Département de Sciences Biologiques, Université de Montréal, 4101 Sherbrooke est, Montréal, Québec, Canada H1X 2B2
* Author for correspondence (e-mail: david.morse{at}umontreal.ca)
Accepted 26 March 2003
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Dinoflagellate, Plastid, Nuclear-encoded protein, Protein import, Signal peptide
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Several mechanisms are known for directing proteins to four membrane-bound
plastids. In one mechanism, such as that used by haptophytes and diatoms,
ribosomes are attached directly to the outer plastid membrane. After passage
of the first (outermost) membrane, termed the chloroplast ER
(Gibbs, 1981), proteins either
cross the second membrane through large pores, or traverse the subsequent
intermembrane space inside transport vesicles
(van Dooren et al., 2001
). In
contrast to this mechanism, targeting to the four membrane-bound plastids of
the apicomplexans does not involve ribosomes bound to the outer membrane, and
indeed, many aspects of the mechanism are still unknown
(van Dooren et al., 2001
). It
is known that plastid proteins enter the ER membrane system using an
N-terminal signal peptide, since constructs lacking the signal peptide produce
a cytoplasmic protein (Waller et al.,
2000
). It is also known that a transit peptide, exposed after
cleavage of the signal peptide, is required for entry into the plastid, as
proteins lacking this transit peptide are secreted from the cell
(Waller et al., 2000
).
However, several mechanisms have been suggested for targeting proteins to the
outermost plastid membrane from the ER
(van Dooren et al., 2001
). One
proposal is that all proteins entering the ER eventually pass through the
outermost plastid compartment, and those destined to remain in the plastid
bind to a receptor for the transit peptide. Alternatively, proteins may be
specifically targeted to the outer plastid membrane directly from the Golgi.
While the observation that plastid directed proteins lacking a transit peptide
are secreted suggests passage through the Golgi, it must be stressed that this
has not been directly shown.
Intriguingly, nuclear-encoded proteins destined for the triple
membrane-bound plastids of Euglena clearly transit through the Golgi
(van Dooren et al., 2001).
Details of the mechanism differ, however, as in Euglena the
plastid-directed leader sequences contain two hydrophobic regions separated by
an S/T-rich region reminiscent of a plastid transit sequence
(Henze et al., 1995
;
Kishore et al., 1993
). In
vitro studies have demonstrated that the plastid directed proteins remain
inserted in the ER membranes, unlike the apicomplexans where the plastid
proteins are soluble inside the ER
(Kishore et al., 1993
;
Osafune et al., 1990
;
Sulli et al., 1999
;
Sulli and Schwartzbach, 1996
).
The second hydrophobic region present in the Euglena leaders is
absent in apicomplexan leaders (Foth et
al., 2003
).
In contrast to the above, there are no reports describing the transport
mechanism used by nuclear-encoded plastid proteins in dinoflagellates. This is
an unfortunate gap in our understanding of protein import pathways as
dinoflagellates, along with diatoms, constitute the bulk of the phytoplankton,
and the oceans are responsible for roughly half the primary production of the
biosphere (Field et al., 1998).
The dinoflagellates contain members with several types of plastids, of which
those containing the carotenoid peridinin are the most prevalent. The next
most abundant type of plastids contains fucoxanthin, and these are proposed to
share a common ancestor with the peridinin-containing plastids
(Yoon et al., 2002
). Both
types of plastids are surrounded by three membranes, and may thus employ a
similar mechanism for protein import. On the one hand, this mechanism might be
expected to be similar to that of apicomplexans because of their close
phylogenetic relationship to the dinoflagellates
(Sogin et al., 1996
). On the
other hand, protein import into dinoflagellate plastids might also be similar
to that in Euglena, because the leader sequences of dinoflagellate
plastid-directed proteins (Fagan et al.,
1998
; Le et al.,
1997
) are similar to those used by Euglena.
To distinguish between these two possibilities, we have characterized the
import pathway into peridinin-containing plastids of the dinoflagellate
Gonyaulax polyedra. We have studied import of two proteins for which
we have previously raised antibodies
(Nassoury et al., 2001), the
carbon fixing enzyme Ribulose bisphosphate carboxylase/oxygenase (Rubisco) and
the soluble light-harvesting peridinin-chlorophyll a-protein (PCP).
