The goal of modern molecular cell biology is nothing short of understanding the
biochemical, cellular, and organismal functions of all the proteins encoded in the
genome. In the preceding sections, we have discussed the isolation and analysis of
mutants, the genetic mapping of mutations, and finally the isolation and cloning of
mutation-defined genes. This approach can provide valuable information about the
molecular mechanisms underlying the cellular processes affected by the original
mutations and the in vivo functions of the normal proteins encoded by the affected
genes. As discussed in Chapter 7,
however, many genes have been identified based on the biochemical properties of
their encoded protein, the sequence similarity of the encoded protein with proteins
of known function, or their interesting patterns of expression in development. In
the absence of mutant forms of such genes, their in vivo functions may be unclear.
By mutating a specific gene in vitro and then replacing the normal copy in the
genome with a mutant form, scientists can assess its in vivo function. This
technique, referred to as gene-targeted knockout, or simply
“knockout,” is in essence the reverse of the approach described
in the previous sections. The process of isolating normal genes to be mutated will
be greatly simplified as sequencing of the genomes of several model organisms and of
the human genome progresses (Chapter
7). Whether starting from a normal protein or sequenced genome, this approach
can be summarized as follows:
Specific Sites in Cloned Genes Can Be Altered in Vitro
Specific sequences in cloned genes can be altered in vitro and then introduced
into experimental organisms. This approach has been exploited primarily to study
two questions. First, what is the relationship between the structure of a
particular protein and its biological function? And second, what are the
specific DNA sequences required to determine the expression pattern of a
particular gene?
Figure 8-29
.
In vitro mutagenesis of cloned genes with chemically
synthesized oligonucleotides
In this example, a cloned gene that normally contains three
segments (A, B, and C) is mutagenized by deletion of segment B.
An oligonucleotide consisting of A′ and C′
segments, which are complementary to A and C, is chemically
synthesized and then is hybridized to the complementary
single-stranded DNA containing the gene to be mutagenized. In
the hybrid molecule, segment B forms a loop. The
A′-C′ oligonucleotide serves as a primer for
synthesis of a complementary strand; one of the nucleotide
triphosphates contains sulfur in place of one of the oxygens at
the α position. The heteroduplex DNA is treated with
the restriction endonuclease NciI; because this
enzyme cannot cleave the sulfur-substituted strand (green), a
single-strand nick is created in the original template strand
(purple). The nicked molecule then is treated with exonuclease
III, which cleaves nucleotides in the 3′ to
5′ direction, thereby removing the B segment. The
resulting gap in the template strand is filled in by DNA
polymerase and closed by DNA ligase.
A variety of enzymatic and chemical methods are available for producing
site-specific
mutations in vitro. In recent years, however, the most common
methods use specific oligonucleotides as
mutagens. Because oligonucleotides of
any desired sequence can be chemically synthesized (see
Figure 7-9), oligonucleotide-based mutagenesis can
generate precisely designed deletions, insertions, and
point mutations in a DNA
sequence. illustrates the
use of this strategy to produce a deletion.
DNA Is Transferred into Eukaryotic Cells in Various Ways
Production of both knockout and transgenic organisms requires the transfer of DNA
into eukaryotic cells. Many types of cells can take up DNA from the medium.
Yeast cells, for instance, can be treated with enzymes to remove their thick
outer walls; the resulting spheroplasts will take up DNA added
to the medium. Plant cells also can be converted to spheroplasts, which will
take up DNA from the medium. Cultured mammalian cells take up DNA directly,
particularly if it is first converted to a fine precipitate by treatment with
calcium ions. Another popular method for introducing DNA into yeast, plant, and
animal cells is called electroporation. Cells subjected to a
brief electric shock of several thousand volts become transiently permeable to
DNA. Presumably the shock briefly opens holes in the cell membrane allowing the
DNA to enter the cells before the holes reseal. DNA also can be injected
directly into the nuclei of both cultured cells and developing embryos.
Once the foreign DNA is inside the host cell, enzymes that probably function
normally in DNA repair and recombination join the fragments of foreign DNA with
the host cell’s chromosomes. Since only a relatively small fraction of
cells take up DNA, a selective technique must be available to identify the
transgenic cells. In most cases the exogenous DNA includes a gene encoding a
selectable marker such as drug resistance. The introduced DNA can insert into
the host genome in a highly variable fashion showing no site specificity, can
replace an endogenous gene by homologous recombination, or can remain as an
independent extrachromosomal DNA molecule referred to as an
episome.
