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Annu. Rev. Plant Biol. 2004. 55:289–313
doi: 10.1146/annurev.arplant.55.031903.141633
c 2004 by Annual Reviews. All rights reserved
Copyright First published online as a Review in Advance on January 7, 2004
PLASTID TRANSFORMATION IN HIGHER PLANTS
Pal Maliga
Waksman Institute, Rutgers University, Piscataway, New Jersey 08854-8020;
Department of Plant Biology, Rutgers University, New Brunswick, New Jersey 08901;
email: maliga@waksman.rutgers.edu
Key Words plastid genetics, plastid markers, protein expression, gene knockouts,
gene containment
■ Abstract Plastids of higher plants are semi-autonomous organelles with a small,
highly polyploid genome and their own transcription-translation machinery. This review provides an overview of the technology for the genetic modification of the plastid
genome including: vectors, marker genes and gene design, the use of gene knockouts and over-expression to probe plastid function and the application of site-specific
recombinases for excision of target DNA. Examples for applications in basic science
include the study of plastid gene transcription, mRNA editing, photosynthesis and evolution. Examples for biotechnological applications are incorporation of transgenes in
the plastid genome for containment and high-level expression of recombinant proteins
for pharmaceutical and industrial applications. Plastid transformation is routine only
in tobacco. Progress in implementing the technology in other crops is discussed.
CONTENTS
INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
SOMATIC CELL GENETICS OF THE PLASTID . . . . . . . . . . . . . . . . . . . . . . . . . . .
APPROACHES TO PLASTID TRANSFORMATION . . . . . . . . . . . . . . . . . . . . . . . .
Stable Transformation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Episomal Maintenance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Transient Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
DNA DELIVERY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
GENETIC MARKERS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Primary Positive Selection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Secondary Positive Selection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Negative Selection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Reporter Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Combination of Visual and Selective Markers . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
GENE DESIGN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
PROBLEMS WITH DATA INTERPRETATION . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
IMPLEMENTATION IN DIFFERENT SPECIES . . . . . . . . . . . . . . . . . . . . . . . . . . . .
APPLICATIONS IN BASIC SCIENCE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Gene Knockouts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Overexpression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Transcription . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
RNA Editing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Evolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
APPLICATIONS IN BIOTECHNOLOGY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Agronomic Traits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Expression of Recombinant Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Marker Gene Elimination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Containment by Plastid Localization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
PERSPECTIVES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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INTRODUCTION
Plastids are plant cellular organelles with their own genome and transcriptiontranslation machinery. The plastid genome (plastome or ptDNA) is a highly polyploid, circular double-stranded DNA 120 kb to 180 kb in size, encoding ∼120
genes. A salient feature of the plastid genome in most higher plant species is
duplication of a large (∼25 kb) region in an inverted orientation (112, 148, 160).
Plastid is the general organelle category encompassing proplastids, the progenitors
of all plastid types and chloroplasts (green plastids), chromoplasts (yellow or red,
in some fruits and flowers), and different types of white plastids such as the amyloplasts (starch containing) and elaioplasts (oil containing) (48). Plastid functions
include photosynthesis and starch, amino acid, lipid, and pigment biosynthesis.
Most plastid proteins are encoded in 2100–3600 nuclear genes, the products of
which are translated on cytoplasmic ribosomes and imported into plastids (89).
Boynton, Gillham, and colleagues (18) first achieved plastid transformation
in 1988 in a unicellular alga, Chlamydomonas reindhartii, followed in 1990 by
transformation of the plastid genome in tobacco, which occurred in my laboratory
(151). Plants with transformed plastid genomes are termed transplastomic (96).
This review focuses on the methodology of plastid transformation in higher plants
and selected applications in basic science and biotechnology. General and specialized reviews on this topic have been published elsewhere (3, 12, 14, 59, 98, 100,
138).
SOMATIC CELL GENETICS OF THE PLASTID
Each particular type of plastid carries identical ptDNA copies, which are attached
to membranes (77, 124, 125) in clusters called plastid nucleoids (86). The number
of plastids and ptDNA copies per cell is highly variable depending on the cell type
(9). In tobacco, the meristematic cells contain 10–14 proplastids, each containing
1–2 nucleoids per organelle, whereas leaf cells may contain 100 chloroplasts, with
10–14 nucleoids each, ∼10,000 ptDNA per cell (9, 157). Plastid transformation
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involves transforming one or a few ptDNA copies, followed by gradually diluting
plastids carrying nontransformed copies on a selective medium. Although there
may be as many as 10,000 ptDNA copies in a cell, physical contact and recombination are probably feasible only within a nucleoid. This suggests that the genetic
unit of transformation and sorting is probably the nucleoid (Figure 1A).
A study of plastid sorting in heteroplastomic tobacco cells shows preferential
propagation of plastids with antibiotic resistance markers on a selective medium
(106). The plastid markers were streptomycin and lincomycin resistance; the heteroplastomic cells were obtained by protoplast fusion. Cell lines established in the
presence of streptomycin or lincomycin had only one parental plastid type with the
appropriate antibiotic resistance marker; in the absence of antibiotic selection both
plastid types were maintained. Reaching the homoplastomic state was estimated
to take ∼20 cell divisions.
