation step of the first cycle, the addition taking
place at a temperature at or above the annealing temperature and just prior to the
annealing step of the first cycle. The hot start overcomes the problem that would arise if
the DNA polymerase were added to complete the assem bly of the PCR reaction mixture
at a relatively low temperature. At low temperature, below the desired hybridization
temperature for the primer (typically in the region 45–60°C), mismatched primers will
form and may be extended somewhat by the polymerase .
Once extended, the mismatched primer is stabilized at the unintended position. Having
been incorporated into the extended DNA during the first cycle, the primer will
hybridize efficiently in subsequent cycles and hence may cause the amplification of a
spurious product.
Alternatives to the hot-start protocol include the use of Taq polymerase antibodies,
which are inactivated as the temperature rises (Taylor & Logan 1995), and AmpliTaq
GoldTM, a modified Taq polymerase that is inactive until heated to 95°C (Birch 1996).
Yet another means of inactivating Taq DNA polymerase at ambient temperatures is the
SELEX method (systematic evolution of ligands by exponential enrichment). Here the
polymerase is reversibly inactivated by the binding of nanomolar amounts of a 70-mer,

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which is itself a poor polymerase substrate and sho uld not interfere with the
amplification primers (Dang & Jayasena 1996).
As noted above, the Taq DNA polymerase lacks a 3′–5′ proofreading exonuclease. This
lack appears to contribute to errors during PCR amplification due to misincorporation
of nucleotides (Eckert & Kunkel 1990). Partly to ov ercome this problem, other
thermostable DNA polymerases with improved fidelity have been sought, although the
Taq DNA polymerase remains the most widely used enzyme for PCR. In certain
applications, especially where amplified DNA is cloned, it is important to check the
nucleotide sequence of the cloned product to reveal any mutations that may have
occurred during the PCR. The fidelity of the amplification reaction can be assessed by
cloning, sequencing and comparing several independently amplified molecules.
TAQ POLYMERASES
The thermostable DNA-dependent DNA polymerase I, Ta q DNA polymerase (Taq pol), is
the most extensively used DNA polymerase for amplif ying genetic material by the
polymerase chain reaction (PCR) and for sequencing DNA. Taq pol is obtained from
Thermus aquaticus, an archeabacterium that thrives at elevated temperatures in deep
thermal vents, and the enzyme is highly resistant t o denaturation at elevated
temperatures. Taq DNA polymerase is closely related to DNA polymerase I of Escherichia
coli in its primary and three-dimensional, tertiary structures.
Both are single subunit enzymes with C-terminal pol ymerase and N-terminal 5′-3′
exonuclease domains. However, Taq pol lacks the 3′-5′ exonuclease (proofreading)
function found in E. coli pol I and therefore cannot excise polymerization errors incurred
during synthesis.
Properties
Taq pol has been cloned and overexpressed in E. coli. The purified full-length protein is
composed of a single subunit of 94 kDa with two distinct activities: 5′-3′ polymerase
and 5′-3′ exonuclease. Taq DNA polymerase is highly thermostable, loses only 10% of its
activity after 30 min incubation at 72 °C (which is the normal growth temperature for
Thermus aquaticus), and 50% activity after 30 minute incubation at 95 °C (M. Suzuki
and L.A. Loeb, unpublished results). Like DNA pol I of E. coli, Taq pol is susceptible to
proteolytic cleavage and yields a small N-terminal fragment that has 5′-3′ exonuclease
activity and a large C-terminal fragment that has p olymerase activity. The large

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Cterminal fragment is analogous to the Klenow fragment of DNA pol I and is also called
the KlenTaq or Stoffel fragment. It is thermostable, loses only 10% activity after 30 min
incubation at 95 °C, and therefore is frequently used in polymerase chain reactions
(PCR) and various sequencing protocols that involve prolonged incubations at elevated
temperatures.
Taq pol conducts DNA-templated DNA synthesis with m oderate accuracy, on average
misincorporating only 1 in 9000 nucleotides (2). The predicted fidelity during PCR is
one error per 400 bases after 25 cycles. The fidelity during PCR is enhanced twofold by
using the truncated KlenTaq fragment instead of full-length Taq pol (3) and up to 10-
fold by using other thermostable polymerases that h ave a 3-5′ exonuclease
(proofreading) function. Specific methods that enhance the fidelity of Taq pol include
incubation at low pH (5 to 6) or with reduced concentrations of MgCl
2. However, both of
these methods decrease Taq pol activity and are gen erally incompatible with PCR,
especially for amplifying long fragments.
The two components of fidelity include
• Nucleotide misinsertion and
• Extension of the misinserted base.
• Taq pol extends mispairs at a very low efficiency: 10
–3
for a T–G mispair to 10
–6
for A–A
mispair
• Because of its relative specificity to extend only properly matched primers, Taq pol is
frequently used to detect specific in vivo mutations.
Structure
The X-ray crystallographic structures of all polymerases determined to date, including
Taq DNA polymerase, resemble in overall morphology a cupped human right hand,
complete with subdomains corresponding to the finge rs (which bind the single
stranded-template), palm (which binds the incoming deoxynucleoside triphosphate,
DNTP) and thumb (which binds double-stranded DNA) (Fig. 1).
The polymerase subdomains of Taq pol and the Klenow fragment are highly homologous
and have 51% amino acid identity. In addition, the overall folding of the two polymerase
domains in the three-dimensional structures are virtually identical and have an average
(root mean square) deviation in alpha carbons of ap proximately 1.4 Å. The overall
folding and location of the 3′-5′ exonuclease domain, which is 30 Å away from the