Normal Genes Can Be Replaced with Mutant Alleles in Yeast and Mice
Gene knockout is a technique for selectively inactivating a gene by replacing it
with a mutant allele in an otherwise normal organism. This technique of
disrupting gene function, which has been widely used in yeast and mice, is a
powerful tool for unraveling the mechanisms by which basic cellular processes
occur.
Gene Knockout in Yeast
Figure 8-30
.
Replacement of the normal HIS3 gene (blue)
by homologous recombination with a mutant his3
gene (yellow) in the yeast S.
cerevisiae.
Recombinant DNA technology is used to prepare a plasmid
containing his3, a deletion mutant that encodes
only part of the sequence of one enzyme in the histidine
biosynthetic pathway, and URA3, which encodes
an enzyme in the uracil biosynthetic pathway. The
URA3 gene is included simply as a
selectable marker. The recipient cell (a uracil-requiring
strain) takes up the plasmid, which can integrate into the
ura3 gene (not of interest in this case) or
into the HIS3 gene. Recombinant cells are
selected by their ability to grow in the absence of uracil.
Subsequent intrachromosomal recombination yields cells that have
lost URA3 and retain either the wild-type
HIS3 gene or the his3
mutant. Cells carrying the his3 mutant gene are
detected by replica plating in the presence and absence of
histidine. [See S. Scherer and R. W. Davis, 1979, Proc.
Nat’l. Acad. Sci. USA
76:4951.]
Figure 8-31
.
Demonstration that actin gene is required for yeast viability
by gene-targeted knockout
A recombinant plasmid containing the URA3 gene,
a selectable marker, and a mutant actin gene (yellow) is
introduced into uracil-requiring (Ura3−)
diploid yeast cells. (The ura3 gene, which is
located on a different chromosome, is not depicted.) Cells that
integrate the plasmid are selected by their growth in the
absence of uracil. Since the recombinant cells contain one
normal and one disrupted actin gene, they can synthesize actin.
To determine whether the actin gene is essential, meiosis and
sporulation are induced by starving the cells; each diploid cell
produces four haploid spores (see Figure 10-54). The wild-type spores,
which are Ura3−, can grow on uracil, but
those with the disrupted actin gene do not. [Adapted from D.
Shortle et al., 1982, Science
217:371.]
After foreign, or exogenous, yeast DNA is taken up by
diploid yeast cells,
recombination generally occurs between the introduced DNA and the homologous
chromosomal site in the recipient cell. Because of this specific, targeted
recombination of identical stretches of DNA, called
homologous
recombination, any
gene in yeast
chromosomes can be replaced
with a mutant
allele ().
The resulting
heterozygous yeast cells, carrying one mutant
allele and one
wild-type
allele, generally grow normally. To determine whether the
knocked-out
gene controls an obligatory function, recombinants containing
the mutant
allele on one
chromosome are treated to induce
meiosis and
sporulation; each
diploid cell produces four
haploid spores, which are
tested for viability. One of the first
genes tested in this way was the one
encoding
actin, a prominent cytoskeletal
protein in yeast and higher
organisms.
Haploid yeast spores without a normal
actin gene cannot grow
().
This technique also is useful in assessing the role of proteins identified
solely on the basis of DNA sequence. For instance, the entire sequence of
the S. cerevisiae genome has been determined. As described
in Chapter 7, analysis of
genomic sequences can identify stretches of DNA that exhibit long open
reading frames or homology to genes encoding known proteins; such stretches
are likely to be transcribed and translated into as yet unidentified
proteins (see Figure 7-31). Gene
knockouts can be used to determine whether such regions are important for
specific cellular functions that are phenotypically detectable. This
technique thus provides a powerful approach to identifying and studying new
genes and the proteins that they encode.
This gene-knockout approach already has been used to analyze yeast chromosome
III. Analysis of the DNA sequence indicated that this chromosome contains
182 open reading frames of sufficient length to encode proteins longer than
100 amino acids, which is assumed in this analysis to be the minimum length
of a naturally occurring protein. The sequences of the proteins that could
be encoded by 116 of these putative protein-coding regions exhibited no
obvious homology to any known proteins. Gene knockout of 55 of these regions
showed that 3 were required for viability; further analysis of 42
nonessential genes revealed that 14 showed a mutant phenotype and 28 did
not. The large number of putative genes with no detectable mutant phenotype
is quite surprising. In some cases the lack of a phenotype could indicate
the existence of backup or compensatory pathways in the cell. Alternatively,
the mutations may give rise to subtle defects that would require more
in-depth phenotypic analysis to uncover.