Current protocols for plastid transformation employ strategies to obtain homoplastomic plants by segregating genome copies and organelles in somatic cells.
The most common approach to plastid transformation in tobacco is transformation
of chloroplasts in leaves and regeneration of shoots from the transformed cells
on a selective medium (151, 153). Formation of homoplastomic cells is accelerated by chloroplast to proplastid dedifferentiation, with a concomitant reduction
in nucleoid (ptDNA) number in tissue culture cells (157), then a rebuilding of the
organelle and nucleoid (ptDNA) numbers in regenerated plants (Figure 1B).
Transplastomic shoots regenerated from leaves after bombardment are always
chimeras. Spectinomycin and kanamycin resistance, conferred by the expression
of chimeric genes, is not cell autonomous in regenerating shoots or in seedling
cotyledons. [An exception is rrn16-based spectinomycin resistance in seedling
cotyledons (151).] Lack of cell autonomous expression means that, in chimeric
shoots, nontransformed sectors also have a resistant (green) phenotype (Figure 2A,
see color insert), although they become bleached when cut out and placed in direct contact with the selective medium (see Genetic Markers, below). Resistant
phenotype of nontransformed cells in a chimeric plant is due to cross-protection
by transformed cells. However, transformed and nontransformed sectors can be
readily identified by color (green or white) in knockout plants lacking a photosynthetic gene (2) or by green fluorescent protein (GFP) accumulation (Figure 2B)
(72), which are cell autonomous traits.
The preferred method to obtain homoplastomic tobacco plants is regenerating
new shoots from the transplastomic sectors, which are then rooted (151, 153).
Homoplastomic plants from the chimeric shoots can also be obtained in the seed
progeny, as long as the transplastomes are present in the cell layer that contributes
to the maternal germline (2). In dicots this is the L2 layer, the phenotype of
which is visible at the leaf margins (113). Homoplastomic plants can be obtained
directly from tissue culture cells if cells (protoplasts) are first cultured to form
undifferentiated callus, and plant regeneration is delayed until plastid segregation
is complete. However, extended propagation of cells in tissue culture is undesirable
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because it causes chromosome rearrangements and polyploidization that affect
plant fertility.
APPROACHES TO PLASTID TRANSFORMATION
Two approaches have been applied to stable genetic modification of plastids: integration of transforming DNA by homologous targeting and introduction of independently replicating shuttle vectors. There have also been attempts to study
gene expression in plastids from transiently introduced DNA. Thus far only stable
integration of the transforming DNA yielded satisfactory results.
Stable Transformation
Plastid transformation vectors are E. coli plasmid derivatives with cloned ptDNA
sequences (1–2 kb) that flank both sides of a selectable marker gene and cloning
sites. The ptDNA sequences serve as targeting regions to direct integration into the
plastid genome (Figure 3A). The plastid vector is propagated in E. coli, and then
introduced into plastids where the marker gene and the gene of interest integrate in
the targeted region by two homologous recombination events. The E. coli vector
part does not carry a plastid replication origin and is subsequently lost.
Plastid targeting sequences do not have special properties and may derive from
any part of the plastid genome. Inserting a marker gene did not interfere with
expression of flanking plastid genes at 14 intergenic regions listed in Table 1.
(An exception is the petB-petD intergenic region, where insertion of aadA made
the plants dependent on sucrose) (107). Plastid DNA fragments containing the
14 neutral intergenic insertion sites could potentially be developed into plastid
Figure 3 Plastid transformation to obtain marker-free tansplastomic plants. (A) Plastid
vectors target insertion of marker gene (m) and gene of interest (goi) via the left (L) and right
(R) targeting sequences (153). Two loxP sites (open triangles) flank the marker gene (m is
floxed) in T1-ptDNA to facilitate its removal by CRE, the P1 phage site-specific recombinase
(26, 56). (B) Plastid-targeted CRE is expressed from a nuclear gene and excises floxed marker
gene to yield T2 marker-free transplastome.
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TABLE 1 Intergenic regions where transgenes were inserted in the plastid genome
Insertion site
Species
References
trnH/pbA
N. tabacum
(21)
trnG/trnfM
N. tabacum
L. esculentum
(17)
(121)
ycf3/trnS
N. tabacum
(64, 74)
rbcL/accD
N. tabacum
(153)
petA/psbJ
N. tabacum
(15, 64, 74)
50 rps12/clpP
N. tabacum
(85, 132)
petD/rpoA
N. tabacum
(74, 149)
ndhB/rps7
B. napus
(62)
3 rps12/trnV
N. tabacum
A. thaliana
O. sativa
L. fendleri
(140, 173)
(134)
(72)
(135)
trnV/rrn16
N. tabacum
(140, 141)
rrn16/trnI
N. tabacum
(140, 151)
trnI/trnA
N. tabacum
(33, 109, 140)
trnN/trnR
N. tabacum
(64, 172)
rpl32/trnL
N. tabacum
(43, 80, 158)
0
vectors. So far only two of these, the trnV/30 rps12 (173) and trnG/trnfM (121)
targeting regions, were made convenient for cloning by removing unnecessary
restriction sites and providing a cluster of unique cloning sites.