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polymerase active site, is also similar in both enzymes. Taq pol, however, lacks four
loops found in the Klenow fragment on one side of the 3′-5′ exo site. In addition, four
key amino acid residues believed to bind metal cofactors involved in the exonuclease
reaction in the Klenow fragment (Asp424, Asp501, Asp355, and Glu357) are replaced by
nonacidic residues in Taq pol (Leu356, Arg405, Gly308, Val310). These changes may
explain why the 3′-5′ nuclease domain of Taq pol is inactive. A direct comparison of the
Klenow and Taq pol structures also shows that Taq contains a more hydrophobic core
and more favorable electrostatic interactions. Both of these properties may contribute to
the greater thermostability of Taq pol.

Interestingly, the location of the 5′-3′ exonuclease active site, which probably functions
in nick translation during DNA repair and during removal of Okazaki Fragments, is 70
Å from the polymerase active site. How polymerase and 5′-3′ exonuclease sites so far
apart work in concert to leave a “repaired” double-stranded DNA with only a nick
remains a mystery. The structure of Taq pol bound t o blunt-end DNA has been
described. Similar to the complexes with DNA of pol b-DNA and HIV reverse
transcriptase, the DNA in the Taq pol active site adopts a structure that is a hybrid
between A and B forms. As a consequence, the minor groove, which interacts with
amino acid residues in the polymerase active site, is especially wider than in B-form
DNA. It is thought that protein side-chains may form hydrogen bonds with the O
2 atom
of the pyrimidine ring and with the N3 atoms of purines.
Because the positions of these two atoms are unchanged in A:T and G:C base pairs,
these putative hydrogen-bond interactions between protein side chains and the last

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base pair may insure proper base pairing before nuc leotide incorporation and thus
enhance the fidelity of the polymerase.
Uses
Taq pol is used extensively in polymerase chain reactions to amplify genetic material.
PCR involves incubation with the gene that is to be amplified in the presence of a
polymerase, PCR primers that flank the gene of interest, all four dNTP, and magnesium.
Briefly, each cycle of PCR involves three steps: (1) incubation at elevated temperature to
separate the double-stranded DNA (typically 95°C); (2) incubation at lower temperature
to allow primer/template annealing, and (3) incubat ion at a temperature for
polymerization. Before the discovery of Taq pol, PCR involved manually adding a small
amount of polymerase to the incubation mixture befo re each polymerization step
because the polymerization was inactivated during incubation at elevated temperatures.
The thermostability of Taq allows heat denaturation of DNA after each cycle without
enzyme inactivation, thus allowing automation of PC R. In general, current PCR
technology using full-length Taq pol allows amplification from one molecule to more
than 105 copies of a target sequence, which can be as large as 5 kilobases. Even larger
sequences are amplified by combining a truncated KlenTaq fragment with low levels of a
thermostable polymerase that contains 3′-5′ proofre ading activity (eg, Vent DNA
polymerase or Pfu polymerase).
PCR is used widely in research and clinical laborat ories for diverse procedures
including amplifying genes for cloning, diagnosis of diseases, and detecting levels of
viruses. Taq DNA polymerase, which possesses weak R NA-templated DNA polymerase
activity, is also used to synthesize doublestranded DNA from messenger RNA templates.
However, this process, termed RT-PCR, is generally more efficient if a reverse
transcriptase is used during the first cycle to generate singlestranded
DNA and Taq pol is used in subsequent amplification cycles. Alternatively, a polymerase
from the thermophilic bacteria Thermus thermophilus (TTH POL), which contains
efficient RNA-templated and DNA-templated DNA polym erase activities in the presence
of Mn2+, can be used for RT-PCR. Because of its rel ative specificity to extend only
properly matched primers, Taq pol I is used to detect specific mutations through PCR
analysis, simply by choosing primers with a 3′ -terminus that is complementary to the
mutant. Efficient amplification of the DNA in this procedure suggests the presence of a

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specific mutation. This protocol has been used successfully to detect mutations in the
ras oncogene and common mutations resulting in inherited disorders.
Taq Pol in DNA Sequencing
Until recently, enzymatic DNA sequencing using Taq pol was marred because
dideoxynucleotides (DDNTP) are not efficiently incorporated by Taq pol (its usage of
ddNTP is 1000 times less efficient than that of dNTP in the presence of Mg2+; However,
it has been recently shown that the substitution Ph e667Tyr of Taq pol increases
incorporation of ddNTP relative to dNTP 250 by 8000-fold. This mutated Taq (Thermo
sequanase) enables cycle sequencing, thus producing accurate and analyzable
sequences using either radioactive or fluorescent sequencing technologies.
Taq Pol in T/A Cloning
Taq pol shares a characteristic common to all polymerases that lack 3′-5′ exonuclease
proofreading activity in that it incorporates nontemplated nucleotides (usually adenine)
onto blunt-end DNA. This property is frequently used in molecular biology to ligate
fragments of DNA in T/A cloning protocols. Briefly, PCR amplification of the sequence of
interest results in a significant proportion of products containing a nontemplated
adenine incorporated onto both 3′ ends. Then this PCR product is incubated in the
presence of a ligase with a linear vector that contains 3′ thymine residues. The
completed reaction results in a circular DNA that contains a cloned insert.
REVERSE TRANSCRIPTASE
In all organisms the major flow of genetic information is from DNA to RNA using DNA-
dependent RNA polymerases, with the DNA itself being replicated by DNA-dependent
DNA polymerases . Some of the DNA synthesis, primarily of specific viruses
(retroviruses) and retroelements, is from RNA by reverse transcriptase. However, there
are many viruses of both prokaryotes and eukaryotes that have RNA genomes and that
replicate by the synthesis of RNA from the genome. This replication is performed by
RNA-dependent RNA polymerases (EC 2.7.7.48) (RdRp) (also termed “RNA replicases”)
and usually involves other additional enzyme activities such as RNA Helicases and
capping and methylating enzymes; these are frequently on the same protein molecule
as the polymerase itself. There is also some evidence for RdRp activity in uninfected
cells, and there was initially considerable controversy as to whether RNA viruses were
replicated by host or viral enzyme (s)