Dinoflagellate Rubisco is unusual, in that it is a form II enzyme and is
formed only of large 55 kDa subunits
(Morse et al., 1995
). This
enzyme is synthesized as a polyprotein, which must be processed to form the
mature large subunits by cleavage of small linker peptides
(Rowan et al., 1996
). The
mechanism and location of this processing event is unknown, although the final
destination of the protein is the stroma, as determined by immunolocalization
(Nassoury et al., 2001
). The
soluble PCP is not a polyprotein, and it is located in the thylakoids, also as
determined by immunolocalization (Nassoury
et al., 2001
). The synthesis of both proteins is under control of
the circadian clock in vivo, and their expression times differ slightly. When
the algae are grown under a 12:12 light:dark (LD) regime, Rubisco is
synthesized from about midnight to midday (LDT 18 to 6), while PCP is
synthesized from dawn to dusk (LDT 0 to 12)
(Markovic et al., 1996
). We
found that nuclear-encoded proteins passed through the Golgi on route to the
plastid, as has been shown for Euglena
(Schiff et al., 1991
).
Furthermore, we show that the dinoflagellate plastid-directed protein leader
sequences are functionally similar to those found in Euglena. We
suggest that the second hydrophobic region present in the leader is a
mechanistic requirement dictated by the triple membrane architecture of the
plastids.
![]() |
Materials and Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In vitro translation and processing experiments
Constructs 1-2LUC and 2LUC were translated using a T7 TnT-coupled rabbit
reticulocyte system (Promega Biotech) containing 35S-methionine
(ICN) in the presence or absence of 4 µl canine pancreatic microsomal
membranes (Promega) as described by the supplier. Protease protection assays
were performed by incubating the translation mix with 0.1 mg/ml trypsin in the
presence or absence of 0.5% Triton X-100. Sedimentation of membrane-bound
translation products was performed by centrifugation in an airfuge using an
A-100/30 fixed angle rotor for 1 hour at 100,000 g.
Translation products were visualized by autoradiography after separation on
12% SDS polyacrylamide gels and electrophoretic transfer onto nitrocellulose
membranes.
Immunocytochemistry and transformation
Gonyaulax cells were cultured and prepared for electron
immunocytochemistry as described, using antibodies directed against a
Gonyaulax Rubisco expressed from a cDNA in bacteria or PCP purified
by column chromatography from the algae as described
(Nassoury et al., 2001). A 20
nm gold-labeled goat anti-rabbit was used for detection (Ted Pella). A similar
procedure was followed for immunolabeling of S. chacoense cells,
except that a commercial anti-luciferase (Promega Biotech) was used as a
primary antibody. A 20 nm gold-labeled rabbit anti-goat was used for detection
(Ted Pella).
Sequence analysis
The sequences used for multiple alignments were obtained from GenBank using
the accession numbers (top to bottom in
Fig. 2) X94549, U93077,
AF298221, AJ009670, AF028561, D14702, X89768, L21904, X15743, X66617 and
S53593. Sequences for hydrophobicity plots (in
Fig. 4) were AF087139, AF028561
and L21904. All analyses were performed using MacVector software, including
ClustalW multiple alignments and neighbour-joining phylogenetic analyses.
Bootstrap values are given for 10,000 trees.
|
|
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
The plastid targeting sequence contains a signal peptide
To rule out the possibility that the observed Golgi labeling was a
non-specific response to the antibodies, we immunolabelled sections from cells
isolated at midday (LDT 6) with the anti-Rubisco. At this time in their daily
cycle, the in vivo synthesis rate of Rubisco is low
(Markovic et al., 1996). We
observed a constant labeling of the plastid
(Fig. 1C), in agreement with
previous results showing no difference in the total Rubisco levels detected on
Western blots and by enzyme-linked immunosorbent assay at different times over
the daily cycle (Nassoury et al.,
2001
). However, no label was found over the Golgi, confirming that
the labeling seen at time of high synthesis rates was indeed due to the
presence of Rubisco. This control experiment was also performed using cells
isolated at midnight (LDT 19) and the anti-PCP antiserum. At this time, PCP
synthesis rates are low (Markovic et al.,
1996
), and again, no labeling of the Golgi was observed
(Fig. 1F). We conclude that
both proteins must enter the Golgi from the ER at times of high synthesis
rates.