Episomal Maintenance
Plastid replication origins can be incorporated in transformation vectors for episomal maintenance. Such plasmids are termed shuttle vectors because they are
maintained as plasmids in E. coli by replication using the ColE1 replication origin
and as extrachromosomal elements in plastids using a plastid ori sequence. An
opportunity to develop such vectors was provided by the serendipitous discovery
of a small (868 bp; NICE1), breakaway circular ptDNA fragment derived from a
vector, which was transmitted through seed (142). A shuttle vector was constructed
by incorporating NICE1 sequences in an E. coli plasmid and a plastid selectable
marker. The shuttle vectors were present in both extrachromosomal and integrated
form, and were used to demonstrate gene conversion between vector and ptDNA
sequences (144). The shuttle vectors were not practical because they were rapidly
lost in the absence of selection.
A conceptual design for plastid shuttle vectors utilizing chloramphenicol resistance as a selective marker gene has been described (32). No published data confirm
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plastid transformation with the vectors, or recovery of transplastomic clones by
selection for chloramphenicol resistance.
Transient Expression
Expression of transiently introduced DNA was also studied in plastids. β-glucuronidase (GUS) expression was shown in isolated chloroplasts after polyethylene
glycol (PEG) treatment introduced the transforming DNA into protoplasts (136).
Evidence for transient green fluorescent protein (GFP) expression in plastids was
reported after biolistic DNA delivery (60) or microinjection of plastid reporter
genes (76).
During the early 1990s several papers were published on transient expression
of chloramphenicol acetyltransferase and GUS from plastid gene constructs after
biolistic delivery of chimeric genes to cells (for example, 37, 168). Because activity
derived from plastid and nuclear expression was not distinguished, the conclusions
of these papers are now considered obsolete.
DNA DELIVERY
A critical development for the progress of organelle biology was the biolistic DNA
delivery enabling transformation of Chlamydomonas chloroplasts (18), yeast mitochondria (67), and higher plant chloroplasts (151) pioneered by John Sanford
and colleagues during the late 1980s. The original gun with particles accelerated
by a gunpowder charge (75) was quickly replaced by a cleaner version, using highpressure He gas as propellant (66). The particle gun remains unchanged since the
early 1990s, except that an adaptor was introduced to simultaneously accommodate seven macrocarriers (Hepta adaptor; Biorad, Hercules, CA). Tungsten or gold
particles work equally well. Biolistic delivery is the system of choice for most
laboratories, as manipulation of leaves, cotyledons, or cultured cells in tissue culture requires less experience than the alternative PEG treatment of protoplasts.
Protocols for DNA coating of particles (97), selection in tobacco leaf cultures
(153), and transformation of chloroplasts in leaves (10) have been published.
PEG-mediated transformation of plastids requires enzymatically removing the
cell wall to obtain protoplasts, then exposing the protoplasts to purified DNA in
the presence of PEG. The protoplasts first shrink in the presence of PEG, then lyse
due to disintegration of the cell membrane. Removing PEG before the membrane
is irreversibly damaged reverses the process. PEG treatment was first used to test
transient expression of GUS in chloroplasts (136), then for stable transformation
of the plastid genome (49, 111). A review of the method (80) and a step-by-step
protocol of plastid transformation by PEG treatment are available (79).
Additional approaches for DNA delivery to plastids have also been tried. The
first attempt at plastid transformation used an Agrobacterium binary transformation vector (38). We now understand that sophisticated nuclear targeting of the
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transferred DNA makes use of Agrobacterium for plastid transformation a challenge. Microinjection, which looks promising for transient gene expression (76),
has not yet yielded stable transplastomic clones.
GENETIC MARKERS
Primary Positive Selection
Primary markers are suitable for selectively amplifying a small number of transformed ptDNA copies. Currently known primary markers are resistance to spectinomycin, streptomycin, and kanamycin, which inhibit protein synthesis on
prokaryotic-type plastid ribosomes. These antibiotics inhibit greening, cell division, and shoot formation in tobacco culture. Therefore, greening, faster proliferation, and shoot formation were used to identify transplastomic clones on a selective
medium. The first transplastomic clones were obtained by spectinomycin selection.
Because spectinomycin allows slow proliferation of nontransformed tobacco cells
it was assumed that the choice of a drug that enables such “nonlethal” selection
is important to recover transplastomic clones (96, 151). However, transplastomic
clones were soon identified by kanamycin selection using an antibiotic concentration that is considered “lethal” (50 mg/L) (20). Thus, slow proliferation of nontransformed cells on a selective medium is not an essential feature of the selection
scheme.