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RdRp can be considered to be an ancient enzymatic activity. It is widely suggested that,
at an early stage in the evolution of life, there was the “RNA World” with the initial
synthetic and processing activities being performed by catalytic RNAs, or ribozymes. In
this scene, the early proteins would have been bett er enzymes than their RNA
counterparts and thus would eventually dominate. It is likely that RdRp activity was
among these early proteins.
Uninfected Cells
RdRp activity has been found in uninfected cells from both animals and plants. Mouse
erythroleukemia cells contain a cytoplasmic RNA synthesis activity which is resistant to
actinomycin D, leads to the formation of (–) strand globin messenger RNA and can use
poly(A)-oligo(U) as a template-primer combination. This activity required Mg2+ but was
inhibited by Mn2+. The best-characterized plant RdRp is that from tomato leaves. This
128-kDa polypeptide chain sedimented at 6.6 S in its native form, which indicated that
it was probably a monomer. Its optimum activity was at pH 7.8. It required a divalent
cation, Mg2+, which is much more effective than Mn2+.
The enzyme activity could transcribe RNA from both RNA and DNA templates, most of
the transcripts being very short. Transcription could take place either with or without a
primer; in the latter case priming was at, or close, to the 3′ end of the template.
As noted above, there was controversy about the rol e of host RdRp in viral RNA
replication, which was exacerbated by the fact that the host RdRp activity increases
upon virus infection. However, it is now generally assumed that the host RdRp is not
involved, at least significantly, in viral replication.
Its function is unknown, but the short lengths of RNA that it transcribes might be
involved in some control mechanism, such as post-transcriptional suppression of gene
expression.
Viral RdRps
Viral RdRps have three remarkable features: (i) they amplify their RNA template many-
fold during a very short infection period; (ii) they specifically replicate the viral genomic
RNA in the presence of a great excess of host RNAs; (iii) they copy the entire template
RNA, and also express subgenomic RNAs, in most case s without using endogenous
primers. In all viral enzyme complexes, the catalytic polymerase activity is virus-
encoded. The detailed mechanisms by which viral RNA replicases operate are unknown,

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but there would appear to be differences between them, as is exemplified by the current
understanding of different viruses.
Bacteriophage Qb
The RdRp holoenzyme of coliphage Qb is capable of in vitro synthesis of (–)-strand RNA
from a (+)-strand template; it consists of four subunits, three of which are host-derived
proteins [ribosomal protein S1 (subunit a) and translation elongation factors EF-Tu(g)
and EF-TS(d)], and the fourth(b) is virus-encoded and is the catalytic subunit. Details of
the function and structure of the basic holoenzyme are reviewed by Blumenthal and
Carmichael and by Ishihama and Barbier and a workin g model has been proposed.
Recent evidence indicates that a host factor is also required for efficient in vivo RNA
replication. As with other RdRps, that of Qb is template-specific, and the template
recognition involves complex interactions between the holoenzyme and the RNA. The
tertiary structure of the RNA is recognized as being important in bringing together the
sites with which the proteins of the holoenzyme interact. Recent studies suggest that
template recognition and specificity involve both the carboxyl-terminal region of the
virus-encoded subunit and the hostderived S1 protein.
Poliovirus
The core enzyme of poliovirus RdRp (the 3D gene product) is processed by the adjacent
gene product (3C) from the polyprotein translated from the viral genome, and it forms
a membraneassociated complex with the primer VPg protein (which covalently attaches
to the 5′ end of the viral RNA) and protein 3C. Although there is not a clear picture of
the detailed mechanism by which poliovirus, and other picornaviruses, replicates their
RNA, there are various observations that have to be incorporated in any model. These
include: (i) the need for the replication complexes to be membrane-associated for
initiation and elongation of minus- and plus-strand synthesis; (ii) the VPg is covalently
linked to all newly synthesized RNA; (iii) the primer for the processive RdRp is unknown
although there are various hypotheses; (iv) the precursor 3CD protein binds to the 5′-
terminal cloverleaf structure of the plus strand in the presence of cellular and/or viral
factors; (v) various cellular proteins have been implicated in RNA synthesis, including a
report that human protein Sam68 interacts with poliovirus 3D protein.
Plant (+)-Strand RNA Viruses

