Upload
lengoc
View
221
Download
0
Embed Size (px)
Citation preview
REVIEW
Chloroplast Translation: Structural and Functional Organization,
Operational Control, and Regulation
Reimo Zoschke* and Ralph Bock*
Max Planck Institute of Molecular Plant Physiology, 14476 Potsdam, Germany
* To whom correspondence should be addressed. Email: [email protected] or
Short title: Chloroplast Translation
One-sentence summary:
We review the structure of the plastid ribosome, the mechanisms of chloroplast protein biosynthesis, the control and regulation of plastid translation, and the methods available to study translation.
ABSTRACT
Chloroplast translation is essential for cellular viability and plant development. Its positioning at the intersection of organellar RNA and protein metabolism makes it a unique point for the regulation of gene expression in response to internal and external cues. Recently obtained high-resolution structures of plastid ribosomes, the development of approaches allowing genome-wide analyses of chloroplast translation (i.e., ribosome profiling), and the discovery of RNA-binding proteins involved in the control of translational activity have greatly increased our understanding of the chloroplast translation process and its regulation. In this review, we provide an overview of the current knowledge of the chloroplast translation machinery, its structure, organization, and function. In addition, we summarize the techniques that are currently available to study chloroplast translation and describe how translational activity is controlled and which cis-elements and trans-factors are involved. Finally, we discuss how translational control contributes to the regulation of chloroplast gene expression in response to developmental, environmental, and physiological cues. We also illustrate the commonalities and the differences between the chloroplast and bacterial translation machineries and the mechanisms of protein biosynthesis in these two prokaryotic systems.
Keywords: ribosome, chloroplast, plastid, gene expression, translational regulation, translational control, translation initiation, translation elongation, translation termination
Plant Cell Advance Publication. Published on April 2, 2018, doi:10.1105/tpc.18.00016
©2018 American Society of Plant Biologists. All Rights Reserved
2
INTRODUCTION
Chloroplasts are the characteristic organelle of plant cells. They host numerous essential metabolic
pathways including photosynthesis, which makes chloroplasts the primary source of chemical energy on
earth. All chloroplasts are likely derived from a single ancient photosynthetic cyanobacterium that was
engulfed by a mitochondriate eukaryotic cell more than a billion years ago. During subsequent host-
endosymbiont co-evolution, the genome of the endosymbiont shrank significantly (e.g., Timmis et al.,
2004). While some genes were lost, many others were transferred to the host genome (Martin et al., 2002;
Bock and Timmis, 2008). The proteome of today’s chloroplasts consists of ~3000 proteins, most of which
are nucleus-encoded and post-translationally imported into the organelle.
Present-day chloroplasts still harbor a genome, which comprises ~120 genes in green plants. Most plastid
genes are essential for plant viability because they encode crucial components of the photosynthesis
machinery (the large subunit of Rubisco and approximately half of the subunits of the thylakoidal protein
complexes involved in the light reactions: photosystem I and II [PSI and PSII], cytochrome b6f complex
[Cyt b6f] and ATP synthase) and the gene expression system of the plastid (including a complete set of
bacterial-type RNA polymerase core subunits, rRNAs and tRNAs, and approximately one third of the
ribosomal proteins; Allen et al., 2011; Green, 2011). The reasons for retention of this particular set of
genes in the plastid genome are not fully understood. Several hypotheses that are not necessarily mutually
exclusive have been put forward, including constraints on importability of proteins (and RNAs) into
plastids and requirements for efficient and organelle-specific redox-regulation of gene expression (Allen,
2015). An intriguing consequence of endosymbiont-host co-evolution are multimeric chloroplast protein
complexes, whose subunits are encoded in different compartments (i.e., the plastid and the nucleus). This
necessitates the tight orchestration of nuclear and chloroplast gene expression (e.g., Jarvis and Lopez-
Juez, 2013; Kleine and Leister, 2016).
The bacterial origin of chloroplast gene expression is evident, for example, from the operon-like structure
of plastid gene clusters and the highly similar composition of the translation machinery. However, several
features clearly distinguish chloroplast gene expression from that of bacteria. For example, chloroplasts
possess an astoundingly complex RNA metabolism that includes the usage of different RNA polymerases
and extensive post-transcriptional RNA processing by splicing, editing, end processing, and intercistronic
processing of polycistronic RNAs (Barkan, 2011; Lyska et al., 2013; Börner et al., 2015). These processes
are nearly exclusively conducted by nucleus-encoded protein factors, most of which were likely
established during host-endosymbiont co-evolution (Barkan, 2011; Lyska et al., 2013; Pfalz and
Pfannschmidt, 2013). Also different from bacteria, the regulatory influence of transcription is limited and
3
post-transcriptional and translational events represent key points in controlling chloroplast gene
expression (e.g., Barkan, 2011; Sun and Zerges, 2015).
In recent years, numerous specific features of chloroplast translation were uncovered, including its
interconnection with co-translational processes in RNA and protein metabolism, its regulation in response
to internal and external triggers and the presence of unusual components of the translation machinery.
Genome-wide analyses have unraveled the suborganellar localization of translation and its participation in
controlling the developmental program of chloroplast gene expression (Zoschke and Barkan, 2015;
Chotewutmontri and Barkan, 2016). Studies of sequence-specific chloroplast RNA-binding proteins that
comprise helical repeat domains, especially the pentatricopeptide repeat (PPR) proteins, revealed their
concerted function in RNA metabolism and promotion of translation (Barkan and Small, 2014; Hammani
et al., 2014). Last but not least, high-resolution structural analyses provided a detailed three-dimensional
picture of the plastid ribosome (e.g., Graf et al., 2016; Bieri et al., 2017). These discoveries were largely
enabled by novel approaches towards the quantitative transcriptome-wide analysis of translational
activity, the identification of specific factors that control protein synthesis, and the structural elucidation
of the translational apparatus in plastids (e.g., by ribosome profiling techniques, RNA co-
immunoprecipitation assays, and refined methods for 3D structural analysis). In the light of these and
other findings, we now can reevaluate classical models of chloroplast translation and reassess
controversially discussed hypotheses.
Chloroplast translation has been mainly studied in the unicellular green alga Chlamydomonas reinhardtii
and in model seed plants such as Arabidopsis, maize, and tobacco. In this review, we focus on
embryophytes and, wherever appropriate, refer to breakthrough discoveries made in Chlamydomonas. For
a broader overview of chloroplast translation in Chlamydomonas, the interested reader is referred to
comprehensive review articles (Stern et al., 2009; Nickelsen et al., 2014; Sun and Zerges, 2015).
METHODS TO ANALYZE CHLOROPLAST TRANSLATION
Methods to determine translational activity either indirectly examine the ribosome coverage of the
translation template (i.e., the mRNA), or directly measure the accumulation of newly synthesized
proteins.
Classical methods to analyze translation
Pulse labeling is the method of choice to directly measure translational activity in vivo. In this approach,
isolated chloroplasts, cells or intact plant tissues are fed with the 35S radiolabeled amino acids methionine
and/or cysteine (e.g., Barkan, 1998). The isotopes are incorporated into newly synthesized proteins to an
4
extent that mirrors their synthesis rate. Subsequently, the radiolabeled proteins can be separated by gel
electrophoresis, visualized and quantified (Figure 1A). An advantage of pulse labeling is that it has the
potential to measure protein synthesis rates independent of the dynamics of ribosome movement along the
mRNA. By pulse labeling, the synthesis rate of especially the large plastid-encoded core subunits of the
photosynthesis machinery can be readily quantified. However, the method has several limitations: (i)
Small subunits and subunits of similar molecular weight are difficult to resolve in protein gels and may
require selective purification by immunoprecipitation (Barkan, 1998). (ii) The synthesis rates of many
plastid-encoded proteins cannot be determined by pulse labeling approaches due to their low expression
levels. (iii) The measured quantities of labeled proteins are determined by their synthesis and degradation
rates. Consequently, for proteins with high turnover rates (e.g., PsbA, the D1 protein of PSII), results of
pulse-labeling experiments are often difficult to interpret (even if followed by a chase with unlabeled
amino acids to examine the stability of the labeled protein). Also, (iv) in multicellular organisms, neither
the pulsing nor the chasing occur homogenously in all cells, thus making quantitative comparisons very
challenging.