Initial transformation vectors carried a plastid 16S rRNA (rrn16) gene with
point mutations that prevent binding of spectinomycin or streptomycin to the 16S
rRNA (140, 141, 151). The rrn16 target site mutations are recessive, and were
∼100-fold less efficient than the currently used dominant aadA gene (153). Streptomycin resistance encoded in the rps12 ribosomal protein gene was also included
in an early vector (140). Unexplored plastid mutations that could be used in vectors are lincomycin resistance, encoded in the 23S rRNA (29, 30), and triazine
resistance, encoded in psbA (31). Plastid mutations conferring resistance to spectinomycin, streptomycin, and lincomycin were also described in Solanum nigrum
(70). Solanum nigrum vectors with spectinomycin resistance (rrn16) and streptomycin resistance (rps12) markers were used to transform tobacco plastids (71).
More efficient primary plastid markers are chimeric genes in which the coding
segment of a bacterial antibiotic detoxifying enzyme is expressed from plastid signals. The aadA gene encodes aminoglycoside 300 -adenylyltransferase (AAD) also
called aminoglycoside nucleotidyltransferase [ANT(300 )-I] (128). AAD inactivates
spectinomycin and streptomycin (GenBank X02340, M10241) and was used to
select transplastomic clones in Chlamydomonas (50) and tobacco (153). The neo
(aph(30 )IIa) gene encodes neomycin phosphotransferase II [NPTII; APH(30 )-II]
(GenBank V00618) (128), and was used to select transplastomic clones in tobacco
(20). The aphA-6 gene encodes aminoglycoside phosphotransferase or APH(30 )VI (GenBank X07753) (128), and was used to select transplastomic clones by
kanamycin and amikamycin resistance in Chlamydomonas (8) and by kanamycin
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resistance in tobacco (64). Direct selection for spectinomycin resistance (153,
173) and for highly expressed kanamycin resistance genes (pHK30, pHK34) (83)
(A.K. Azhagiri, unpublished results), on average, yield one transplastomic line in
a bombarded leaf sample, although the values in this laboratory varied between 0.5
and 5.0.
Another potentially useful marker is a plant nuclear gene encoding betaine aldehyde dehydrogenase (BADH), which confers resistance to the toxic compound betaine aldehyde (BA) (36). BA selection is supposedly 25-fold more efficient than
spectinomycin selection (34). This claim has triggered research in other laboratories to confirm the advantage of selecting transplastomic clones by BA resistance.
So far direct selection of transplastomic tobacco clones by resistance to BA alone
is not confirmed (S.M. Whitney, J.T. Andrews, T. Golds & H.U. Koop, unpublished
results). It may be relevant that no transplastomic plants that carry the BADH gene
as the only selective marker have been reported.
Secondary Positive Selection
Use of secondary selective markers is dose dependent; they are not suitable to
select transplastomic clones when only a few ptDNA copies are transformed,
but will confer a selective advantage when most genome copies are transformed.
Examples for secondary markers are genes that confer resistance to the herbicides phosphinothricin (PPT) (92, 167) or glyphosate (167) or to the antibiotic
hygromycin [based on expression of the bacterial hygromycin phosphotransferase
gene (Z. Svab & S. Corneille, unpublished results)].
Negative Selection
The ability to identify loss-of-function of a conditionally toxic gene forms the
basis of negative selection. A negative selection scheme in plastids utilizes the
bacterial cytosine deaminase (CD) enzyme encoded in the codA gene (126). CD
catalyzes deamination of cytosine to uracil, enabling use of cytosine as the sole
nitrogen and pyrimidine source. CD is present in prokaryotes and in many eukaryotic microorganisms, but is absent in higher plants. 5-fluorocytosine is converted to
5-fluorouracil, which is toxic to cells. This negative selection scheme was utilized
to identify seedlings on 5-fluorocytosine-medium from which codA was removed
by the CRE-loxP site-specific recombinase (26).
Reporter Genes
The E. coli GUS and the Aequorea victoria GFP are reporter enzymes that allow
tracking gene expression, but do not confer a selective advantage or disadvantage
to plastids. GUS enzymatic activity expressed in chloroplasts has been measured
using fluorogenic assays (43, 141, 143, 172) and visualized by histochemical staining (141). GFP is a visual marker, allowing direct imaging of the fluorescent gene
product in living cells. Its chromophore forms autocatalytically in the presence of
oxygen and fluoresces green when absorbing blue or UV light. GFP has been used
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to detect transient gene expression (60) and stable transformation events (117, 130,
133, 135) in chloroplasts. GFP was fused with the aadA gene product (AAD) to
be used as a bifunctional visual and selective (spectinomycin resistance) marker
gene (72).
Combination of Visual and Selective Markers
Koop and colleagues (74) developed an ingenious scheme for rapidly identifying
transplastomic sectors using pigment-deficient tobacco knockout plants as recipients. In the knockout plants, an antibiotic resistance gene (aadA) replaces a plastid
gene that causes chlorophyll deficiency. The transformation vector carries the photosynthetic gene to restore green pigmentation and a second antibiotic resistance
gene to favor maintenance of transformed plastids. Homoplastomic sectors and
plants can be readily identified by the green color, a strategy that significantly
reduces the time required to obtain homoplastomic plants.
GENE DESIGN
A modular approach for transgene assembly was developed to conveniently shuffle
50 -regulatory regions, coding segments, and 30 -regulatory regions based on the
pUC18/19 polycloning site (for reviews, see 98, 100).