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Understanding of the functioning of the RdRps of plant (+)-strand RNA viruses is limited
because of the difficulties in isolating actively replicating systems from the membrane-
bound replication complexes in infected cells. The replication complexes of several
viruses have been shown to contain host-encoded pro teins, as well as the predicted
virus-encoded products. The replication complex of tobacco mosaic virus (TMV)
associates with a protein related to GCD10 protein, which is the RNA-binding subunit
of yeast eIF-3 and that of brome mosaic virus associates with the 41-kDa subunit of
wheat eIF-3; the replication complex of cucumber mosaic virus is associated with a 50-
kDa host protein. As the TMV and brome mosaic virus replicase complexes are
associated with different eIF-3 subunits, it is thought likely that these host proteins
play different roles in their respective virus replications.
The replication of RNA by RdRp is considered to have several stages covering initiation
to elongation. There is recent evidence pointing towards the involvement of the transfer
RNAlike structures at the 3′ end of virion RNAs of several plant viruses in the initiation
and priming of at least (–)-strand synthesis.
Double-Stranded RNA-Viruses and (–)-Strand RNA Viru ses
As the genomes of double-stranded or (–)-strand RNA viruses cannot be translated
directly, the virus particles carry the RdRp to initiate synthesis of mRNA on entry into
the infected cell. The RdRp of influenza virus, which has a ( –)-strand segmented
genome, has essentially two functions, the synthesi s of mRNA from the genome
segments and the replication of the virion RNA via the complementary RNA. Influenza
mRNA synthesis is primed at the 3′ end of the virion RNA by 10- to 13-nucleotide
capped RNA fragments that are “cap-snatched” from the 5′ ends of host heterogeneous
nuclear RNA (hnRNA). The exact details of priming of the complementary RNA are not
yet fully understood.
Groups of RdRps
Kamer and Argos recognized various sequence motifs characteristic of RdRps. The most
conserved of these is the central Gly-Asp-Asp (GDD) triplet flanked by 5-residue
segments that are mainly hydrophobic amino acids. This is taken as being suggestive
of a b-hairpin structure comprising two hydrogen-bonded antiparallel beta-strands
connected by a short exposed loop containing the GDD amino acids. Mutation of the
GDD box can abolish the replicase function.

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Alignment of the RdRp sequences has been extended w ith the recognition of eight
motifs. This led to the classification of the RdRps into three supergroups (Table 1), with
a number of lineages within each supergroups. The supergroups extend across viruses
that infect animals, plants and bacteria and bring together viruses whose genome
structures and expression have properties in common. However, these groupings have
not been fully supported by other analyses and care has to be taken in deriving
evolutionary relationships from these analyses.
Grouping of RdRps
Supergroup 1
1 genome segment. - Polyprotein expression - VPg
Examples
Animals: Picornaviruses, Arteriviruses, Astroviruses, Caliciviruses, Coronaviruses
Insects: Nodaviruses
Plants: Comoviruses, Enamovirus RNA1, Luteoviruses subgrou p II, Potyviruses,
Umbraviruses, Sequiviruses, Sobemoviruses
Fungus: Barnaviruses
Supergroup II
1 to several genome segments - Individual gene translation - Capped RNA
Examples
Bacteria: Leviviruses
Animals: Flaviviruses, Pestiviruses
Plants: Dianthoviruses, Enamovirus RNA2, Luteoviruses subgroup I, Machlomoviruses,
Necroviruses, Tombusviruses
Supergroup III
1 to several genome segments - Individual gene translation - Capped RNA
Examples
Animals: Togaviruses, Caliciviruses
Plants: Bromoviruses, Capilloviruses, Carlaviruses, Closte roviruses, Furoviruses,
Hordeiviruses, Idaeoviruses, Potexviruses, Tobamoviruses, Tobraviruses
Structure
The three-dimensional structure of the RdRp of poliovirus has been determined by X-
ray crystallography to 0.26 nm (2.6 Å) resolution. The overall shape of this polymerase

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resembles those of other polymerases, being likened to a right hand. The palm domain
that contains the catalytic core structure is very similar to that of other polymerases,
but the structures of the “fingers” and the “thumb” subdomains differ from those of
other polymerases. Extensive regions of interactions between neighboring molecules
were observed in the crystals, which suggests that an unusual higher order structure
might be important in polymerase function.
Mechanism Action
As noted above, it has been suggested that the functioning of RdRps has several stages.
By analogy with the functioning of DNA-dependent DN A polymerases, these were
proposed as being: (i) template binding, (ii) promoter localization, (iii) melting the
template to give a transcriptionally open complex, (iv) nucleotide substrate binding, (v)
formation of the first phosphodiester bond, (vi) promoter clearance, and (vii) progressive
elongation. Relatively little is known about any of these functional stages for RdRps,
and it is likely that there will be differences between the various virus systems for at
least some of the stages. For example, there is evidence for priming of some of the plant
(+)-strand viruses at the tRNA-like 3′ terminal structures, whereas other plant viral
RNAs do not have these structures. Also, there appe ar to be two different priming
systems for influenza mRNA and complementary RNA sy nthesis. There also has to be
priming for the formation of the (–) strand, most likely at the 3′ end of the ( + )-strand,
and then for the synthesis of the (+) strand, again most likely at the 3′ end of the ( – )-
strand. The latter is the complement of the 5′ end of the
(+)-strand and, for most viruses, the 5′ and 3′ gen ome sequences bear little
resemblance.
This lack of detailed information is mainly due to the difficulties of studying these
enzymes systems, especially those of eukaryotes that are associated with membranes.
Furthermore, as well as the RdRp itself, the replication complex also contains several
other virus-encoded and host-encoded activities. Some of the other virus-encoded
activities, such as helicases (probably involved in the melting of the template stage),
capping, and methylation enzymes are reviewed by Buck.
RdRps lack proofreading activity, and thus, there is a high rate of error in the synthesis
of the new RNA strand. It is estimated that 1 in 10 3 to 104 nucleotides is