Targeting to the ER typically requires the presence of an N-terminal
hydrophobic signal peptide. Indeed, such a signal, followed by an AXA-type
signal peptidase site, is found in the leader sequences of plastid directed
proteins for both dinoflagellates and Euglena. In Euglena,
the two hydrophobic regions have been proposed to act as start and stop
signals for translocation across the ER membrane
(Sulli et al., 1999). This
topology results in a membrane-bound protein with the first signal peptidase
site inside the ER lumen and the bulk of the protein outside.
We thus tested if dinoflagellate plastid leader sequences acted similarly
to those studied in Euglena. For our in vitro analyses, we chose to
examine the leader sequence of the thylakoid protein PCP. The leader sequence
of this protein is virtually identical to that of the stromal protein
glyceraldehyde-3-phosphate dehydrogenase (GAP)
(Fagan et al., 1999),
suggesting that the information to distinguish between stromal and thylakoid
locations might be found within the coding sequence of the mature PCP. To
avoid potentially conflicting signals, the complete PCP leader sequence,
encoding both hydrophobic regions and the S/T-rich regions separating them
(Le et al., 1997
), was thus
fused with a luciferase reporter gene. These constructs were transcribed and
translated in vitro and the protein products analyzed by SDS PAGE. The size of
the protein produced is as predicted when the transcripts are translated in
the absence of canine microsomes, and is smaller when translated in the
presence of canine microsomes. As expected, the processed form is larger than
luciferase alone (Fig. 2A). We
conclude that the size change upon addition of microsomes is consistent with
cleavage of the signal peptide at the expected site.
This cleavage of the signal peptide following the signal peptide suggested that the first hydrophobic region is inserted into the microsomal membrane with the N-terminus in the cytoplasm. If so, then the second hydrophobic region should act as a stop transfer signal, and the final topology of the protein, after cleavage by the signal peptidase, would be a single pass membrane protein with its C-terminal end in the cytoplasm. To test this, we digested with trypsin the protein produced in the presence of microsomes. This treatment reduced the level of radiolabelled protein, and no further decrease in the amount of protein could be obtained by inclusion of Triton X-100 in the digestion (Fig. 2A). This indicated that the small amount of signal resisting digestion did not represent luciferase inside the microsomes. The topology of the protein deduced from this experiment is unambiguous and is illustrated at right (Fig. 2A).
The leader contains a functional plastid transit sequence
These results also suggested that the second hydrophobic region might act
as a signal peptide if the first hydrophobic region was deleted, as has been
observed in Euglena (Sulli et
al., 1999). We thus prepared and translated a construct lacking
the first hydrophobic region. The new construct showed no size decrease upon
addition of microsomes, and the protein product was sensitive to trypsin
digestion (Fig. 2B). To test
the possibility that the hydrophobic region was inserted into the membrane,
the labeled protein was centrifuged at 100,000 g for one hour
following the translation. The labeled protein is found in the pellet,
suggesting that the protein is either membrane bound or is tightly associated
with the microsomal membrane. A likely topology for the protein, based on both
these results and by analogy to results obtained with Euglena, is
illustrated in the schema at right (Fig.
2B).
The topology of the plastid-directed proteins deduced from the in vitro
experiments described above suggests that plastid precursor proteins may
arrive at the plastid anchored in the membrane of a transport vesicle with
their S/T-rich region inside the vesicle lumen. Fusion of the vesicle with the
outer plastid membrane would make the S/T-rich sequence available to protein
translocators spanning the inner two plastid membranes. We thus predicted that
S/T-rich sequence would be equivalent to the transit sequence of higher plant
plastid proteins. To test this, we asked if the translocators in the double
membrane-bound plastids of higher plants could recognize this sequence in
vivo. In vivo experiments allow a potential import into plastids to be
evaluated in the context of competition with other compartments, unlike in
vitro tests where only import into isolated pea chloroplasts is assessed
(DeRocher et al., 2000;
Wastl and Maier, 2000
).