The 50 regulatory regions are provided in a PL cassette, which includes a promoter (P) and translation control sequences (L, leader). The translation control
sequences may be the mRNA 50 -UTR, or the 50 -TCR that includes the 50 -UTR and
the coding region N terminus (82, 83). The plastid genome contains many promoters (90). Biotechnological applications have focused on the strong, sigma70-type
rRNA operon (Prrn) promoter recognized by the plastid-encoded plastid RNA
polymerase. Prrn was fused with translation control sequences of plastid and phage
origin to facilitate translation of the encoded recombinant proteins. The 50 -UTR is
typically a truncated and mutant form of native sequences. Protein accumulation
from the same (Prrn) promoter may vary as much as 10,000-fold depending on the
choice of translation control signals (43, 82, 83, 169, 172; reviewed in 100).
The 30 -regulatory region or T cassette encodes the mRNA 30 -UTR, typically
including a stem-loop structure. The 30 -regulatory region is important for mRNA
stability (108). Most commonly used T cassettes derive from the plastid psbA,
rbcL, and rps16 genes (100).
PROBLEMS WITH DATA INTERPRETATION
Although the plastid marker genes are designed for expression in plastids, some
copies may fortuitously integrate in the nucleus and express from an upstream promoter to yield spectinomycin/streptomycin or kanamycin resistant clones (20, 28).
Transplastomic clones can be readily distinguished from the products of nuclear
insertion by the altered ptDNA on DNA gel blots.
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Spontaneous mutation to spectinomycin resistance is very common in all species
tested so far, including tobacco (151, 152), potato (133), Arabidopsis (134), and
Lesquerella (135). [So far, no spontaneous mutation for kanamycin resistance
has been found in tobacco, but it is common in Chlamydomonas chloroplasts
(57).] Transplastomic clones differ from spontaneous mutants by resistance to
both spectinomycin and the unselected streptomycin (Figure 2C) (153).
Plastid genome fragments, at least in some species, are present in the nuclear (5,
118) and/or in the mitochondrial genomes (110). For example, rice chromosome 10
alone carries two large ptDNA insertions (33 kb and 131 kb) encompassing almost
the entire ptDNA (118). Such nuclear or mitochondrial copies of the plastid rbcL
(5), ndh (78), and ycf9 (7, 95) genes are indicated by a persistent, weak wild-type
signal on DNA gel blots of total cellular DNA. However, the weak signal should
be absent on blots prepared with ptDNA from purified chloroplasts (101, 120),
and nontransformed, wild-type seedlings should be absent in the seed progeny
(101). A technically more demanding approach to verifying the plastid location
of transgenes utilizes PCR analysis of pulsed-field gel electrophoresis-purified
ptDNA (154).
IMPLEMENTATION IN DIFFERENT SPECIES
Although plastid transformation in higher plants was achieved in 1990 (151), it
is routine only in tobacco (153, 173). Plastid transformation has also been successful in two other solanaceous species, potato (133) and tomato (121). Plastid transformation in Arabidopsis thaliana (134) and the related Brassica napus
(62) and Lesquerella fendleri (135) was feasible but inefficient. Plastid transformation of embryogenic cultures in rice could be readily obtained. Rice plants
regenerated from the transformed culture were heteroplastomic (72), suggesting
that only refining the tissue culture system is required to obtain homoplastomic
plants. Spectinomycin resistance (aadA) was the marker of choice in each species,
except rice, in which streptomycin resistance was used to select transplastomic
clones. Spectinomycin selection in cereals is not an option because rice and maize
plastid rRNAs naturally have the mutations that prevent spectinomycin binding
(46, 72).
APPLICATIONS IN BASIC SCIENCE
Gene Knockouts
Targeted knockout of plastid genes involves construction of a transformation vector in which a selectable marker replaces the target gene in a larger ptDNA fragment (Figure 4). Selection for antibiotic resistance results in replacement of the
target gene with the selective marker in the ptDNA. Table 2 contains the list of
28 genes that have been targeted for deletion. Twenty-five genes have been deleted.
Positively identifying essential plastid genes has been problematic since attempts to
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Figure 4 Targeted deletion of a plastid gene (G2) by replacement with marker
gene (M1) in vector.
obtain homoplastomic ycf1, ycf2 (42), and clpP1 (132) knockout lines by targeted
insertion of a selective marker have failed. Evidence for an essential role for the
ClpP1 protease subunit was obtained by the CRE-loxP site-specific recombination
system (85). The clpP1 gene in the plastid genome was first flanked with directly
oriented loxP sites (floxed). A CRE gene was then introduced into the nucleus
by pollination. The nuclear-encoded, plastid-targeted CRE entered the plastids
and excised the floxed clpP1 copies. Loss of the clpP1 gene product, the ClpP1
protease subunit, led to ablation of the shoot system of tobacco plants, suggesting
that ClpP1-mediated protein degradation is essential for shoot development.