Gene Knockout in Mice
Gene-targeted knockout mice are a powerful experimental system for studying
development, behavior, and physiology; they also may be useful model systems
for studying certain human genetic diseases. The procedure for producing
gene-targeted knockout mice involves the following steps:
- 1
Mutant alleles are introduced by homologous recombination into
embryonic stem (ES) cells.
- 2
ES cells containing a knockout mutation in one allele of the gene
being studied are introduced into early mouse embryos. The resultant
mice will be chimeras
containing tissues derived from both the transplanted ES cells and
the host cells. These cells can contribute to both the germ-cell and
somatic-cell populations.
- 3
Chimeric mice are mated to assess whether the mutation is
incorporated into the germ line.
- 4
Mice each heterozygous for the knockout mutation are mated to produce
homozygous knockout mice.
Figure 8-32
.
Preparation of embryonic stem (ES) cells
Fertilized mouse eggs divide slowly at first; after 41/2 days,
they form the blastocyst, a hollow structure composed of about
100 cells surrounding an inner cavity called the
blastocoel. Only ES cells, which constitute
the inner cell mass, actually form the embryo. Other cells form
the trophectoderm, which gives rise to the membranes (amnion and
placenta) by which the embryo is attached to the uterine wall.
Embryonic stem cells can be removed from the blastocyst and
grown on lethally irradiated “feeder cells.”
[See E. Robertson et al., 1986, Nature
323:445.]
The isolation and culture of embryonic
stem cells, which are derived from the
blastocyst, are illustrated in . These cells can be grown in culture through many
generations. Exogenous DNA containing a mutant
allele of the
gene being
studied is introduced into ES cells by
transfection. The introduced DNA
recombines with chromosomal sequences in about 1 cell out of 100 (i.e., 1
percent
recombination frequency). In some cells, the added DNA recombines
with the homologous chromosomal site, but
recombination at other chromosomal
sites (i.e., nonhomologous
recombination) occurs
10
3 – 10
4 times
more frequently. The small fraction of cells in which homologous
recombination takes place can be identified by a combination of positive and
negative selection: positive selection to identify cells in which any
recombination occurs and negative selection to remove cells in which
recombination takes place at nonhomologous sites.
Figure 8-33
.
Isolation of mouse ES cells with a gene-targeted disruption
by positive and negative selection
(a) When exogenous DNA is introduced into ES cells, random
insertion via nonhomologous recombination occurs much more
frequently than gene-targeted insertion via homologous
recombination. Recombinant cells in which one copy of the gene
X (orange) is disrupted can be obtained by
using a recombinant vector that carries gene X
disrupted with neor (light red), a
neomycin-resistance gene, and, outside the region of homology,
tkHSV (purple), the thymidine
kinase gene from herpes simplex virus. The viral thymidine
kinase, unlike the endogenous mouse enzyme, can convert the
nucleotide analog ganciclovir into the monophosphate form; this
is then modified to the triphosphate form, which inhibits
cellular DNA replication in ES cells. Thus ganciclovir is
cytotoxic for recombinant ES cells carrying the
tkHSV gene. Nonhomologous
insertion includes the tkHSV gene,
whereas homologous insertion doesn’t; therefore, only
cells with nonhomologous insertion are sensitive to ganciclovir.
(b) Recombinant cells are selected by treatment with neomycin,
since cells that fail to pick up DNA or integrate it into their
genome are neomycin-sensitive. The surviving recombinant cells
are treated with ganciclovir. Only cells with a targeted
disruption in gene X, and therefore lacking the
tkHSV gene, will survive. [See
S. L. Mansour et al., 1988, Nature
336:348.]
For this selection scheme to work, the DNA constructs introduced into ES
cells need to include, in addition to sequences used to knock out the
gene
of interest, two selectable marker
genes (). One of these additional
genes
(neor) confers neomycin
resistance; it permits positive selection of cells in which either
homologous (specific) or nonhomologous (random)
recombination has occurred.
The second selective
gene, the thymidine
kinase gene from herpes simplex
virus (tkHSV) confers
sensitivity to ganciclovir, a cytotoxic
nucleotide analog; this
gene permits
negative selection of ES cells in which nonhomologous
recombination has
occurred. Only ES cells that undergo homologous
recombination (i.e.,
gene-targeted specific insertion of the DNA construct) can survive this
selection scheme.
Figure 8-34
.