We prepared a reporter gene construct containing the leader sequence
lacking the first hydrophobic region. This gene fusion, containing a
Solanum chacoense 5'UTR and driven by the constitutive 35S
promoter, was used to transform the wild potato S. chacoense
(Matton et al., 1997). Leaves
of eight transformed plants were screened and two selected for analysis on the
basis of high signal with the antibody. The anti-luciferase antibody detects
only the luciferase and has no reaction with untransformed plants and plants
with low expression of the transgene (Fig.
3C). However, in the highly expressing plants, the subcellular
location of the fusion protein was clearly the chloroplast. The presence of
the reporter in the plastids confirms that the S/T-rich region acts as an
authentic plastid targeting sequence in higher plants as well
(Fig. 3A,B). To quantify the
immunolabeling, five random pictures were taken from each sample, and the
number of gold beads determined for plastid, vacuolar and cytoplasmic
compartments. The label density in the plastids of transformed plants is 5- to
10-fold greater than that in other compartments within the same plant, and
over 10-fold greater than the background labeling observed in the plastids of
untransformed plants (Fig. 3D).
Presumably, this conservation of function reflects the common evolutionary
ancestor to all extant plastids. We have not observed staining in the ER or
nuclear membranes in these experiments. However, we cannot completely rule out
some ER targeting, as we did not detect any signal above background in plants
transformed with a luciferase reporter fused to the full-length leader
sequence. If proteins directed to the ER membrane in this manner were
unstable, for example, a direct comparison of the staining intensity in the
two compartments would be impossible. In any event, it is clear that the
S/T-rich region acts as an authentic transit peptide in higher plants.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Similar to the leader sequences used to target other complex plastids, the dinoflagellate leader contains a hydrophobic N-terminal signal peptide and an internal plastid transit sequence. Unlike most other leader sequences, the dinoflagellate sequences contain a second hydrophobic region at the C-terminal end. This unusual feature is also found in Euglena, as shown by an alignment of five dinoflagellate sequences and six Euglena sequences (Fig. 4A). This alignment is color coded for visualization of the hydrophobic character (blue boxes). An AXA-type signal peptidase site follows the first hydrophobic regions (arrow) here manually aligned for comparison. The serine/threonine-rich regions (yellow boxes) that constitute the plastid transit sequence separate the two hydrophobic regions (Fig. 4A). Since Euglena and the dinoflagellates share a triple bound plastid architecture, we suggest that the C-terminal hydrophobic region, and the unusual protein topology that results from its insertion in the membrane of the transport vesicles, are required by the organelle's structure.
It is instructive to consider the dinoflagellate targeting mechanism in the
light of the phylogenetic relationships between the different algal classes.
First, it is generally accepted that a close phylogenetic relationship exists
between apicomplexans and dinoflagellates
(Fast et al., 2002;
Sogin et al., 1996
). This
relationship can be illustrated by the molecular phylogeny of
glyceraldehyde-3-phosphate dehydrogenase (GAP). Many examples of these trees
are available in the literature, and the neighbor-joining tree shown here
(Fig. 4B) is illustrative of
much larger taxonomic samplings (Fast et
al., 2001
). These trees show the close relationship between
apicomplexan and dinoflagellate hosts, as well as the relationship between
euglenoids and trypanosomes. An interesting feature of the GAP phylogeny is
the clustering of plastid directed sequences in the apicomplexans and the
dinoflagellates (Fig. 4B)
(Fast et al., 2001
). This
phylogeny is supported by some plastid sequences
(Fast et al., 2002
;
Wilson et al., 1996
;
Zhang et al., 2000
), but not
by a tufA phylogeny (Kohler et
al., 1997
) or the presence of a green algal-like mitochondrial
cox2 gene in the apicomplexan nucleus
(Funes et al., 2002
) that
suggest green algal ancestry for the apicoplast. In any event, plastids from
dinoflagellates and Euglena have had different evolutionary origins,
even though both have three membranes and a second hydrophobic region in the
leader sequences (Fig. 4B). In
the model discussed below, we consider the case where dinoflagellate and
apicomplexan plastids share a common ancestor. However, it is important to
note that similar conclusions can also be reached in other scenarios, provided
the plastids of dinoflagellates initially had four bounding membranes.