Overexpression
The advantage of overexpression as a research tool was shown by the unexpected
discovery of site-specific RNA editing trans-factors based on over-expression of an
edited RNA segment (23) and a clP1-specific mRNA maturation factor based on
overexpression of the clpP1 50 -UTR in a chimeric transcript (84). Overexpression
was used to probe the plastid accD function by replacing the weak accD promoter
with the strong rrn promoter (94). Another example for overexpression involved
relocating the nuclear anthranilate synthase gene to plastids to boost tryptophan
production (170).
Transcription
The field of plastid gene transcription also benefits from plastid transformation.
The plastid rpoA (127) and rpoB (1) tobacco knockout plants played a critical role
in recognizing that the plastid-encoded plastid RNA polymerase (PEP) and the
nuclear-encoded phage-type RNA polymerase (NEP) transcribe distinct groups
of genes (55, 127). Reporter genes expressing GUS and GFP have been utilized
to identify PEP and NEP promoter elements. Of the PEP promoters, dissection
in vivo was reported for the blue-light-regulated psbD promoter (1, 158), the rRNA
operon PEP promoter (150), the rbcL promoter (129), and the psbA promoter (58).
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TABLE 2 Plastid genes targeted for deletion
Gene
Function
References
rpoA
PEP
(40, 127)
rpoB
PEP
(2, 40)
rpoC1
PEP
(40, 127)
rpoC2
PEP
(127)
trnV
tRNA-Val(GAC)
(26, 56)
sprA
RNA
(147)
oriA
DNA replication
(109)
DNA replication
(109)a
ndhA
Ndh
(78)
ndhB
Ndh
(61, 131)
ndhC
Ndh
(19, 78)
ndhH
Ndh
(78)
ndhI
Ndh
(78)
ndhJ
Ndh
(19)
ndhK
Ndh
(19, 78)
rbcL
Rubisco
(68)
psbA
PSII
(6)
psbE
PSII
(156)
psbF
PSII
(156)
psbL
PSII
(156)
psbJ
PSII
(54, 156)
clpP1
oriB
∗
Protease
(132)∗ (85)
a
ycf1
?
(42)a
ycf2a
?
(42)a
ycf3
PSI
(122)
ycf6/petN
cyt. B6/f
(53)
ycf9/lhbA/psbZ
PSII
(7, 95, 120, 155)
ycf10
?
(154)
a
No homoplastomic knockout plant was reported.
Of the promoters recognized by the NEP, the clpP1 (137) and atpB (166) promoters
have been subjected to in vivo dissection.
RNA Editing
RNA editing in plastids involves posttranscriptional C to U nucleotide conversions
(11, 160). Plastid transformation has been an important tool in studying RNA
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editing in chimeric transcripts consisting of a series of small mRNA segments
transcriptionally fused with a reporter gene (23). Sequences required for RNA
editing are contained in a small segment (13, 24); RNA editing depends on sitespecific depletable trans-factors (22, 23, 116); if a site is not edited in species it is
an indication that the capacity to edit the site is absent (15) unless the species has
sites that share specificity factor(s) with the heterologous site (22).
Photosynthesis
Because many photosynthetic genes are encoded in the plastid genome, plastid
transformation is a necessary tool to probe and improve photosynthesis. Studies
in higher plants focused on photosynthetic gene knockouts (Table 2) and Rubisco
engineering (for a review, see 3). Rubisco in higher plants is composed of plastidencoded large and nucleus-encoded small subunits (LS and SS, encoded in the
plastid rbcL and nuclear rbcS genes, respectively). The plastid-encoded tobacco
rbcL gene was replaced with a heterologous sunflower gene (69), genes from
nongreen algae (165), cyanobacteria (Synechococcus PCC6301) (69), and the αproteobacterium Rhodospirillum rubrum (163, 164). Replacing the tobacco rbcL
gene with the Rhodospirillum rubrum Form II Rubisco yielded a fully photoautotrophic, fertile plant (163). Allotopic expression of the Rubisco subunits has been
explored by relocating the plastid rbcL gene to the nucleus (68), and the nuclear
small subunit gene to the plastid genome (162, 171).
Evolution
During evolution most of the ∼3000 genes encoded in the genome of the photosynthetic endosymbiont migrated to the nucleus (89, 102). Plastid transformation
enabled addressing experimentally the mechanistic details of gene transfer. The
frequency of DNA transfer from plastids to the nucleus was tested by incorporating a nuclear kanamycin resistance gene in the plastid genome, then selecting
for a transfer event to the nucleus by selecting for expression of a kanamycin resistance gene. The frequencies of transfer were 1 in 16,000 in the seed progeny
(63) and significantly lower (1 in 5,000,000) in somatic cells (145). Of course,
acquiring a nuclear promoter by a transferred plastid gene is several orders of magnitude less likely than expressing a plastid gene that brings along its own nuclear
promoter. The feasibility of relocating a plastid gene to the nucleus was also tested
(68). First, the plastid-encoded rbcL copy was removed by targeted deletion yielding a Rubisco-deficient plant. Absence of a functional Rubisco in the knockout
plants established that none of the ∼15 nuclear rbcL copies contained in chunks
of incorporated ptDNA fragments are functional. The LS coding region was then
extended at the N terminus to provide a plastid-targeting transit sequence and was
incorporated in the nucleus of knockout plants. Rubisco levels were restored up
to ∼10% of the wild-type levels. Successfully relocating the plastid rbcL gene to
the nucleus supports the view that gene migration from plastids to the nucleus is
an ongoing process.