Polysome analysis is a widely used method that indirectly measures translational activity using the
association of mRNAs with ribosomes as a proxy. Polysomes are high molecular weight assemblies of
actively translating ribosomes held together by the strands of mRNA being translated. They can be
separated from free mRNAs and ribosomes (monosomes) by ultracentrifugation in sucrose density
gradients (Figure 1B). Following RNA extraction from gradient fractions and RNA gel blot analysis, the
distribution of specific mRNAs across the gradient is visualized and provides a qualitative measure of
their translational activity (Barkan, 1998). For genome-wide analysis (translatomics), mRNAs recovered
from different density fractions can be examined by microarray hybridization (e.g., Kahlau and Bock,
2008). Polysome analyses are often complicated by the operon-like organization of genes in the plastid
genome. The processing of transcripts produced from polycistronic transcription units frequently gives
rise to a multitude of mono-, oligo- and polycistronic RNA species, all of which represent potential
translation templates (e.g., Barkan, 1988). Due to the physical linkage of reading frames located on the
same transcript, their individual translation rates cannot be resolved. In addition, only translational
regulation at the level of initiation can be detected, because the molecular weight of mRNAs loaded with
actively elongating ribosomes is indistinguishable from those with paused or stalled ribosomes.
Other elegant though labor intensive methods have been used to examine the regulatory capacity of cis-
elements in chloroplast translation: (i) Chloroplast in vitro translation systems have been established and
used to analyze the regulatory influence of putative cis-elements residing in the 5’ untranslated region
(5’UTR) on translation (e.g., Hirose and Sugiura, 1996; Yukawa et al., 2007). (ii) Reporter genes (e.g.,
A Pulse labeling
3s5_1abeledcysteine/methionine
• + ••
Cell lysis
+
Proteins \\ � J'-� 1D or 2D gel electrophoresis
Labeled Proteins
+
1- PsaA/B-AtpA/B ( PsbB (CP47)
1
_ PsbC (CP43)
1- PsbD (D2)- PsbA (D1)
B Polysome analysis
Polysomes, monosomes, free mRNA
Cell lysis
+ ;-e5
�_rJ
Ultracentrifugation
Separated RNPs
Sucrose gradien
Free mRNA
+ Polysomes
RNA purification and detection
RNA gel blot
+
---�-1
Figure 1. Common methods to analyze chloroplast translation.
C Ribosome profiling
Polysomes, monosomes, free mRNA
Cell lysis
+ ;-e5
�_rJ
Nuclease treatment
Monosomes
Ribosome footprints
+ B B
B B
RNA purification
+
NGS or microarray analysis
Ribosome distribution
(A) Pulse labeling. Plant cells (chloroplast: large green oval, nucleus: white circle) are fed with radiolabeled cysteine and/ormethionine (red dots) which is incorporated together with unlabeled amino acids (black dots) into nascent peptides bytranslation (for simplicity, only chloroplast ribosomes are shown). Proteins are then isolated, separated by gel electrophoresisand visualized/quantified by radio-detection methods (gel picture kindly provided by Karin Meierhoff).
(B) Polysome analysis. Plant cell lysates are loaded on sucrose gradients (white to black: low to high concentration) toseparate ribonucleoprotein particles (RNPs) according to their molecular weight by ultracentrifugation. RNA is isolated fromgradient fractions and examined by RNA gel blot analysis to determine the ribosome loading of specific mRNAs.
(C) Ribosome profiling. Plant cell lysates are treated with nuclease to degrade ribosome-free mRNA sequences. Thisgenerates monosomes, whose protected mRNA fragments (ribosome footprints) are subsequently purified. The positions andabundances of the ribosome footprints are determined by next-generation sequencing or microarray hybridization and reflectprotein synthesis rates.
5
6
GFP or GUS) have been fused to different presumed cis-elements and inserted into the plastid genome by
chloroplast transformation to examine the translational activity conferred by these sequences (e.g., Staub
and Maliga, 1994; Eibl et al., 1999; Drechsel and Bock, 2011).
Ribosome profiling: genome-wide analysis of translation at high resolution
The above described classical methods have been informative, but they are labor intensive, limited in
resolution, and none of them is suited to genome-wide and/or high-throughput analyses. These deficits
were addressed by ribosome profiling, an approach that enables the quantitative genome-wide analysis of
translation in unprecedented depth and resolution (Ingolia et al., 2009). Ribosome profiling takes
advantage of the remarkable stability of translating ribosomes, which protect the mRNA sequence they
physically cover from attack by nucleases, thereby producing protected fragments, so-called ribosome
footprints (Wolin and Walter, 1988; Figure 1C). Next-generation sequencing analysis of these footprints
determines the in vivo positions and abundances of translating ribosomes. Considering that each
elongating ribosome produces one protein, ribosome footprint abundances reflect the protein synthesis
rate for each reading frame (Ingolia et al., 2009). Footprint abundance is typically normalized to mRNA
abundance (assayed by RNA sequencing), so that relative translation efficiencies can be inferred (Ingolia
et al., 2009). Consequently, the approach measures the two determinants of gene expression that define
the final protein output: transcript amount and translational activity. In recent years, ribosome profiling
has been extensively used to study translation in prokaryotes and eukaryotes (e.g., Ingolia, 2016).
In chloroplasts, ribosome profiling was first applied in a modified approach, exchanging the next-
generation sequencing analysis of footprints by microarray hybridization (Zoschke et al., 2013a; Figure
1C). More recently, deep sequencing was used to study chloroplast translational dynamics in maize and
Arabidopsis (Chotewutmontri and Barkan, 2016; Lukoszek et al., 2016; Gawronski et al., 2018).
Despite the compelling attractions of ribosome profiling, it should be noted that the method cannot
distinguish actively translating from paused ribosomes. This may be problematic if translation is
regulated at the level of elongation, as described for some chloroplast genes (see below). Application of
inhibitors of initiation or early elongation (e.g., lincomycin) and examination of the run-off kinetics of
ribosomes over time should allow to distinguish pausing from elongating ribosomes.
THE PLASTID TRANSLATION MACHINERY – VARIATIONS ON A BACTERIAL THEME
A bacterial-like translation machinery whose components are encoded by two genomes
Chloroplast translation is carried out by prokaryotic-type 70S ribosomes that are composed of a small 30S
and a large 50S subunit and contain orthologues of most proteins and all rRNAs of the Escherichia coli
7
(E. coli) reference ribosome (Kössel et al., 1985; Yamaguchi and Subramanian, 2000; Yamaguchi et al.,
2000). All rRNAs, a complete set of ~30 tRNAs, approximately half of the ribosomal proteins of the 30S
subunit and one quarter of the proteins of the 50S subunit are encoded in the plastid genome (Sugiura,
1995). The remaining ribosomal proteins are nucleus-encoded (Tiller and Bock, 2014;
http://www.bangroup.ethz.ch/research/nomenclature-of-ribosomal-proteins.html). Interestingly, the
genomic distribution of genes for organellar ribosomal proteins is, to some degree, evolutionarily
conserved, suggesting constraints in ribosome assembly that require on-site co-expression of rRNAs and
organelle-encoded core ribosomal proteins (Maier et al., 2013). Most other components of the plastid
translation machinery are nucleus-encoded (e.g., initiation/elongation/termination/ribosome recycling
factors, aminoacyl-tRNA synthetases), except for initiation factor 1 (IF1) which is plastid-encoded in
many plants (Millen et al., 2001).