One possible evolutionary scheme, based on the GAP phylogeny shown
(Fig. 4B), would involve uptake
of a red algal symbiont by the common ancestor to both dinoflagellates and
apicomplexans (Fig. 4C). Genes
that had been transferred to the red algal nucleus, either before or after
this secondary endosymbiotic event, would then be transferred to the new host
cell nucleus. Given the ability of the dinoflagellate S/T-rich transit
sequences to still function in higher plants
(Fig. 3), any sequences
transferred to the new host nucleus presumably included this plastid transit
sequence. To re-enter the plastids, now bounded by four membranes, a signal
peptide would have to be acquired in addition to the transit peptide
(McFadden, 1999). Thus,
according to this scheme, plastid-directed proteins in the last common
ancestor for the apicomplexans and dinoflagellates would be expected to have
both a signal peptide and a transit peptide in their leaders. The divergence
of the dinoflagellates, and the loss of one bounding membrane around their
plastids, must then have been accompanied by the selection of modifications in
the C-terminal end of the leader sequence with an accentuated hydrophobicity.
This new hydrophobic region, common to the sequence of plastid-directed
nuclear proteins of dinoflagellates and Euglena
(Fig. 4A), thus appears to be
an essential element for targeting nuclear-encoded proteins to triple
membrane-bound plastids.
The scheme above posits a transfer of genes to the nucleus of the new host
cell before loss of the outermost plastid membrane. So far, however, it has
not been possible to date the timing of these events. An alternative model, in
which membrane loss occurred prior to gene transfer, would require hydrophobic
signal peptide-like sequences to be inserted both before and after the transit
peptide. A third model, where genes were transferred from the plastid genome
directly to the new host nucleus, is not favored because of the fact that the
reduced gene content of most extant plastids shows considerable overlap
(Palmer and Delwiche, 1996).
In any event, the peptide leader sequence does not recapitulate the gene
transfer process, as at least part of the leader is defined not by phylogeny
but by the mechanistic requirements of protein targeting to triple
membrane-bound plastids.
What mechanistic requirement may be filled by the second membrane-spanning region in the dinoflagellate leader? One possible role may be to present the plastid transit sequence to the protein translocators in the inner (second) membranes. However, it may also act as a recovery mechanism should any of the plastid proteins be misdirected to the plasma membrane, as soluble proteins secreted from the cell would be irretrievably lost. Indeed, as little is known about the targeting mechanism that delivers proteins specifically to the plastid, this now represents the next challenge in elucidating the protein import mechanisms in dinoflagellates and Euglena.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Bruce, B. D. (2000). Chloroplast transit peptides: structure, function and evolution. Trends Cell Biol. 10,440 -447.[CrossRef][Medline]
Delwiche, C. and Palmer, J. (1996). Rampant horizontal transfer and duplication of rubisco genes in eubacteria and plasmids. Mol. Biol. Evol. 13,873 -882.[Abstract]
DeRocher, A., Hagen, C. B., Froehlich, J. E., Feagin, J. E. and
Parsons, M. (2000). Analysis of targeting sequences
demonstrates that trafficking to the Toxoplasma gondii plastid branches off
the secretory system. J. Cell Sci.
113,3969
-3977.
Fagan, T., Hastings, J. and Morse, D. (1998). Phylogeny of Glyceraldehyde-3-phosphate dehydrogenase indicates lateral gene transfer from cryptomonads to dinoflagellates. J. Mol. Evol. 47,633 -639.[Medline]
Fagan, T., Morse, D. and Hastings, J. (1999). Circadian synthesis of a nuclear encoded chloroplast Glyceraldehyde-3-phosphate dehydrogenase in the dinoflagellate Gonyaulax polyedra is translationally controlled. Biochemistry 38,7689 -7695.[CrossRef][Medline]
Fast, N. M., Kissinger, J. C., Roos, D. S. and Keeling, P.