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misincorporated, which gives a high rate of genome mutation. This, coupled with the
high rate of replication, can lead to rapid evolution of RNA viruses.
PCR TYPES
Recent developments have sought to minimize amplification times. Such systems have
used small reaction volumes in glass capillaries to give large surface area-to-volume
ratios. This results in almost instantaneous temperature equilibration and minimal
annealing and denaturation times. This, accompanied by temperature ramp rates of 10–
20°C/s, made possible by the use of turbulent force d hot-air systems to heat the
sample, results in an amplification reaction completed in tens of minutes.
While the PCR is simple in concept, practically there are a large number of variables
which can influence the outcome of the reaction. This is especially important when the
method is being used with rare samples of starting material or if the end result has
diagnostic or forensic implications. For a detailed analysis of the factors affecting the
PCR, the reader should consult McDowell (1999). There are many substances present in
natural samples (e.g. blood, faeces, environmental materials) which can interfere with
the PCR, and ways of eliminating them have been rev iewed by Bickley and Hopkins
(1999).
RT-PCR
The thermostable polymerase used in the basic PCR requires a DNA template and hence
is limited to the amplification of DNA samples. There are numerous instances in which
the amplification of RNA would be preferred. For example, in analyses involving the
diffierential expression of genes in tissues during development or the cloning of DNA
derived from an mRNA (complementary DNA or cDNA), particularly a rare mRNA. In
order to apply PCR methodology to the study of RNA, the RNA sample must first be
reverse-transcribed to cDNA to provide the necessar y DNA template for the
thermostable polymerase. This process is called reverse transcription (RT), hence the
name RT-PCR. Avian myeloblastosis virus (AMV) or Mo loney murine leukaemia virus
(MuLV) reverse transcriptases are generally used to produce a DNA copy of the RNA
template. Various strategies can be adopted for first-strand cDNA synthesis (Fig.).

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Long accurate PCR (LA-PCR)
Amplification of long DNA fragments is desirable for numerous applications of gene
manipulation. The basic PCR works well when small f ragments are amplified. The
efficiency of amplification and therefore the yield of amplified fragments decrease
significantly as the size of the amplicon increases over 5 kb. This decrease in yield of
longer amplified fragments is attributable to partial synthesis across the desired
sequence, which is not a suitable substrate for the subsequent cycles. This is
demonstrated by the presence of smeared, as opposed to discrete, bands on a gel.
Barnes (1994) and Cheng et al. (1994) examined the factors affecting the thermostable
polymerization across larger regions of DNA and identified key variables affecting the
yield of longer PCR fragments. Most significant of these was the absence of a 3′–5′
exonuclease (proofreading) activity in Taq polymerase. Presumably, when the Taq
polymerase misincorporates a dNTP, subsequent exten sion of the strand either
proceeds very slowly or stops completely. To overco me this problem, a second
thermostable polymerase with proofreading capability is added. Thermostable DNA
polymerases with proofreading capabilities are listed in Table.

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PCR-NESTED
In order to minimize further the amplification of spurious products, the strategy of
nested primers may be deployed. Here the products of an initial PCR amplification are
used to seed a second PCR amplification, in which one or both primers are located
internally with respect to the primers of the first PCR. Since there is little chance of the
spurious products containing sequences capable of hybridizing with the second primer
set, the PCR with these nested primers selectively amplifies the sought-after DNA.
INVERSE PCR
Inverse polymerase chain reaction (IPCR). IPCR is another method for isolating host
sequences flanking a transposon insertion. As for plasmid rescue, genomic DNA from a
tagged individual is isolated and digested with a restriction enzyme that releases the
end of the transposon and a piece of adjoining host DNA. Ligation is used to circularize
the linear host DNA–transposon fragment. PCR using two transposon-specific
oligonucleotide primers, each reading outward from the ends of the transposon
sequence into the flanking DNA, are used to amplify the flanking host DNA. Then the
amplified product is cloned.
Another approach that uses incomplete sequence information to amplify a target gene is
inverse PCR. In this case a sequence of part of a long DNA mol ecule, say a
chromosome, is known. The objective is to extend the analysis along the DNA molecule
into the unknown regions. To synthesize the primers for PCR, the unknown target
sequence must be flanked by two regions of known sequence. The present situation is
exactly the opposite of that. To circumvent this problem, the target molecule of DNA is
converted into a circle. Going around a circle brings you back to the beginning. In
effect, even though, only one small stretch of sequence is known, the circular form
allows you to have that one region on both sides of the target sequence.
A restriction enzyme, usually one that recognizes a six-base sequence, is used to make
the circle. This enzyme must not cut into the known sequence, therefore, eventually;
this enzyme will cut either upstream or downstream from the known region. The
resulting fragment will have unknown sequence first , the known sequence in the
middle, followed by more unknown sequence. The two ends of the fragment will have
compatible sticky ends that are easily ligated together to make a circle of DNA (Fig.).
Two primers corresponding to the known region and facing outwards around the circle

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are used for PCR. Synthesis of new DNA will proceed around the circle clockwise from
one primer and counter-clockwise from the other. Overall, inverse PCR gives multiple
copies of a segment of DNA containing some DNA to the right and some DNA to the left
of the original known region.