Plastid deviations from the bacterial ribosome and their structural and functional consequences
Despite the generally bacterial structure of the chloroplast ribosome, there are some features that clearly
distinguish plastid ribosomes from the E. coli reference ribosome. Chloroplast ribosomes contain the full
set of bacterial rRNAs (23S, 16S, and 5S rRNAs), which comprise the peptidyl transferase activity (23S
rRNA) and the decoding center (16S rRNA) and serve as scaffold for ribosomal proteins during ribosome
assembly (Shajani et al., 2011; Maier et al., 2013). However, the 23S rRNA gene was split into two genes
in the plastid genome: a large 5’ portion encoded by the 23S rRNA gene and a small 3’ fragment encoded
by the 4.5S rRNA gene (Whitfeld et al., 1978). In addition, the 23S rRNA is post-transcriptionally
processed at two so-called “hidden breaks” into three fragments, whose abundances and precise sizes vary
among species (e.g., ~0.5, ~1.2 and ~1.1 kb from 5’ to 3’ in Arabidopsis plastids). These fragments are
found in the mature 70S ribosome and held together by intermolecular base pairing (Kössel et al., 1985;
Bieri et al., 2017). Despite their general homology, some structural elements of the E. coli 16S and 23S
rRNAs are absent from chloroplasts and, conversely, the chloroplast 23S rRNA contains additional
secondary structures (Kössel et al., 1985). Particularly well conserved are the catalytic domain V in the
23S rRNA, which carries the peptidyl transferase activity, and the anti-Shine-Dalgarno sequence in the
16S rRNA, which is crucial for translation initiation (Scharff et al., 2017).
In the course of evolution, significant changes also occurred in the proteinaceous part of the chloroplast
ribosome. The Rpl25 and Rpl30 proteins were completely lost and, in some species, the bacterial Rpl23
was replaced by its counterpart from the cytosolic 80S ribosome (Bubunenko et al., 1994; Yamaguchi and
Subramanian, 2000). Furthermore, the first complete inventory of plastid ribosomal proteins identified six
proteins that were assumed to lack bacterial orthologues and, consequently, were designated as plastid-
specific ribosomal proteins (PSRP) 1-6 (Yamaguchi and Subramanian, 2000; Yamaguchi et al., 2000).
8
However, later it was shown that PSRP1 is not a genuine ribosomal protein, but the orthologue of the
bacterial cold-shock protein pY, which is associated with the small subunit of the ribosome but not a
structural part of it (Sharma et al., 2007; Sharma et al., 2010). PSRP4 also shows homology to a bacterial
protein: THX, an intrinsic part of the 30S ribosomal subunit in Thermus thermophilus, which so far, has
not been found in other bacteria (Yamaguchi and Subramanian, 2003). At present, PSRP2 and PSRP3 in
the 30S ribosomal subunit, and PSRP5 and PSRP6 in the 50S subunit are considered genuine plastid-
specific ribosomal proteins and, consequently, their renaming to Rps22/23 and Rpl37/38 was suggested
(Bieri et al., 2017). After their discovery, PSRPs were hypothesized to act in light-regulation of
translation (Yamaguchi and Subramanian, 2003; Manuell et al., 2007). Whereas plastid pY may indeed
perform this function (see below), a number of studies have suggested that the major function of PSRPs
lies in the structural compensation of evolutionarily modified rRNA domains (Sharma et al., 2007; Tiller
et al., 2012; Ahmed et al., 2016; Graf et al., 2016; Bieri et al., 2017). However, this role does not
necessarily exclude additional functions in translational regulation.
Some of the conserved plastid ribosomal proteins also exhibit N- or C-terminal extensions (or internal
expansions) compared to their E. coli orthologues (Yamaguchi and Subramanian, 2000; Yamaguchi et al.,
2000). Many of these extensions mediate new interactions with rRNAs or ribosomal proteins and may
structurally compensate for missing or modified rRNA domains (Ahmed et al., 2016; Graf et al., 2016;
Bieri et al., 2017). However, the extensions of some ribosomal proteins (e.g., S2, S18, S21) represent
potential new contact sites with the mRNA and, therefore, were hypothesized to be involved in
translational regulation (Manuell et al., 2007; Sharma et al., 2007; Graf et al., 2016). Notably, several
alterations (e.g., extensions of Rps5 and Rps1 and an insertion in Rps4) narrow the mRNA entry site of
the chloroplast ribosome compared to the E. coli reference ribosome (Bieri et al., 2017). Furthermore,
structural changes in the polypeptide exit tunnel and the tunnel exit site were hypothesized to support the
co-translational binding of the chloroplast signal recognition particle (SRP) which diverges substantially
from that of bacteria (Ahmed et al., 2016; Graf et al., 2016; Bieri et al., 2017).
Altogether, the chloroplast ribosome has a substantially higher protein mass (by ~170 kDa) and a slightly
lower RNA content (by ~0.4 kDa) than the E. coli ribosome, resulting in a considerably increased protein
to RNA ratio (~2:3 compared to 1:3 in E. coli). The partial replacement of rRNA domains by protein
elements in chloroplast ribosomes follows the general evolutionary trend of reducing RNA components in
enzymatically active chloroplast ribonucleoprotein particles (RNPs) and substituting them by protein
constituents (Barbrook et al., 2006). Other examples include the chloroplast tRNA processing enzyme
RNase P that lost its RNA component in the Viridiplantae lineage (Pinker et al., 2013) and the chloroplast
SRP of seed plants, that lacks the SRP RNA (Ziehe et al., 2017). The tendency to lose RNA functions in
9
chloroplasts may be driven by the evolutionary genome reduction and the massive transfer of protein-
coding genes to the nucleus. Whereas proteins can be reimported post-translationally into plastids, this
route seems to be blocked for RNA components, which may have enforced their evolutionary loss,
replacement by proteins or retention in the chloroplast genome (in the case of rRNAs and tRNAs;
Barbrook et al., 2006). Also, it was predicted that ribosome composition (including its relative protein and
rRNA contents) is optimized for the production of the translation machinery itself, a process that strongly
limits cell division rates in prokaryotes (Reuveni et al., 2017). However, in plastids, the synthesis of
ribosomal proteins and rRNAs is partially uncoupled from translation, due to the transfer of genes for
ribosomal proteins to the nucleus and the presence of nucleus-encoded RNA polymerases, both of which
are produced by cytosolic rather than organellar ribosomes. Consequently, the evolutionary constraints on
ribosome composition are somewhat relaxed in organelles, which may have facilitated the observed shifts
in protein-to-rRNA ratios in plastid and mitochondrial ribosomes (Reuveni et al., 2017).
Some ribosomal proteins are dispensable under standard growth conditions
Most constituents of the plastid translation machinery are essential for chloroplast biogenesis and
consequently, for plant viability (Tiller and Bock, 2014). In many species, plastid translation is essential
even under heterotrophic growth conditions (Ahlert et al., 2003; Sosso et al., 2012), presumably due to
the necessity to express a few essential plastid genes such as accD, clpP, ycf1 and ycf2 (Bock, 2007).
However, some ribosomal proteins and tRNAs are non-essential, at least under standard greenhouse
conditions.
Non-essential ribosomal proteins have been identified in both E. coli and plastids. Surprisingly, despite
the shared ancestry of bacterial and plastid ribosomes, the essentiality of ribosomal proteins is not fully
conserved between the two systems (Tiller and Bock, 2014). The nucleus-encoded chloroplast ribosomal
proteins RPL11, RPL24, RPS17, RPS21, PSRP3, PSRP6 and the plastid-encoded Rps15, Rpl33 and
Rpl36 are non-essential (Tiller and Bock, 2014). The phenotypes of viable ribosomal protein mutants
range from wild-type appearance to very strong phenotypes with altered leaf morphology, variegated
leaves, cold-induced bleaching and retarded growth. Non-essential plastid ribosomal proteins have been
speculated to facilitate ribosome assembly and structural integrity, or act in regulation, optimization or
localization of translation (Pesaresi et al., 2001; Yamaguchi and Subramanian, 2003; Tiller et al., 2012;
Tiller and Bock, 2014). However, their distinct molecular functions remain to be elucidated. In bacteria,
ribosomes can differ in their protein composition, for instance, under different growth conditions.