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APPLICATIONS IN BIOTECHNOLOGY
Agronomic Traits
In field crops, the most common transgenic traits are resistance to insects and herbicides expressed from nuclear genes. Both types of genes have been successfully
engineered for plastid expression (Table 3). Expression of the B.t. insecticidal
protein from nuclear genes required construction of synthetic, codon-modified
genes to improve translation, protect the mRNA from degradation, and prevent
early translation termination. In contrast, the B.t. insecticidal protein genes were
expressed in plastids from bacterial coding segments (39, 81, 103, 115; for a review,
see 100). Commercially useful versions of commonly used herbicide resistance
genes are also available. Tobacco plants with some degree of glyphosate tolerance
were obtained by overexpression of the sensitive form of 5-enolpyruvylshikimate3-phosphate synthase (EPSPS) target enzyme in plastids (33). Field-level tolerance
to glyphosate was obtained by expression in plastids of prokaryotic EPSPS genes;
the required protein levels were higher (∼5% of TSP) than when EPSPS was
expressed from nuclear genes (169). An interestingly split EPSPS gene was developed with gene containment in mind, with one half of the protein encoded in
a nuclear gene and the second half in a plastid gene. Intein trans-splicing resulted
TABLE 3 Plastid transgenes for biotechnological applications
Gene
Function
References
bar
Herbicide res./PPT
(65, 92, 167)
CP4
Herbicide res./glyphosate
(167, 169)
Ic-EPSPSc
Herbicide res./glyphosate
(25)
cry1Ac
B.t. Insecticidal prot.
(103)
cry2Aa2
B.t. insecticidal prot
(39, 81)
cry1Ia5
B.t. insecticidal prot.
(115)
Somatotropin
Human growth hormone
(139)
CTB
Cholera toxin B subunit
(35)
TetC
Tetanus vaccine
(159)
HSA
Human serum albumin
(45)
PBP
Protein-based polymer
(51)
ASA2
Tryptophan biosynthesis
(170)
MS1-99
Antimicrobial peptide
(41)
phb operon
Polyhydroxybutyrate
(91)
TPS1
Trehalose phosph. synthase
(88)
merA
Mercuric ion reductase
(123)
merB
Organomercurial lyase
(123)
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in reconstituting the herbicide-resistant EPSPS enzyme in plastids (25). Pollen
transmission from this crop can lead to the transfer of only part of the herbicideresistance gene, which is insufficient to confer glyphosate tolerance. Field-level
tolerance to herbicides containing PPT as an active ingredient was obtained by
expressing bar in the plastid genome (92, 167).
Expression of Recombinant Proteins
To meet the demands for production capacity of recombinant proteins, there is
significant interest in plant-based production of vaccines, antibodies, and industrial enzymes (47, 87, 93, 146). Transgene expression in tobacco plastids reproducibly yields protein levels in the 5% to 20% range (for a review, see 100).
A salient feature of plastid expression is the importance of post-transcriptional
regulation; from the same promoter proteins may accumulate in a 10,000-fold
range (for references, see Gene Design, above). Since all codons are relatively
frequently used, codon optimization yields only a modest 2- to 3-fold increase
in protein accumulation levels (92, 159, 169). Transcripts derived from genes of
diverse sources were stable in plastids, including bacterial genes with relatively
high levels of adenine and thymine (high-AT) (103, 115) (159) and guanine and
cytosine (high-GC) (92), synthetic mRNAs (159), and plant (170) and human (45)
cDNAs. This suggests the compatibility of the plastid’s RNA degradation machinery with mRNAs from diverse sources, avoiding the need to construct synthetic
genes for plastid expression. Furthermore, there is no protein glycosylation in
plastids (159).
Marker Gene Elimination
The interest in developing marker elimination systems for plastids was driven by
regulatory concerns to avoid releasing antibiotic resistance genes in transplastomic
crops, the desire to reuse the relatively few available plastid marker genes, and the
metabolic burden imposed by expressing marker genes. Three systems are available for marker gene elimination. The first system relies on the loop-out of the
marker gene through directly repeated sequences (65). This system is practical
only in exceptional cases, when introducing secondary markers, such as herbicide resistance genes (65), is the desired objective. A second approach involves
cotransforming two independently targeted plastid transgenes and segregating out
the ptDNA with the marker gene at the heteroplastomic stage (167). The third
and most efficient approach uses vectors with floxed marker genes, which can be
removed with the CRE site-specific recombinase. Although convenient vectors
with floxed marker genes have not yet been reported for the introduction of passenger genes, the feasibility of the approach was shown by excising aadA (56),
codA (26) and clpP1 (85) genes (Figure 3B). Important for the application of
CRE in plastids is that no detrimental ptDNA rearrangements persist once CRE is
removed (27).