J. (2001). Nuclear-encoded, plastid-targeted genes suggest a
single common origin for apicomplexan and dinoflagellate plastids.
Mol. Biol. Evol. 18,418
-426.
Fast, N. M., Xue, L., Bingham, S. and Keeling, P. J. (2002). Re-examining alveolate evolution using multiple protein molecular phylogenies. J. Eukaryot. Microbiol. 49, 30-37.[Medline]
Field, C. B., Behrenfeld, M. J., Randerson, J. T. and Falkowski,
P. (1998). Primary production of the biosphere: integrating
terrestrial and oceanic components. Science
281,237
-240.
Foth, B. J., Ralph, S. A., Tonkin, C. J., Struck, N. S.,
Fraunholz, M., Roos, D. S., Cowman, A. F. and McFadden, G. I.
(2003). Dissecting apicoplast targeting in the malaria parasite
Plasmodium falciparum. Science
299,705
-708.
Funes, S., Davidson, E., Reyes-Prieto, A., Magallon, S., Herion,
P., King, M. P. and Gonzalez-Halphen, D. (2002). A
green algal apicoplast ancestor. Science
298, 2155.
Gibbs, S. (1981). The chloroplasts of some algal groups may have evolved from endosymbiotic eukaryotic algae. Ann. N. Y. Acad. Sci. 361,193 -208.[Medline]
Henze, K., Badr, A., Wettern, M., Cerff, R. and Martin, W. (1995). A nuclear gene of eubacterial origin in Euglena gracilis reflects cryptic endosymbioses during protist evolution. Proc. Natl. Acad. Sci. USA 92,9122 -9126.[Abstract]
Kishore, R., Muchhal, U. S. and Schwartzbach, S. D. (1993). The presequence of Euglena LHCPII, a cytoplasmically synthesized chloroplast protein, contains a functional endoplasmic reticulum-targeting domain. Proc. Natl. Acad. Sci. USA 90,11845 -11849.[Abstract]
Kohler, S., Delwiche, C. F., Denny, P. W., Tilney, L. G.,
Webster, P., Wilson, R. J., Palmer, J. D. and Roos, D. S.
(1997). A plastid of probable green algal origin in Apicomplexan
parasites. Science 275,1485
-1489.
Le, Q. H., Markovic, P., Jovine, R., Hastings, J. and Morse, D. (1997). Sequence and genomic organization of the peridinin-chlorophyll a-protein from Gonyaulax polyedra. Mol. Gen. Genet. 255,595 -604.[CrossRef][Medline]
Markovic, P., Roenneberg, T. and Morse, D. (1996). Phased protein synthesis at several circadian times does not change protein levels in Gonyaulax. J. Biol. Rhythms 11,57 -67.[Medline]
Martin, W. and Schnarrenberger, C. (1997). The evolution of the Calvin cycle from prokaryotic to eukaryotic chromosomes: a case study of functional redundancy in ancient pathways through endosymbiosis. Curr. Genet. 32,1 -18.[CrossRef][Medline]
Martin, W., Stoebe, B., Goremykin, V., Hapsmann, S., Hasegawa, M. and Kowallik, K. V. (1998). Gene transfer to the nucleus and the evolution of chloroplasts. Annu. Rev. Genet. 393,162 -165.
Matton, D. P., Maes, O., Laublin, G., Xike, Q., Bertrand, C.,
Morse, D. and Cappadocia, M. (1997). Hypervariable domains of
self-incompatibility RNases mediate allele-specific pollen recognition.
Plant Cell 9,1757
-1766.
McFadden, G. I. (1999). Plastids and protein targeting. J. Eukaryot. Microbiol. 46,339 -346.[Medline]
Morden, C., Delwiche, C., Kuhsel, M. and Palmer, J. (1992). Gene phylogenies and the endosymbiotic origin of plastids. Biosystems 28,75 -90.[CrossRef][Medline]
Morse, D., Salois, P., Markovic, P. and Hastings, J. W. (1995). A nuclear encoded form II rubisco in dinoflagellates. Science 268,1622 -1624.[Medline]
Nassoury, N., Fritz, L. and Morse, D. (2001).