Moreover, there is growing evidence that this ribosome heterogeneity (additionally involving post-
transcriptional and post-translational modifications of rRNAs and ribosomal proteins) creates regulatory
capacity by conferring translational selectivity of ribosomal sub-pools for specific mRNAs (e.g., Sauert et
10
al., 2015; Shi et al., 2017). However, whether non-essential plastid ribosomal proteins can act as
modulators of ribosome affinity to specific sets of mRNAs, is currently unknown.
Translation with a minimal set of tRNAs
Similar to some bacterial species, plastids do not encode the full set of 32 tRNAs that are required to
serve the 61 codons by standard and wobble base pairing between codon and anticodon. For example,
only 30 tRNA genes are present in the plastid genome of most seed plants. Since there is no evidence for
tRNA import from the cytosol, superwobble base pairing has been considered as a mechanism enabling
translation with a reduced set of tRNAs. A systematic reverse genetic screen revealed that the chloroplast
tRNAs trnG-GCC, trnL-CAA, trnS-GGA, trnT-GGU, and trnV-GAC are not essential for plant viability
(Alkatib et al., 2012; and references therein). These non-essential tRNAs decode codons with a
pyrimidine in the third position and, for all of them, essential plastid isoacceptor tRNAs with a uracil (U)
in the first anticodon position exist in plastids. These isoacceptor tRNAs can serve the respective codons
by superwobble base pairing (i.e., the U in the wobble position of the anticodon can pair with all four
nucleotides in third codon position; Alkatib et al., 2012). The diverse phenotypes of these mutants suggest
that superwobble base pairing causes distinct molecular constraints on translation (Alkatib et al., 2012),
possibly explaining the evolutionary conservation of some non-essential tRNAs.
CONSERVATION AND MODIFICATION OF BACTERIAL TRANSLATION MECHANISMS
IN CHLOROPLASTS
The overall structural conservation of essential functional elements of bacterial ribosomes in chloroplasts
is generally assumed to reflect the functional preservation of the bacterial translation mechanisms in
plastids (e.g., Peled-Zehavi and Danon, 2007).
Initiation
The initiation process starts with the contact of a pre-initiation complex consisting of the 30S subunit and
the initiator tRNA (for N-formylmethionine, fMet) to the initiation site in the mRNA. Bacterial initiation
depends on the initiation factors IF1, 2 and 3 which facilitate initiator tRNA binding and ribosome
subunit assembly. Functional chloroplast orthologues were identified for all three IFs (Sijben-Müller et
al., 1986; Campos et al., 2001; Miura et al., 2007; Zheng et al., 2016). Interestingly, many plants contain
two or more paralogous genes for plastid IF3, whose differential expression was proposed to regulate
chloroplast translation initiation (Nesbit et al., 2015). Also similar to the standard bacterial translation
initiation, approximately two thirds of the chloroplast reading frames are preceded by the purine-rich
Shine-Dalgarno sequence (SD) (Shine and Dalgarno, 1974; Scharff et al., 2011). In bacteria, the SD
11
interacts by base pairing with a pyrimidine-rich sequence in the 16S rRNA (the anti-Shine-Dalgarno
sequence, aSD) to ensure proper positioning of the initiation complex at the start codon. The aSD is fully
conserved in plant plastids, and evidence for its functionality has been provided for many genes with SD
(e.g., Kim and Mullet, 1994; Hirose and Sugiura, 2004). Nevertheless, the significance of SD-dependent
initiation in chloroplasts has been questioned (e.g., Fargo et al., 1998). To provide ultimate clarification,
mutations were introduced into the aSD in the tobacco chloroplast genome and shown to cause reduced
translation for many reading frames with upstream SDs, thus confirming the functionality of SD-aSD
interactions (Scharff et al., 2017). On the other hand, roughly a third of the chloroplast genes and also
many bacterial genes do not contain SD sequences (or contain a putative SD but display SD-independent
translation initiation). It was shown that low amounts of mRNA secondary structure around the start
codon facilitate SD-independent translation initiation (e.g., Scharff et al., 2011; Nakagawa et al., 2017;
Scharff et al., 2017). Furthermore, in bacteria, the ribosomal protein S1 preferentially binds
polypyrimidine tracts and acts as an RNA chaperone that unfolds structured regions in mRNAs, thereby
enabling efficient SD-independent translation initiation (Qu et al., 2012). Chloroplast S1 binds RNA with
a preference for adenine or uracil-rich sequences, but whether this supports SD-independent translation
initiation is unknown (e.g., Franzetti et al., 1992; Shteiman-Kotler and Schuster, 2000).
Studies in both bacteria and chloroplasts have pointed to a 5’-to-3’ ribosome scanning mechanism and the
preferential utilization of the 5’-most start codon (e.g., Drechsel and Bock, 2011; Yamamoto et al., 2016).
Moreover, extended interactions of mRNA sequences upstream of the start codon and adjacent to the SD
with bases downstream of the anticodon of the initiator tRNA-fMet and bases next to the aSD,
respectively, have been suggested to facilitate chloroplast translation initiation (e.g., Ruf and Kössel,
1988; Esposito et al., 2003; Kuroda et al., 2007). However, the exact mechanism and the quantitative
contribution of these interactions to the efficiency of translation initiation remain to be elucidated. Similar
to bacteria, the triplets AUG, GUG and UUG can be utilized as start codons in chloroplasts (e.g., Hirose
et al., 1999; Kuroda et al., 2007; Rott et al., 2011; Moreno et al., 2017), with the recognition efficiency of
non-AUG start codons presumably depending on the sequence context (Boeck and Kolakofsky, 1994).
Elongation
After binding of the 50S subunit to the pre-initiation complex, the functional 70S ribosome is completed
and starts moving along the coding sequence of the mRNA to translate it into a polypeptide chain.
Bacterial translation elongation depends on the factors EF-Tu, EF-G and EF-Ts for which conserved
chloroplast orthologues were identified (Breitenberger et al., 1979; Fox et al., 1980; Sreedharan et al.,
1985). Notably, the expression of chloroplast elongation factors is regulated by light and other stimuli
12
(temperature, phytohormones, developmental cues), suggesting involvement of elongation factors in the
regulation of translation (Akkaya and Breitenberger, 1992; Bhadula et al., 2001; Singh et al., 2004;
Albrecht et al., 2006; Liu et al., 2010; Schröter et al., 2010).
Analogous to bacterial gene expression, many chloroplast genes are co-transcribed from operon-like gene
clusters. The resulting polycistronic transcripts contain reading frames that are separated by (often short)
spacer sequences or even overlap by a few nucleotides. In bacteria, translation of adjacent and
overlapping reading frames is frequently coupled in that translation of the second coding sequence
depends on that of the first one (Jackson et al., 2007). In some cases, the strong RNA helicase activity of
the ribosome translating the upstream reading frame is needed to unfold RNA secondary structures that
mask initiation elements (SD and/or start codon) of the downstream reading frame (Jackson et al., 2007).
Recently, also direct coupling of termination on the upstream reading frame (without ribosome
disassembly) with subsequent scanning and re-initiation on the downstream reading frame has been
demonstrated (Yamamoto et al., 2016). In chloroplasts, cases of coupled (ndhC/K, psbD/C) and
uncoupled (atpB/E) translation were identified by in vitro and in vivo analyses, respectively, but the
detailed mechanisms of translational coupling are unknown (Yukawa and Sugiura, 2008; Adachi et al.,
2012; Zoschke et al., 2013a). A recently designed in vivo expression system exploits coupled translation
in chloroplasts and suggests that the RNA helicase function of the ribosome may also mediate
translational coupling in chloroplasts (Martin Avila et al., 2016).