Inverse PCR allows unknown sequences to be amplified by PCR provided that they are
located next to DNA whose sequence is already known . The DNA is cut with a
restriction enzyme that does not cut within the region of known sequence, as shown in
Step I. This generates a fragment of DNA containing the known sequence flanked by two
regions of unknown sequence. Since the fragment has two matching sticky ends, it may
be easily circularized by DNA ligase.

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Finally, PCR is performed on the circular fragments of DNA (Step 2). Two primers are
used that face outwards from the known DNA sequence. PCR amplification gives a
single linear product that includes unknown DNA from both left and right sides. This
PCR product can now be cloned and/or sequenced.
RAPD-PCR,
Randomly Amplified Polymorphic DNA, or RAPD , is usually found in the plural as
RAPDs and is pronounced “rapids,” partly because it is a quick way to get a lot of
information about the genes of an organism under investigation. The purpose of RAPDs
is to test how closely related two organisms are. In practice, DNA samples from
unknown organisms are compared with DNA from a prev iously characterized organism.
For example, traces of blood from a crime scene may be compared to possible suspects,
or disease-causing microorganisms may be related to known pathogens to help trace an
epidemic.
The principle of RAPDs is statistically based. Given any particular five-base sequence,
such as ACCGA, how often will this exact sequence a ppear in any random length of
DNA? Since there are four different bases to choose from, one in every 45 (or 4 X 4 X 4
X 4 X 4 = 1,024) stretches of five bases will—on average—be the chosen sequence. Any
arbitrarily chosen 11-base sequence will be found once in approximately every 4 million
bases. This is approximately the amount of DNA in a bacterial cell. In other words, any
chosen 11-base sequence is expected to occur by cha nce once only in the entire
bacterial genome. For higher organisms, with much m ore DNA per cell, a longer
sequence would be needed for uniqueness.
For RAPDs, the arbitrarily chosen sequence should b e rare but not unique. PCR
primers are made using the chosen sequence and a PCR reaction is run using the total
DNA of the organism as a template. Every now and th en a primer will find a correct
match, purely by chance, on the template DNA. For PCR amplification to occur there
must be two such sites facing each other on opposite strands of the DNA. The sites
must be no more than a few thousand bases apart for the reaction to work well. The
likelihood of two correct matches in this arrangement is quite low.
In practice, the length of the primers is chosen to give five to 10 PCR products.
For higher organisms, primers of around 10 bases are typical.The bands from PCR are
separated by gel electrophoresis to measure their sizes. The procedure is repeated

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several times with primers of different sequence. The result is a diagnostic pattern of
bands that will vary in different organisms, depending on how closely they are related.
Although we do not know in which particular genes the PCR bands originate, this does
not matter in measuring relatedness. Diagnosis therefore relies on having a primer (or
set of primers) that reliably give a band of a particular size with the target organism and
give different bands with other organisms, even those closely related. RAPD results
using such a primer are shown in Figure. Grey mold, due to Botrytis cinerea, is one of
the most destructive infections of strawberries and also attacks other plants. Classical
diagnosis involves culturing the fungus on nutrient agar. It is slow and difficult due to
the presence on the plants of other harmless fungi, which often grow faster in culture.
As can be seen, RAPD analysis clearly identifies the pathogens from other related fungi,
including other species from the genus Botrytis.

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The first step of RAPD analysis is to design primers that will bind to genomic DNA at
random sites that are neither too rare nor too common. In this example, the primers
were sufficiently long to bind the genomic DNA at a dozen places. For PCR to be
successful, two primers must anneal at sites facing each other but on opposite strands.
In addition, these paired primer sites must be close enough to allow synthesis of a PCR
fragment in a reasonable time. In our example, there are three pairs but only two of
these pairs were close enough to actually make the PCR product. Consequently, this
primer design will result in two PCR products as seen in the first lane of the gel (marked
“First organism”). The same primers are then used to amplify genomic DNA from other
organisms that are suspected of being related. In this example, suspect #2 shows the
same banding pattern as the first organism and is presumably related. The other two
suspects do not match the first organism and are therefore not related.
RT-PCR (REAL TIME PCR)
As the name suggests, real time PCR is a technique used to monitor the progress of a
PCR reaction in real time. At the same time, a relatively small amount of PCR product
(DNA, cDNA or RNA) can be quantified. Real Time PCR is based on the detection of the
fluorescence produced by a reporter molecule which increases, as the reaction
proceeds. This occurs due to the accumulation of the PCR product with each cycle of
amplification. These fluorescent reporter molecules include dyes that bind to the
double-stranded DNA (i.e. SYBR® Green) or sequence specific probes (i.e. Molecular
Beacons or TaqMan® Probes).
Real time PCR facilitates the monitoring of the reaction as it progresses. One can start
with minimal amounts of nucleic acid and quantify the end product accurately.
Moreover, there is no need for the post PCR processing which saves the resources and
the time. These advantages of the fluorescence based real time PCR technique have
completely revolutionized the approach to PCR-based quantification of DNA and RNA.
In a real time PCR protocol, a fluorescent reporter molecule is used to monitor the
PCR as it progresses. The fluorescence emitted by the reporter molecule manifolds as
the PCR product accumulates with each cycle of amplification. Based on the molecule
used for the detection, the real time PCR techniques can be categorically placed under
two heads:

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Non-specific Detection using DNA Binding Dyes
Specific Detection Target Specific Probes
Non-specific Detection using DNA Binding Dyes:

In real time PCR, DNA binding dyes are used as fluorescent reporters to monitor the
real time PCR reaction. The fluorescence of the reporter dye increases as the product
accumulates with each successive cycle of amplification. By recording the amount of
fluorescence emission at each cycle, it is possible to monitor the PCR reaction during
exponential phase. If a graph is drawn between the log of the starting amount of
template and the corresponding increase the fluores cence of the reporter dye
fluorescence during real time PCR, a linear relationship is observed.