General procedure for producing gene-targeted knockout
mice
Embryonic stem (ES) cells heterozygous for a knockout mutation in
a gene of interest (X) and homozygous for a
marker gene (here, black coat color) are transplanted into the
blastocoel cavity of 4.5-day embryos that are homozygous for an
alternate marker (here, white coat color). The early embryos
then are implanted into a pseudopregnant female. Some of the
resulting progeny are chimeras, indicated by their black and
white coats. Chimeric mice then are backcrossed to white mice;
black progeny from this mating have ES-derived cells in their
germ line. By isolating DNA from a small amount of tail tissue,
it is possible to identify black mice heterozygous for the
knockout allele. Intercrossing of these black mice produces
individuals homozygous for the disrupted allele, that is,
knockout mice. [Adapted from M. R. Capecchi, 1989,
Trends Genet.
5:70.]
Once ES cells
heterozygous for a knockout
mutation in the
gene of interest
are obtained, they are injected into a recipient mouse blastocyst, which
subsequently is transferred into a surrogate pseudopregnant mouse (). If the ES cells also are
homozygous for a visible marker trait (e.g., coat color), then chimeric
progeny carrying the knockout
mutation can be identified easily. These are
then mated with mice
homozygous for another
allele of the marker trait to
determine if the knockout
mutation is incorporated into the
germ line.
Finally, mating mice, each
heterozygous for the knockout
allele, will
produce progeny
homozygous for the knockout
mutation.
Cell-Type-Specific Gene Knockout in Mice
In most cases, investigators are interested in examining the effects of
knockout mutations in a particular region of the mouse, at a specific stage
in development, or both. Since most genes function in different parts of the
organism and at different times, a knockout mouse may die or have defects in
various tissues prior to the stage to be analyzed. To address this problem,
mouse geneticists have devised a clever technique using site-specific DNA
recombination sites (called loxP sites) and the enzyme,
called Cre, that catalyzes recombination between them. The
loxP-Cre recombination system is present in bacteriophage P1, but also
promotes recombination when placed in mouse cells.
Figure 8-35
.
Cell-type-specific gene knockouts using the loxP-Cre
recombination system
Two loxP sites are inserted on each side of an essential exon (2)
of the gene of interest (i.e., gene X) (blue) by homologous
recombination. These sites do not disrupt gene function. The
loxP-containing mouse is crossed to a transgenic mouse carrying
a cell-type-specific promoter controlling expression of the Cre
recombinase, which induces recombination between loxP sites.
This mouse is heterozygous for a constitutive gene
X knockout. In the resulting loxP-Cre
mouse, Cre protein is produced only in those cells in which the
promoter is active, and in those cells recombination therefore
occurs between the loxP sites, leading to deletion of exon 2.
Since the other allele is a constitutive gene X
knockout, deletion between the IoxP sites results in complete
loss of function in all cells expressing Cre.
Homologous
recombination strategies discussed in the previous section are
used to obtain mice in which loxP sites are inserted so they flank the
gene
of interest or an essential
exon. Since the inserted loxP sites are not
within
exons, they do not by themselves disrupt
gene function.
Transgenic
mice also are prepared carrying the
cre gene linked to a
cell-type-specific
promoter. As depicted in , mating of these two types of mice will yield
progeny that carry the
gene of interest modified by insertion of flanking
lox P sites and the
cre gene controlled by a
cell-type-specific
promoter. In these mice,
recombination between the loxP
sites, which disrupts the
gene of interest, will occur only in those cells
in which the
promoter is active and therefore producing the Cre
protein
necessary to induce the
recombination.
One important example of this technique comes from studies on learning and
memory. Earlier pharmacological and physiological studies had indicated that
normal learning requires a specific neurotransmitter receptor, the NMDA
class of glutamate receptors, in a specific region of the brain called the
hippocampus. But mice in which the gene encoding an NMDA receptor subunit
was knocked out died neonatally, precluding analysis of the
receptor’s role in learning. Cell-type-specific inactivation of
the receptor was achieved by constructing mice carrying a Cre gene expressed
in a subclass of hippocampal neurons and two different alleles of the
receptor subunit gene, an allelle containing the loxP sites and a
conventional knockout allele. These mice survived to adulthood and showed
learning and memory defects, confirming a role for these receptors in normal
learning and memory.
Use of Knockout Mice to Study Human Genetic Diseases
Gene knockout can produce model systems for studying inherited human
diseases. Such model systems are powerful tools for investigating the nature
of genetic diseases and the efficacy of different types of treatment, and
for developing effective gene therapies to cure these often devastating
diseases.