Termination and ribosome recycling
When one of the three stop codons is reached, the orthologous release factors RF1/PrfA (serving UAA
and UAG) and RF2/PrfB1 (serving UAA and UGA) set the synthesized protein free by hydrolysis of the
ester bond (Buckingham et al., 1997; Meurer et al., 2002; Motohashi et al., 2007). Additionally, PrfB1 is
involved in the stabilization of plastid mRNAs containing reading frames with UGA stop codons (Meurer
et al., 2002). PrfB3, a non-functional chloroplast-targeted paralog of PrfB1, lacks domains that are
essential for stop codon recognition and hydrolytic activity of release factors. Remarkably, PrfB3 gained a
new function in transcript stabilization of the petB mRNA (Stoppel et al., 2011). Another release factor,
RF3, facilitates dissociation of RF1 and RF2 from the ribosome in bacteria (Buckingham et al., 1997) and
likely also in chloroplasts (Beligni et al., 2004). In the final step, ribosome recycling factor (RRF; Rolland
et al., 1999), EF-G and IF3 facilitate the release of mRNA and tRNA, and the disassembly of the small
and large ribosomal subunits, thereby recycling them for the next round of translation initiation.
13
PLASTID TRANSLATION IS INTERCONNECTED WITH RNA AND PROTEIN
METABOLISM
Relaxed coupling of translation and transcription
In bacteria, translation initiates and elongates co-transcriptionally, thus ensuring efficient transcription
(e.g., by preventing RNA polymerase backtracking), conferring RNA stability (by translating ribosomes
protecting the mRNA from ribonucleolytic attack), enabling timely translation, and maintaining genome
integrity (e.g., by avoiding extended hybridization of RNA and DNA that would cause collision of the
transcription and replication machineries; e.g., McGary and Nudler, 2013). A similar coupling of
transcription and translation was proposed for chloroplast gene expression based on early electron
micrographs that were interpreted as evidence for ribosomes being associated with nascent transcripts
(e.g., Rose and Lindbeck, 1982; Figure 2). Additional evidence for coupling of transcription and
translation has come from the findings that (i) ribosomal proteins are associated with the transcription
machinery (Pfalz et al., 2006), and (ii) translation factors and other proteins involved in translation are
enriched in plastid nucleoids in a ribonuclease-sensitive manner, suggesting tethering by nascent
transcripts (Majeran et al., 2011). Moreover, orthologues of the Nus proteins that couple transcription and
translation in bacteria have been identified in chloroplast nucleoids (Majeran et al., 2011). However,
chloroplast transcripts have a longer half-life than bacterial transcripts, are stable when not covered by
ribosomes, and many translated RNA species are generated by RNA processing (see below). Together,
this implies that, simply due to the kinetics of mRNA processing and turnover, there may be a
quantitative shift towards post-transcriptional translation in chloroplasts.
The chloroplast genome is regularly transcribed by bacterial-type and phage-type RNA polymerases that
have very different properties (Börner et al., 2015). For example, the two polymerases transcribe with
different speeds and recognize different promoters thus producing primary transcripts with divergent 5’
ends. Notably, in phage-infected E. coli cells, the speedy transcription by the RNA polymerase of
bacteriophage T7 is not coupled with translation and thus produces initially “naked” (i.e., ribosome-free)
transcripts with a higher decay rate (e.g., Makarova et al., 1995). How transcription by the different
plastid RNA polymerases is coordinated with translation and whether or not the utilized RNA polymerase
influences the kinetics of protein synthesis, is currently unknown.
Unprocessed transcripts can be translated
Primary chloroplast transcripts undergo extensive RNA processing, including splicing of group I and II
introns, RNA editing (changing cytosine to uracil residues to restore codons for conserved amino acids or
start or stop codons), 5’ and 3’ end trimming, and intercistronic processing that generates diverse
-
Thylakoids
Internal triggers:
- developmental stage
- cell type
- phytohormones- redox state
- circadian clock- alarmone - (p)ppGpp- metabolites
Unprocessed RNA
Transcription
External triggers:
- light (quality/quantity)- temperature
- osmotic status- nutrient availability
Processed RNA
R:licing,� Protein proce�sing,e�iting,_ intra: & �����11;
1targeting,
1nterc1stronic Y processing
Chloroplast ----i Protein
___ _, complex
DNA
Thylakoids
RNA polymerase
RNA with reading frame, --�-- and translating ribosome
with nascent peptide
Group II intron
Transcript-specific RNA-binding proteins (or complexes) involved in RNA processing and/or translational activation
Factors involved in protein processing, folding, targeting and assembly
Membrane-embedded protein import
apparatus (TOC/TIC)
Figure 2. Overview of internal and external triggers that cause regulatory adjustments of translation in chloroplasts, the mechanisms that control translation, the coupling of RNA and protein metabolisms to chloroplast translation, and the localization of the chloroplast translation machinery.
Chloroplast translation is regulated in response to internal and external triggers (listed in the upper part). Nucleus-encoded factors are translated in the cytosol (shown in the upper left part) and imported into the chloroplast, where they control and/or regulate chloroplast protein synthesis directly (by altering chloroplast translation activity) or indirectly (by controlling co-translational chloroplast RNA or protein metabolisms). Chloroplast translation occurs co-transcriptionally (left), however, due to the slow mRNA turnover, the majority of ribosomes act post-transcriptionally (right). RNA-binding proteins assist co-translational RNA processing and/or facilitate translation initiation. Ribosomes initiate and elongate regularly on both processed and unprocessed transcripts, the extent of which seems to mainly depend on the kinetics of the processing events (see text and Figure 3). Many of the factors involved in protein processing, folding, targeting and assembly act co-translationally on the nascent polypeptide. See text for details.
14
15
transcript isoforms from polycistronic primary transcripts (e.g., Barkan, 2011; Lyska et al., 2013). There
is no obvious spatial separation that would compartmentalize RNA metabolism and translation.
Interestingly, many of the factors known to be involved in transcript processing (e.g., RNA-binding
proteins), stabilization and translation co-localize with processed and unprocessed transcripts in nucleoids
and transcription complexes (e.g., Pfalz et al., 2006; Majeran et al., 2011; Lehniger et al., 2017). This
raises the question whether unprocessed mRNAs, often referred to as “precursors” or “immature
transcripts”, are utilized as templates for translation, or whether any partitioning (e.g., temporal
separation) exists between RNA processing and translation in plastids.
Up to twelve reading frames are interrupted by group II introns in chloroplast genomes of seed plants
(e.g., Sugiura, 1995). In all of them, a substantial fraction of the coding region is located downstream of
the intron and, consequently, splicing is essential to produce functional proteins (Barkan, 2011).
Surprisingly, recent ribosome profiling studies in maize chloroplasts have demonstrated that translation
initiates on unspliced atpF, ndhA, ndhB and ycf3 transcripts and also elongates (Zoschke et al., 2013a;
Alice Barkan, personal communication; Figure 3A). Whether translation elongation pauses at the robust
intron structure or, alternatively, the RNA helicase function of the ribosome allows translation to proceed
into the intron until it terminates at the first in-frame stop codon, thus producing non-functional proteins,
has not yet been possible to resolve. Nevertheless, it is clear that ribosomes initiate and elongate on
unspliced transcripts, strongly arguing against a spatial or temporal separation of splicing and translation
processes.
High-resolution ribosome profiling analysis of maize chloroplast translation also demonstrated that the
degree of editing in mRNA footprints of actively translating ribosomes is not substantially different from
that in the general transcript pool (Chotewutmontri and Barkan, 2016). Specific editing sites that are only
partially edited in the transcript pool showed a similar degree of partial editing in ribosome footprints.