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SYBR® Green is the most widely used double-strand DNA-specific dye reported for real
time PCR. SYBR® Green binds to the minor groove of the DNA double helix. In the
solution, the unbound dye exhibits very little fluorescence. This fluorescence is
substantially enhanced when the dye is bound to dou ble strand DNA. SYBR® Green
remains stable under PCR conditions and the optical filter of the thermocycler can be
affixed to harmonize the excitation and emission wavelengths.
Although these double-stranded DNA-binding dyes pro vide the simplest and cheapest
option for real time PCR, the principal drawback to intercalation based detection of PCR
product accumulation is that both specific and nonspecific products generate signal.
Specific Detection using Target Specific Probes:
Specific detection of real time PCR is done with some oligonucleotide probes labeled
with both a reporter fluorescent dye and a quencher dye. Probes based on different
chemistries are available for real time detection, these include:
a. Molecular Beacons
b. TaqMan® Probes
c. FRET Hybridization Probes
d. Scorpion® Primers
Molecular Beacons: Molecular beacons are single stranded hairpin shap ed
oligonucleotide probes. In the presence of the target sequence, they unfold, bind and
fluoresce. The molecular beacon chemistry is one of the chemistries used to carry out a
real time experiment.
Structure: A molecular beacon consists of 4 parts, namely
• Loop: This is the 18-30 base pair region of the molecula r beacon which is
complementary to the target sequence.
• Stem: The beacon stem sequence lies on both the ends of the loop. It is typically 5-7 bp
long at the sequences at both the ends are complementary to each other.

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• 5' fluorophore: Towards the 3' end of the molecular beacon, is attached a dye that
fluoresces in presence of a complementary target.
• 3' quencher (non fluorescent): The quencher dye is covalently attached to the 3' end of
the molecular beacon and when the beacon is in clos ed loop shape, prevents the
fluorophore from emitting light.


Molecular Beacons Functioning
Molecular beacons can report the presence of specif ic nucleic acids from a
homogeneous solution. In the presence of a complementary target, the "stem" portion of
the beacon separates out resulting in the probe hybridizing to the target. In the absence
of a complimentary target sequence, the beacon rema ins closed and there is no
appreciable fluorescence. When the beacon unfolds i n the presence of the
complementary target sequence, the fluorophore is n o longer quenched, and the
molecular beacon fluoresces. The fluorescence is easily detected in a thermal cycler.
The amount of fluorescence at any given cycle, or following cycling, depends on the
amount of specific product. For quantitative PCR, m olecular beacons bind to the
amplified target following each cycle of amplification and the resulting signal is
proportional to the amount of template. Fluorescence is monitored and reported during
each annealing step when the beacon is bound to its complementary target. This
information is then used during PCR or RT-PCR (reve rse transcriptase PCR)
experiments to quantify initial copy number.

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Applications of Molecular Beacons Include
1. SNP detection
2. Real-time nucleic acid detection
3. Real-time PCR quantification
4. Allelic discrimination and identification
5. Multiplex PCR assays
6. Diagnostic clinical assays
Molecular Beacons Vs Linear Probes
While other systems use fluorescence to detect the accumulation of PCR product,
molecular beacons add another level of specificity due to the presence of a distinct
molecular probe apart from the primers. In addition, the stem probe structure of a
molecular beacon makes it better able to discriminate single base-pair mismatches
(compared to linear probes) because the hairpin makes mismatched hybrids thermally
less stable than hybrids between the corresponding linear probes and their mismatched
target.
Furthermore, unlike linear hydrolysis probes, quenching of molecular beacons has been
shown to occur through a direct transfer of energy from the fluorophore to quencher.
Consequently, a common quencher molecule can be use d, increasing the number of
possible fluorophores that can easily be used as re porters. This is an important
advantage when designing Real time PCR experiments in which several molecular
beacons with different coloured fluorophores are used to detect multiple targets in the
same tube (multiplexing).
The use of molecular beacons in diagnostic assays h as thus been ever increasing.
Diagnostic assays that aim at detecting single nucleotide polymorphisms, screening
genetically diverse species and developing drugs through pharmacogenetic applications
are now using molecular beacons based real time PCR assays. The ability of molecular
beacons to fluoresce when bound specifically to a double stranded target and their
accuracy to discriminate alleles, make them a powerful tool in the research lab armory.
Molecular beacons can be successfully used to ascer tain not only the presence or