Recent studies on cystic fibrosis illustrate this use of the knockout
technique. Cystic fibrosis, which afflicts about 1 in 2000 Caucasians, is
caused by an autosomal recessive mutation in the CFTR gene
(see Figure 8-17b). This gene was
cloned by positional cloning strategies, and the biochemical function of its
encoded protein studied. Using the human gene, researchers isolated the
homologous mouse gene and subsequently introduced mutations in it. The
gene-knockout technique was then used to produce homozygous mutant mice,
which showed symptoms (i.e., a phenotype) similar to those of humans with
cystic fibrosis. These knockout mice are currently being used as a model
system for studying this genetic disease and developing effective
therapies.
Foreign Genes Can Be Introduced into Plants and Animals
In the previous section we discussed techniques for replacing one form of a gene
with another through homologous recombination. In this section we discuss
methods for producing transgenic organisms, which carry cloned genes that have
integrated randomly into the host genome.
Transgenic technology has numerous experimental applications and potential
agricultural and therapeutic value. For instance, dominantly acting alleles of
tumor-causing genes can be used to produce transgenic mice, thus providing an
animal model for studying cancer. In Drosophila, transgenes
often are used to determine whether a cloned segment of DNA corresponds to a
gene defined by mutation. If the cloned DNA is indeed the gene in question, then
introducing it as a transgene into a mutant fly will transform the mutant into a
phenotypically normal individual. Transgenic plants may be commercially valuable
in agriculture. Plant scientists, for example, have developed transgenic
tomatoes that exhibit reduced production of ethylene, which promotes fruit
ripening. The ripening process is delayed in these transgenic tomatoes, thus
prolonging their shelf life. Finally, transgenic technology is a critical
component in the burgeoning field of gene therapy for human genetic
diseases.
Transgenic Mice
As noted in the discussion of knockout mice, specific integration of
exogenous DNA into the genome of mouse cells by homologous recombination
occurs at a very low frequency. In contrast, the frequency of random
integration of exogenous DNA into the mouse genome at nonhomologous sites is
very high. Because of this phenomenon, the production of transgenic mice is
a highly efficient and straightforward process.
Figure 8-36
.
General procedure for producing transgenic mice
[See R. L. Brinster et al., 1981, Cell
27:223.]
As outlined in , foreign
DNA containing a
gene of interest is injected into one of the two pronuclei
(the male and female
haploid nuclei contributed by the parents) of a
fertilized mouse egg before they fuse. The injected DNA has a good
likelihood of being randomly integrated into the
chromosomes of the
diploid
zygote. Injected eggs then are transferred to foster mothers in which normal
cell growth and
differentiation occurs. About
10 – 30 percent of the progeny will contain
the foreign DNA in equal amounts (up to 100 copies per cell) in all tissues,
including
germ cells. Immediate breeding and backcrossing (parent-offspring
mating) of the 10 – 20 percent of these mice
that breed normally can produce pure
transgenic strains
homozygous for the
transgene.
Numerous examples of the use of
transgenic mice for studying various aspects of normal mammalian biology are
presented in other chapters. They also provide a model system for studying
disease processes. For example, many forms of cancer are promoted by normal
cellular genes acting in a dominant fashion owing to their misregulated
activity. Although transgenic mice carrying one of these genes, called
myc, develop normally, tumors form at a high frequency.
The observation that only a small number of cells expressing the transgene
develop tumors supports a model in which additional genetic changes are
necessary for tumors to form. These mice may provide an important tool for
identifying those changes.
Transgenic Fruit Flies
Figure 8-37
.
Generation of transgenic fruit flies by P-element
transformation
The P element, a mobile genetic element, can move from one place
in the genome to another. This movement (transposition) is
catalyzed by transposase, which is encoded by the P element; the
3′ and 5′ ends of the P element are
recognized by transposase and are required for transposition to
occur. To produce transgenic fruit flies by this method, the
functionally different regions of the P element are incorporated
into two different bacterial plasmids. The donor plasmid
contains three necessary elements: the transgene (orange); a
marker gene (green) used to indicate flies in which the plasmid
DNA is transposed to a recipient chromosome; and both ends of
the P element (dark
purple) — 3′ P and
5′ P — flanking the
other two genes. It does not contain transposase. In this
example, the marker is the dominant
w+ allele, which
confers red eye color. The red bracket indicates the segment of
the donor plasmid that can transpose into the fly genome. The
other plasmid carries the P element (encoding transposase) with
mutations in one end, which prevent it from transposing. The two
plasmids are co-injected into blastoderm embryos homozygous for
the recessive w− allele,
which confers white eye color. Transposase synthesized from the
gene on the P-element plasmid catalyzes transposition of the
donor plasmid DNA into the fly genome. Because transposition
occurs only in germ-line cells (not in somatic cells), all the
G0 adults that develop from injected embryos have
white eyes. Mating of these flies with white-eyed flies will
yield some G1 red-eyed progeny carrying the transgene
and the marker allele
(w+)
in all cells.