This is in line with the earlier finding that editing of tobacco rps14 is not a requirement for efficient in
vitro translation (Hirose et al., 1998). Together, these data imply that ribosomes cannot distinguish
between edited and unedited transcripts (Figure 3B). However, two exceptions were identified in maize:
the transcripts of rpl2 and ndhA which are only translated in their edited form (Chotewutmontri and
Barkan, 2016). In rpl2, ACG-to-AUG editing restores the start codon (Hoch et al., 1991), thereby
activating translation (Chotewutmontri and Barkan, 2016). This is consistent with the earlier finding that
restoration of the tobacco ndhD start codon by editing is essential for efficient translation initiation in
vitro (Hirose and Sugiura, 1997). In some instances, start codon restoration by editing is subject to tissue-
specific or developmental variation, thus raising the intriguing possibility that editing may control
translational activity (Ichinose and Sugita, 2016). In the case of ndhA, splicing of its group II intron is
A Unspliced
5' 3'
Exon 2
Unedited
5' 3'
C
Polycistronic
RF 1 RF 2 RF 3
Processing
\ •
w
Spliced
5'
�
3'
Edited
5' 3'
Monocistronic
5'�3' 5'�3' RF 1 RF 3
5'�3 RF 2
Figure 3. Ribosomes translate unprocessed chloroplast transcripts (see text for details).
(A) Several chloroplast reading frames are interrupted by group II intrans. Left: Translating ribosomes cover exon 1 ofunspliced atpF, ndhA, ndhB, and ycf3 transcripts (Zoschke et al., 2013a; Alice Barkan, personal communication). Middle andright: Splicing releases the intron and ligates the exons. Consequently, both exons of the spliced transcript are occupied byribosomes, producing full-length proteins (chain of black dots: nascent polypeptide).
(B) Chloroplast transcripts are edited at specific sites by modification of cytosine (C) to uracil (U) residues, often restoringcodons for conserved amino acids (change from yellow to white dot in the nascent peptide). Actively translated mRNAs havethe same editing status as the total transcriptome (Chotewutmontri and Barkan, 2016), indicating that, in a partially editedtranscript pool, also unedited transcripts are translated.
(C) Polycistronic chloroplast transcripts often undergo post-transcriptional processing that generates smaller transcriptisoforms (represented by the three monocistronic transcripts on the right; RF : reading frame). Often all transcript isoforms areused as translation templates. The extent to which transcript processing may enhance translation efficiency needs to bedetermined on a case-by-case basis.
16
17
required to enable editing of the first editing site in exon II (which is separated by the intron; Schmitz-
Linneweber et al., 2001; and references therein). Consequently, ndhA transcripts that are unedited at this
particular site are unspliced, and elongating ribosomes cannot get through to the editing site in the second
exon (Chotewutmontri and Barkan, 2016). Leaving aside these special cases, it can be assumed that,
normally, unedited and unspliced transcripts are translated (Figure 3) and potentially give rise to the
synthesis of low amounts of non-functional and potentially deleterious proteins. In accordance with this
assumption, failure to restore conserved amino acids in subunits of PSII, the cytb6f complex and the ATP
synthase by mRNA editing strongly impairs the function of the respective complexes (Bock et al., 1994;
Zito et al., 1997; Schmitz-Linneweber et al., 2005b). Hence, rapid proteolytic removal of the
dysfunctional proteins synthesized from unedited and unspliced transcripts has to be assumed to prevent
deleterious effects on chloroplast function.
End trimming and intercistronic processing of plastid transcripts have been proposed to enhance their
translational activity (e.g., Drechsel and Bock, 2011). However, internal reading frames on polycistronic
transcripts derived from the maize psbB-psbT-psbH-petB-petD transcription unit are actively translated,
despite the fact that they exist also as 5’ reading frames on processed transcripts (Barkan, 1988).
Likewise, unprocessed tobacco atpH and rbcL mRNAs were translated as efficiently as processed
mRNAs in vitro (Yukawa et al., 2007). Furthermore, synthetic transcription units that were engineered
into plastids gave rise to polycistronic transcripts that were efficiently translated in the absence of
processing (e.g., Staub and Maliga, 1995). Finally, a genome-wide ribosome profiling study revealed that
several polycistronic mRNAs are efficiently used as translation template, despite the known co-existence
of monocistronic transcript isoforms (Zoschke and Barkan, 2015). In all of these examples, the translation
of downstream reading frames in polycistronic mRNAs was not dependent on transcript processing into
monocistronic units, indicating that internal start codons can be efficiently recognized. Also, a number of
plastid reading frames are only present in di- or polycistronic transcripts, indicating that these must
undergo translation. Altogether, these data provide compelling evidence that transcript end processing and
intercistronic processing are not general requirements for efficient translation (Figure 3C). However, in a
few cases there is good evidence that transcript processing stimulates translation. For instance, in tobacco,
a base-pairing interaction between the psaC coding region and the ndhD 5’UTR in the dicistronic
transcript was shown to prevent efficient ndhD translation in vitro, whereas processed monocistronic
transcripts were translationally active, suggesting that, in this operon, processing is required to activate
translation (Hirose and Sugiura, 1997). Similarly, in vitro translation provided evidence that unprocessed
atpB, psbB and psbD transcripts from tobacco chloroplasts are less efficiently translated than their
processed isoforms (Yukawa et al., 2007; Adachi et al., 2012). Moreover, in tobacco chloroplasts,
18
heterologous expression of GFP from engineered polycistronic mRNAs was more efficient when GFP
was placed at the 5’ end of the synthetic operon (Drechsel and Bock, 2011). Finally, in maize, the
monocistronic forms of psaI and rps14 show the highest accumulation in those developmental stages
where the highest translation rates of these reading frames occur, a finding that would be consistent with
the monocistronic RNA species being better translatable (Chotewutmontri and Barkan, 2016).
In sum, although in some cases, translation is indeed stimulated by RNA processing, there is no general
dependence of translation on processing. Unprocessed, unspliced and unedited transcripts have been
shown to be used as translation templates (Figure 3), and, therefore, these transcript isoforms are not
necessarily “immature” or “precursors”.
Several RNA-binding proteins act dually in transcript processing/stabilization and promotion of
translation
In recent years, plastid RNA-binding proteins, many of them with helical repeat domains, were shown to
be involved in specific RNA end trimming and intercistronic processing events (Barkan and Small, 2014;
Hammani et al., 2014). Mutants of some of these factors (Table 1) displayed defects in RNA processing
that were accompanied by translation deficiencies (e.g., Barkan et al., 1994; Felder et al., 2001;
Hashimoto et al., 2003). Initially, these were interpreted as RNA processing-dependent translation defects
(in that processing was required for efficient translation). However, later, it was observed that the
knockout of PPR10, an RNA-binding protein involved in the processing and stabilization of specific atpH
transcript isoforms, caused much stronger translation and protein accumulation defects than expected
from its RNA processing defect, suggesting a more direct role of PPR10 in translation of atpH (Pfalz et
al., 2009). In vitro assays showed that PPR10 binds to the atpH 5’UTR and protects transcripts from 5’-
to-3’exonucleolytic degradation. Consequently, the PPR10-binding site defines the 5’ end of these
transcripts (Prikryl et al., 2011). In addition, PPR10 binding remodels the atpH 5’UTR such that an RNA
stem-loop structure that occludes the putative SD sequence of atpH is dissolved and the ribosome-binding
site becomes exposed (Prikryl et al., 2011). This suggests a dual function of PPR10 in RNA stabilization
and stimulation of atpH translation. A similar mode of action was shown, or is discussed, for other
chloroplast RNA-binding proteins such as HCF107, PGR3, CRR2 and CRP1 (see Table 1 and references
therein). The additional translation-promoting function should be independent of processing in that it
should also occur in polycistronic transcripts where the target reading frame is located downstream of
other reading frames. In fact, reanalysis of ppr10, crp1, pgr3 and hcf107 maize mutants by ribosome
profiling revealed substantial translation defects in vivo (and less severe transcript accumulation defects)
for the reading frames downstream of the RNA-binding sites of the respective protein (in atpH, petD,
petL, and psbH expression, respectively; Zoschke et al., 2013a; Alice Barkan, personal communication).