Circadian changes in ribulose-1,5-bisphosphate carboxylase/oxygenase
distribution inside individual chloroplasts can account for the rhythm in
dinoflagellate carbon fixation. Plant Cell
13,923
-934.
Osafune, T., Yokota, A., Sumida, S. and Hase, E. (1990). Immunogold localization of ribulose-1,5-bisphosphate carboxylase with reference to pyrenoid morphology in chloroplasts of synchronized Euglena gracilis cells. Plant Physiol. 92,802 -808.
Palmer, J. D. and Delwiche, C. F. (1996).
Second-hand chloroplasts and the case of the disappearing nucleus.
Proc. Natl. Acad. Sci. USA
93,7432
-7435.
Rowan, R., Whitney, S. M., Fowler, A. and Yellowlees, D.
(1996). Rubisco in marine symbiotic dinoflagellates: form II
enzymes in eukaryotic oxygenic phototrophs encoded by a nuclear multigene
family. Plant Cell 8,539
-553.
Saba-El-Leil, M., Rivard, S., Morse, D. and Cappadocia, M. (1994). The S11 and S13 self incompatibility alleles in Solanum chacoense Bitt. are remarkably similar. Plant Mol. Biol. 24,571 -583.[Medline]
Schiff, J. A., Schwartzbach, S. D., Osafune, T. and Hase, E. (1991). Photocontrol and processing of LHCP II apoprotein in Euglena: possible role of Golgi and other cytoplasmic sites. J. Photochem. Photobiol. B 11,219 -236.[CrossRef][Medline]
Sogin, M. L., Morrison, H. G., Hinkle, G. and Silberman, J. D. (1996). Ancestral relationships of the major eukaryotic lineages. Microbiologia 12, 17-28.[Medline]
Strub, A., Lim, J. H., Pfanner, N. and Voos, W. (2000). The mitochondrial protein import motor. Biol. Chem. 381,943 -949.[Medline]
Sulli, C. and Schwartzbach, S. D. (1996). A
soluble protein is imported into Euglena chloroplasts as a membrane-bound
precursor. Plant Cell 8,43
-53.
Sulli, C., Fang, Z., Muchhal, U. and Schwartzbach, S. D.
(1999). Topology of Euglena chloroplast protein precursors within
endoplasmic reticulum to Golgi to chloroplast transport vesicles.
J. Biol. Chem. 274,457
-463.
van Dooren, G. G., Schwartzbach, S. D., Osafune, T. and McFadden, G. I. (2001). Translocation of proteins across the multiple membranes of complex plastids. Biochim. Biophys. Acta 1541,34 -53.[CrossRef][Medline]
Vothknecht, U. C. and Soll, J. (2000). Protein import: the hitchhikers guide into chloroplasts. Biol. Chem. 381,887 -897.[Medline]
Waller, R. F., Reed, M. B., Cowman, A. F. and McFadden, G.
I. (2000). Protein trafficking to the plastid of Plasmodium
falciparum is via the secretory pathway. EMBO J.
19,1794
-1802.
Wastl, J. and Maier, U. G. (2000). Transport of
proteins into cryptomonads complex plastids. J. Biol.
Chem. 275,23194
-23198.
Wilson, R. J., Denny, P. W., Preiser, P. R., Rangachari, K., Roberts, K., Roy, A., Whyte, A., Strath, M., Moore, D. J., Moore, P. W. et al. (1996). Complete gene map of the plastid-like DNA of the malaria parasite Plasmodium falciparum. J. Mol. Biol. 261,155 -172.[CrossRef][Medline]
Yoon, H. S., Hackett, J. D. and Bhattacharya, D.
(2002). A single origin of the peridinin- and
fucoxanthin-containing plastids in dinoflagellates through tertiary
endosymbiosis. Proc. Natl. Acad. Sci. USA
99,11724
-11729.
Zhang, Z., Green, B. R. and Cavalier-Smith, T. (2000). Phylogeny of ultrarapidly evolving dinoflagellate chloroplast genes: a possible common origin for sporozoan and dinoflagellate plastids. J. Mol. Evol. 51, 26-40.[Medline]