Foreign DNA can be incorporated into the
Drosophila
germ-line
genome by the technique of P-element
transformation (). This technique makes use
of a segment of the P element, a highly
mobile DNA element, which can
transpose (jump) from an extrachromosomal element into a
chromosome. (Mobile
DNA elements are discussed in detail in
Chapter 9.) Generally, this procedure results in
incorporation of a single copy of the
transgene into the
Drosophila genome. In contrast,
transgenic mice carry
multiple copies of the
transgene incorporated into their
chromosomes. In
both organisms, however, the chromosomal insertion site is highly
variable.
Flies that develop from injected embryos will carry some germ cells that have
incorporated the transgene; some of their progeny will carry the transgene
in all somatic and germ-line cells, giving rise to pure transgenic lines.
Individuals carrying the transgene are recognized by expression of a marker
gene (e.g., one affecting eye color) that is also present on the donor DNA.
Although the transgenes in Drosophila and mice insert in
chromosomal sites different from the position of the corresponding
endogenous gene, they usually are expressed in the right tissue and at the
right time during development. Examples of the importance of this technology
for studying development are discussed in Chapter 14.
Transgenic Plants
Figure 8-38
.
Production of transgenic plants with recombinant Ti
plasmids
In nature, the Ti (tumor-inducing) plasmid in A.
tumefaciens gains entry into a plant and is
integrated into the plant DNA owing to the action of the
VIR (virulent) region of the plasmid.
Tumor-like growths (galls) result from action of the T region of
the gene. By recombinant DNA techniques, a foreign gene (orange)
is introduced within the T region of a Ti plasmid, thus
destroying its tumor-inducing ability. An agrobacterium
containing such a recombinant Ti plasmid then is used to
introduce the foreign gene into plant cells. When a selectable
transgene is used — here one
conferring resistance to herbicide
(herbres) — recombinant
plants can be selected. [See R. T. Fraley et al., 1983,
Proc. Nat’l. Acad. Sci. USA
80:4803.]
In nature, plant cells often live in close association with certain bacteria,
which may provide a convenient vehicle for introducing cloned DNA into
plants.
Agrobacterium tumefaciens, for example,
attaches to the cells of dicotyledonous plants and causes the formation of
plant
tumors known as
galls. (Plants with two leaflets from
each seed are called
dicotyledons, or
dicots; plants with one leaflet are called
monocots .) This bacterium introduces a circular DNA molecule,
called the
Ti (
tumor-inducing)
plasmid,
into the plant cell in a manner similar to bacterial conjugation. The
plasmid DNA then recombines with the plant DNA. Since the Ti
plasmid has
been isolated, new
genes can be inserted into it using
recombinant DNA
techniques and the Ti
genes causing
tumors can be disrupted. The resulting
recombinant
plasmid can then transfer desired
genes into plant cells ().
An especially useful characteristic of plants for transgenic studies is the
ability of cultured plant cells to give rise to mature plants. Meristematic
(growing) cells from dissected plant tissue or cells within excised parts of
a plant will grow in culture to form callus tissue, an
undifferentiated lump of cells. Under the influence of plant growth
hormones, different plant parts (roots, stems, and leaves) develop from the
callus and eventually grow into whole, fertile plants. When an agrobacterium
containing a recombinant Ti plasmid infects a cultured plant cell, the newly
incorporated foreign gene is carried into the plant genome.
As noted above A. tumefaciens readily infects dicots
(petunia, tobacco, carrot) but not monocots; reliable techniques for
introducing genes into monocots are still being developed. Direct
introduction of DNA by electroporation has been successful in rice plants,
which are monocots, and the future looks bright for the manipulation of
other commercially important monocotyledonous crop plants. Also available
for gene-transfer experiments are cells of a tiny, rapidly growing member of
the mustard family called Arabidopsis thaliana. This plant
is well-suited to genetic analysis of a variety of developmental and
physiological processes. It takes up little space, is easy to grow, and has
a small genome, and genes defined by mutations can be cloned by positional
cloning strategies.