19
In line with a dual function of some RNA-binding proteins, PPR53, a member of the small PPR-SMR
family, was recently described to be involved in both promotion of ndhA translation and
processing/stabilization of transcript isoforms with ndhA as the 5’ reading frame (Zoschke et al., 2016).
These examples support the idea that RNA-binding proteins acting in processing/stabilization of specific
5’ transcript ends can also directly promote translation of the reading frame downstream of their binding
site (Figure 2). Such proteins provide a physical link between RNA metabolism and translation, and
therefore, co-occurrence of intercistronic processing and stimulation of translation does not necessarily
imply a strict requirement of RNA processing to facilitate translation.
Table 1. Factors demonstrated or suggested to facilitate translation of specific transcripts in seed plant plastids. Asterisks (*) indicate an additional function in stabilization of the transcript 5’ end upstream of the reading frame whose translation is stimulated. Question marks (?) denote proposed but experimentally unconfirmed functions in stimulation of translation. SDR: short-chain dehydrogenase/reductase, HAT: half a tetratricopeptide repeat, PPR: pentatricopeptide repeat, SMR: small MutS-related.
Factor Protein domain(s)
Reading frames with translation promoted
Species References for translational function
ATP1 unknown atpB maize McCormac and Barkan, 1999; Zoschke et al., 2013a
ATP4/SVR7 PPR, SMR atpB maize, Arabidopsis
Zoschke et al., 2012; Zoschke et al., 2013b; Zoschke et al., 2013a
CRP1 PPR petA, petD*, psaC maize Barkan et al., 1994; Zoschke et al., 2013a
CRR2 PPR ndhB* Arabidopsis Hashimoto et al., 2003
HCF107 HAT psbH* Arabidopsis Felder et al., 2001; Hammani et al., 2012
HCF152 PPR possibly petB* Arabidopsis Meierhoff et al., 2003
HCF173 atypical SDR psbA* Arabidopsis Schult et al., 2007
HCF244 atypical SDR psbA Arabidopsis Link et al., 2012
PGR3 PPR petL*, ndhA? Arabidopsis Yamazaki et al., 2004; Cai et al., 2011
PPR10 PPR atpH* maize Pfalz et al., 2009; Prikryl et al., 2011; Zoschke et al., 2013a
PPR53 PPR, SMR ndhA* maize, Arabidopsis
Zoschke et al., 2016
20
Notably, a widely employed sequence element in chloroplast biotechnology (IEE, for intercistronic
expression element) that enhances the heterologous expression of reading frames located in polycistronic
transcription units includes the HCF107-binding site (Zhou et al., 2007; Hammani et al., 2012). The
insertion of the IEE between reading frames enhances the accumulation of monocistronic transcripts. This
could be related to the RNA secondary structure formed by the IEE (Zhou et al., 2007), the action of
RNase E (Walter et al., 2010) and/or the binding of HCF107 (Hammani et al., 2012; Legen et al., 2018).
The turnover of plastid transcripts is not determined by their translation status
In E. coli, mRNAs are stabilized by translating ribosomes, presumably by ribosome coverage providing
physical protection from ribonucleases (e.g., Laalami et al., 2014). By contrast, the study of maize and
Arabidopsis mutants with transcript-specific translation defects has revealed that atpB, petA, psaC and
psbA transcripts are stable although their translation and, consequently, their ribosome occupancy was
dramatically reduced (Barkan et al., 1994; McCormac and Barkan, 1999; Link et al., 2012; Zoschke et al.,
2012; Zoschke et al., 2013b). Furthermore, exchange of the canonical AUG start codon by the non-
standard initiation codon GUG or UUG in the atpB, clpP and psbD reading frames in tobacco
chloroplasts diminished translation initiation and protein synthesis but did not destabilize the transcripts
(Rott et al., 2011; Moreno et al., 2017; Mark A. Schöttler, personal communication). Also, mutants with
general impairments in chloroplast translation do not exhibit general transcript accumulation defects (e.g.,
Barkan, 1993; Scharff et al., 2017). Similarly, the treatment of wild-type Arabidopsis plants with
lincomycin, an antibiotic that disturbs 70S translation elongation only at the earliest steps, thus causing
run-off of ribosomes (Kallia-Raftopoulos and Kalpaxis, 1999), did not cause a substantial decrease in the
accumulation of any of the analyzed chloroplast transcripts (e.g., Meurer et al., 2002; Stoppel et al.,
2011). Accumulation of chloroplast transcripts was also not increased after treatment with
chloramphenicol, a 70S elongation inhibitor that arrests ribosomes and inhibits their release, thus
resulting in densely ribosome-covered transcripts (Nierhaus and Wittmann, 1980).
Altogether, the available data demonstrate that chloroplast mRNAs are stable in the absence of translation
and do not require physical protection by ribosomes. This may not be surprising given the evolutionary
switch from a largely transcriptional regulation of gene expression, as found in bacteria, to predominantly
post-transcriptional regulation in chloroplasts (which strongly depends on transcripts with long half-
lives). Chloroplasts harbor many endo- and exoribonucleases that potentially could degrade “naked”
transcripts (e.g., Germain et al., 2013). This implies that non-translated chloroplast mRNAs must be
somehow protected against nuclease attack. A small family of RNA recognition motif (RRM) domain-
21
containing proteins, the chloroplast ribonucleoproteins (cpRNPs), were shown to be involved in different
steps of mRNA metabolism, including transcript stabilization (Ruwe et al., 2011). Taking into account the
high abundancy of these proteins in the chloroplast, their broad RNA-binding activity in vivo (in that they
associate with virtually all mRNAs), and the fact that they are specifically bound to non-polysomal
mRNAs (Nakamura et al., 2001; Kupsch et al., 2012; Teubner et al., 2017), cpRNPs are strong candidates
for providing stability to untranslated mRNAs. Notably, mutants of the cpRNPs CP29A and CP31A show
a conditional cold-sensitive phenotype, presumably caused by a reduction in the stability of many
mRNAs (Kupsch et al., 2012). A possible explanation is a run-off of translating ribosomes in the cold and
subsequent transcript degradation in the absence of stabilizing cpRNPs. A recent structural analysis
revealed a narrowed mRNA entry site of the chloroplast ribosome compared to that of E. coli (Bieri et al.,
2017). It seems tempting to speculate that this is because chloroplast ribosomes, different from their
bacterial counterparts, need to strip off abundant RNA-binding proteins such as cpRNPs when translation
reinitiates and elongates on previously “stored” (i.e., untranslated) mRNAs, or even on normally
translated mRNAs with low initiation rates (resulting in larger ribosome spacing).
In sum, untranslated chloroplast mRNAs are stable and cpRNP binding may protect them against
ribonucleolytic attack, thus causing the observed uncoupling of mRNA stability from translation.
However, a direct functional connection between translational activity, cpRNP (un)binding and mRNA
stability remains to be demonstrated.
Co-translational folding, maturation, targeting and assembly of proteins
In bacteria, several steps in protein metabolism, including proteolytic processing, chemical modification,
co-factor binding, folding, targeting and assembly, can occur co-translationally (e.g., Gloge et al., 2014).
It seems clear that this is also the case in chloroplasts, although the knowledge about the intersections
between plastid translation and protein metabolism is still scarce (e.g., Giglione et al., 2015; Breiman et
al., 2016; Figure 2). Removal of the N-terminal N-formylmethionine often represents the first step of
nascent peptide chain processing in bacteria and chloroplasts. It occurs co-translationally by the
consecutive reactions of peptide deformylase and methionine aminopeptidase (Breiman et al., 2016).