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Containment by Plastid Localization
Transgene flow is a problem within a species, when pollen from one stand of the
crop finds its way into another stand (119), or when transgenes are incorporated in
weedy relatives (44). Effectively controlling intraspecific transgene flow in species
with a strict maternal inheritance of plastids can be achieved by incorporating the
transgenes in the plastid genome. Examples for species with strict maternal inheritance of plastids are Zea mays, Glycine max, Oryza sativa, and Arabidopsis
thaliana, a group to which two thirds of the angiosperm species belong (52, 105,
114). Since more sensitive, selectable plastid markers are available, strict maternal inheritance of plastids has been questioned in Nicotiana tabacum and Nicotiana plumbaginifolia. Two publications reported low-level pollen transmission of
plastome-encoded streptomycin resistance (104) and tentoxin resistance (4) at a
frequency of 0.07% to 2.5% of the progeny, respectively. Low frequency (0.03%)
pollen transmission of plastids was also reported in a Setaria italica (foxtail or
birdseed millet) cross (161). In each case, at least one of the parental lines was a
cytoplasmic substitution line, in which the nucleus of one species was combined
with the plastids and/or mitochondria of another species. Strict maternal inheritance is known to break down in species hybrids (73), and that may explain low
frequency of paternal pollen transmission in these examples.
PERSPECTIVES
During the past decade plastid transformation has become a principal tool of plastid biology. The most important task for the coming years will be implementing
plastid transformation in the major crops. The key to progress will be identifying bottlenecks in the recalcitrant species and combining suitable tissue culture
systems with efficient molecular tools. In agronomic applications, incorporating
transgenes in the plastid genome instead of the nucleus will be an excellent tool
to control gene flow in crops with maternal plastid inheritance. Plastid localization will be a significant improvement as compared to the present practice of
incorporating transgenes in the nuclear genome, when 100% of pollen leaving
the field carries the transgenic information. Part of breeding transplastomic crops
will require testing paternal plastid transmission and, if necessary, screening for
lines in which paternal transmission of plastids does not occur. From here on,
the feasibility of functional transfer of plastid genes to the nucleus will also be a
consideration. A predictable outcome is a glut of new smart gene designs, such as
editing-dependent genes (23), genes with plastid introns (16), and split genes (25),
which minimize the opportunity for expression of plastid genes after transfer to
the nucleus. Although basic science applications, such as probing photosynthetic
functions and plastid gene expression will remain important, plastid transformation has now reached a more mature phase when it is expected to make a broader
impact through agricultural and industrial applications.
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ACKNOWLEDGMENTS
I thank members of my laboratory for their comments during the preparation of
the manuscript; Arun Azhagiri for assistance with the figures; and John Andrews,
Ralph Bock, Timothy Golds, Jonathan Gressel, Hans-Ulrich Koop, Jeffrey Staub,
and Spencer Whitney for communicating unpublished results and supplying copies
of their manuscripts prior to publication. The National Science Foundation Biochemistry and Eukaryotic Genetics Program, The Rockefeller Foundation, Monsanto Co., and Rutgers F&A Special Research Grant supported research in this
laboratory.
The Annual Review of Plant Biology is online at http://plant.annualreviews.org
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Figure 1 Sorting ptDNA at the organelle and cellular levels yields homoplastomic
plants. (A) Chloroplast (CHL) to proplastid (PP) dedifferentiation accelerates ptDNA
sorting. Transformed and nontransformed nucleoids (N) in plastids are symbolized
with red and blue circles, respectively. Transformed and nontransformed ptDNA in
enlarged nucleoids are red and blue circles, respectively. ptDNA are anchored to
membranes by proteins (black dots). Nucleoid 1 is heteroplastomic and is the progenitor of homoplastomic transgenic (1a) and wild-type (1b) proplastids. Homoplastomic proplastid differentiates into chloroplast (bottom). Wild-type proplastids (2, 1b)
are antibiotic sensitive and divide more slowly. (B) Reduction in plastid number during chloroplasts (∼100 per leaf cell; green, elongated ) to proplastid (∼10–14 per
meristematic cell; greenish, oval) transition and lack of exact duplication of the cytoplasm accelerates formation of homoplastomic cells. On top is a leaf mesophyll cell
with one transformed chloroplast (red) and nucleus (Nu). Meristematic cell 1 is heteroplastomic. Cleavage of the cytoplasm yields one meristematic cell with transformed chloroplasts only (1a) and one with wild-type plastids (1b). Cell 1a is the
progenitor of homoplastomic mesophyll cells (1c). Reproduced with permission
(99).
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PLASTID TRANSFORMATION
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Figure 2 Selection of transplastomic clones by spectinomycin resistance. (A) Spectinomycin inhibits callus formation, greening, and shoot regeneration from tobacco
leaf segments on shoot regeneration medium. Transplastomic clones are resistant to
spectinomycin and are identified as green shoots or calli (153). (B) The shoots are
chimeric, visualized by accumulation of green fluorescent protein in transplastomic
sectors. Spectinomycin resistance is not cell autonomous as sensitive sectors are also
green. (72) (C) Spontaneous spectinomycin resistant mutants are sensitive (top),
transplastomic clones are resistant to streptomycin (bottom) when cultured on a
selective streptomycin (500 mg/L) medium (153).