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absence of a particular causative agent, but also in screening which antibodies will be
effective against a particular strain of the causative organism.
TaqMan® Probes
TaqMan® probes are dual labeled hydrolysis probes and are a registered trademark of
the Roche Molecular Systems, Inc. TaqMan® probes utilize the 5' exonuclease activity of
the enzyme Taq Polymerase for measuring the amount of target sequences in the
samples. TaqMan® probes consist of a 18-22 bp oligonucleotide probe which is labeled
with a reporter fluorophore at the 5' end and a quencher fluorophore at the 3' end.
TaqMan® Probe Functioning
While carrying out a TaqMan® experiment, a fluorogenic probe, complementary to the
target sequence is added to the PCR reaction mixture. This probe is an oligonucleotide
with a reporter dye attached to the 5' end and a quencher dye attached to the 3' end.
Till the time the probe is not hydrolized, the quencher and the fluorophore remain in
proximity to each other, separated only by the length of the probe. This proximity
however, does not completely quench the flourescenc e of the reporter dye and a
background flourescence is observed. During PCR, th e probe anneals specifically
between the forward and reverse primer to an internal region of the PCR product. The
polymerase then carries out the extension of the primer and replicates the template to
which the TaqMan® is bound. The 5' exonuclease activity of the polymerase cleaves the
probe, releasing the reporter molecule away from the close vicinity of the quencher. The
fluorescence intensity of the reporter dye, as a result increases. This process repeats in
every cycle and does not interfere with the accumulation of PCR product.
TaqMan® Probe Applications
Quantitative real time PCR.
DNA copy number measurements.
Bacterial identification assays.

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SNP genotyping.
Verification of microarray results.
FRET Probes Technology
Fluorescence Resonance Energy Transfer (FRET) occurs due to the interaction
between the electronic excited states of two dye molecules. The excitation is transferred
from one (the donor) dye molecule to the other (the acceptor) dye molecule without
emission of a photon. This is distance-dependent, that is the donor and the acceptor
dye must be in close proximity. FRET has been used for investigating a variety of
biological phenomena that produce changes in molecular proximity.
FRET probes are a pair of fluorescent probes placed in close proximity. Fluorophores
are so chosen that the emission spectrum of one ove rlaps significantly with the
excitation spectrum of the other. During FRET, the donor fluorophore excited by a light
source, transfers its energy to an acceptor fluorophore when positioned in the direct
vicinity of the former.
The acceptor fluorophore emits light of a longer wavelength, which is detected in
specific channels. The light source cannot excite the acceptor dye.
FRET Probes Functioning
The hybridization probe system consists of two oligonucleotides labeled with fluorescent
dyes. The hybridization probe pair is designed to hybridize to adjacent regions on the
target DNA. Each probe is labeled with a different marker dye. Interaction of the two
dyes can only occur when both are bound to their target. The donor probe is labeled
with fluorophore at the 3' end and the acceptor probe at 5' end. During PCR, the two
different oligonucleotides hybridize to adjacent regions of the target DNA such that the
fluorophores, which are coupled to the oligonucleotides, are in close proximity in the
hybrid structure. The donor fluorophore (F1) is excited by an external light source, then
passes part of its excitation energy to the adjacent acceptor fluorophore (F2). The
excited acceptor fluorophore (F2) emits light at a different wavelength which can then be
detected and measured.

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Applications of FRET Probes
1. Quantitative PCR.
2. DNA copy number measurements.
3. Pathogen detection assays.
4. SNP genotyping.
5. Verification of microarray results.
Scorpion Primers & Probes
Scorpion technique was developed by Dr. David Whitc ombe of DxS Ltd. Scorpion
primers are bi-functional molecules in which a primer is covalently linked to the probe.
The molecules also contain a fluorophore and a quencher. In the absence of the target,
the quencher nearly absorbs the fluorescence emitted by the fluorophore. During the
Scorpion PCR reaction, in the presence of the target, the fluorophore and the quencher
separate which leads to an increase in the fluorescence emitted. The fluorescence can
be detected and measured in the reaction tube.
Structure of the Scorpion primer
The Scorpion primer carries a Scorpion probe element at the 5' end. The probe is a self-
complementary stem sequence with a fluorophore at o ne end and a quencher at the
other. The Scorpion primer sequence is modified at the 5'end. It contains a PCR blocker
at the start of the hairpin loop (Usually HEG monomers are added as blocking agent). In
the initial PCR cycles, the primer hybridizes to the target and extension occurs due to
the action of polymerase. Scorpion primers can be used to examine and identify point
mutations by using multiple probes. Each probe can be tagged with a different
fluorophore to produce different colors.
Scorpion® Primer Functioning
After one cycle of PCR extension completes, the newly synthesized target region will be
attached to the same strand as the probe. Following the second cycle of denaturation
and annealing, the probe and the target hybridize. The denaturation of the hairpin loop

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requires less energy than the new DNA duplex produc ed. Consequently, the hairpin
sequence hybridizes to a part of the newly produced PCR product. This results in the
separation of the fluorophore from the quencher and causes emission.
Applications of Scorpion Probes
1. SNP analysis
2. Real-time PCR
3. Allelic discrimination
4. Single tube genotyping assay
Advantages and disadvantages of Real time PCR:
• Real-time reverse-transcriptase (RT) PCR quantitates the initial amount of the template
most specifically, sensitively and reproducibly, and is a preferable alternative to other
forms of quantitative RT-PCR that detect the amount of final amplified product at the
end-point.
• Real-time PCR monitors the fluorescence emitted during the reaction as an indicator of
amplicon production during each PCR cycle (ie, in real time) as opposed to the endpoint
detection.
• It is not influenced by non-specific amplification unless SYBR Green is used.
• The real-time progress of the reaction can be viewed in some systems. Real-time PCR
does not detect the size of the amplicon and thus does not allow the differentiation
between DNA and cDNA amplification.

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