Likewise, the N-terminal signal peptide for thylakoid targeting of PetA (cytochrome f) is co-
translationally cleaved (see below). Another widespread N-terminal modification of the nascent peptide is
the N-α-acetylation of the penultimate amino acid (e.g., Zybailov et al., 2008; Breiman et al., 2016). A
complete list of identified N-terminal processing events in plastid-encoded proteins is provided at
http://www.i2bc.paris-saclay.fr/spip.php?article1261&lang=fr (Breiman et al., 2016).
22
In bacteria and eukaryotes, folding of the nascent peptide chain has been shown to start already in the
ribosome exit tunnel (e.g., Bhushan et al., 2010; Gloge et al., 2014). With dimensions of 10 nm in length
and 1 - 2 nm in width, the 70S ribosome exit tunnel has a sufficient size to shelter 30 - 60 amino acids
(depending on the folding status) and allows the formation of small protein domains consisting of α-
helices (e.g., Holtkamp et al., 2015). Protein folding in the exit tunnel is assisted by the ribosome itself
through interactions of the nascent peptide with the 23S rRNA and ribosomal proteins (e.g., L4, L22 and
L23; Gloge et al., 2014). Given the high conservation of the peptide exit tunnel, co-translational folding is
expected to occur also during chloroplast translation. The ribosome-associated chaperone trigger factor
binds the nascent peptide upon exit from the ribosome and stabilizes it, thus preventing protein
aggregation and assisting co-translational protein folding in bacteria and, most likely, also in chloroplasts
(Breiman et al., 2016; Ries et al., 2017). Subsequently, other chaperones are recruited and take over
(Trösch et al., 2015).
In parallel to folding, co-factors such as chlorophylls, hemes, carotenoids, quinones and metal ions can
associate co-translationally with chloroplast apoproteins (Schöttler et al., 2011; Nickelsen and Rengstl,
2013; Schöttler et al., 2015). Several studies suggest that the plastid-encoded apoproteins of PSI and PSII
must bind chlorophylls co-translationally to ensure faithful complex biogenesis, most likely, because
chlorophyll binding is required for correct protein folding and assembly (e.g., Nickelsen and Rengstl,
2013). This is supported by evidence that chlorophyll stabilizes nascent chlorophyll-binding proteins
(e.g., Mullet et al., 1990; Kim et al., 1994b; Eichacker et al., 1996). Specific pausing sites during psbA,
psaA, psaB, and psaC translation elongation were suggested to facilitate the co-translational binding of
chlorophyll and other co-factors such as pheophytin, quinone, iron sulfur and manganese clusters (Kim et
al., 1991, 1994a; Gawronski et al., 2018). In cyanobacteria, unassembled PsbB and PsbC apoproteins
contain chlorophyll a and β-carotene, suggesting their early co-translational association (Boehm et al.,
2011).
Chloroplasts comprise different suborganellar compartments: stroma, thylakoid membrane, thylakoid
lumen, inner and outer envelope membranes and the intermembrane space. The targeting of some
chloroplast-encoded proteins to the thylakoid membrane has long been recognized to occur co-
translationally (reviewed in Celedon and Cline, 2013; Figure 2). Early on, it was shown that puromycin
treatment (which causes premature translation termination and thereby release of the nascent peptide) also
releases ribosomes from the thylakoid membrane in chloroplasts, suggesting co-translational protein
targeting mechanisms (e.g., Yamamoto et al., 1981). In later studies, chloroplast sub-fractionation
coupled with polysome analysis and pulse labeling studies revealed that polytopic proteins of PSI (PsaA,
PsaB) and PSII (PsbA/D1, PsbB/CP47, PsbC/CP43, PsbD/D2), and the bitopic cytochrome f subunit
23
(PetA) of the cyt b6f complex associate with the thylakoid membrane co-translationally (e.g., Margulies et
al., 1987; Friemann and Hachtel, 1988; Kim et al., 1994b; van Wijk et al., 1996). However, the
interpretation of the results from these experiments was sometimes controversial (e.g., Ibhaya and
Jagendorf, 1984) and complicated by the fact that in chloroplasts, polycistronic transcripts can be used as
translation templates. Consequently, one co-translationally inserted polypeptide produced from a
polycistronic transcript is sufficient to tether all co-transcribed cistrons to the thylakoid membrane. This
difficulty was overcome in a recent study by using ribosome profiling, a method that employs nucleases
to degrade mRNAs in polysomes down to the footprints protected by monosomes (Zoschke and Barkan,
2015). By coupling this approach with fractionation of chloroplasts into thylakoid membranes and stroma,
co-translational membrane insertion could be comprehensively examined at a genome-wide scale. The
study revealed that 19 of the 37 plastid-encoded intrinsic transmembrane domain-containing thylakoid
proteins in maize insert co-translationally into the membrane, and supplied evidence that exposure of the
first transmembrane domain provides the signal and/or the anchor for stable membrane association (with
the sole exception of PetA, as described below). The data suggest a model for ribosome-mediated mRNA
targeting, in which the nascent polypeptide exposed by the first “pioneer” ribosome anchors the
translation machinery together with the translated mRNA at the thylakoid membrane. Continued
translation by the following ribosomes keeps the mRNA tethered to the thylakoid membrane. A similar
model was suggested for the cytosolic ribosomes that are associated with mitochondria and the
endoplasmic reticulum (Jan et al., 2014; Williams et al., 2014). In addition, a recent ribosome profiling
study in Arabidopsis correlated plastid ribosome pausing events with the synthesis and correct integration
of transmembrane domains (Gawronski et al., 2018). Electron microscopic evidence indicates that
membrane-associated chloroplast polysomes are connected with all unstacked thylakoid membrane
regions (i.e., stroma lamellae, grana margins), but not with internal membranes in grana stacks which are
inaccessible due to their tight packing (e.g., Yamamoto et al., 1981). Consequently, once grana stacks are
assembled during chloroplast biogenesis, plastid-encoded grana proteins (mainly PSII subunits) need to
be transported post-translationally from unstacked membrane regions into grana stacks, for example,
during photosystem repair (e.g., Pribil et al., 2014).
The mechanisms involved in suborganellar protein targeting have been best studied for nucleus-encoded
chloroplast proteins that are post-translationally distributed (Celedon and Cline, 2013). Five major
pathways with partially overlapping functions (and some shared components) have been described
(Schünemann, 2007; Celedon and Cline, 2013). The secretory (Sec) pathway and the twin-arginine
translocase (Tat) transport proteins across the thylakoid membrane into the lumen (reviewed in
Schünemann, 2007). The chloroplast signal recognition particle (cpSRP) interacts with the cpSRP
24
receptor cpFtsY and the insertase ALB3 to insert nucleus-encoded light-harvesting complex proteins into
the thylakoid membrane (reviewed in Ziehe et al., 2017). Some proteins apparently insert spontaneously
into the thylakoid membrane, and finally, a recently discovered parallel Sec pathway targets proteins to
the inner envelope membrane (Li et al., 2017b).
Much less is known about the mechanisms involved in co-translational suborganellar targeting of
chloroplast-encoded proteins. Most plastid-encoded proteins are found in either the stroma or the
thylakoid membrane. Plastid-encoded membrane proteins likely utilize one of the above-mentioned
targeting pathways, either co- or post-translationally. So far, the co-translational targeting mechanisms
were elucidated in some detail for only two plastid-encoded proteins: PetA and PsbA. PetA is the only
plastid-encoded protein containing a cleavable signal peptide at its N-terminus which is recognized by
cpSecA. In vitro and genetic data suggest that PetA targeting to the thylakoid membrane occurs co-
translationally (Voelker et al., 1997; Röhl and van Wijk, 2001; and references therein). A recent ribosome
profiling study showed that nascent PetA engages the thylakoid membrane long before its single
transmembrane domain becomes exposed outside the ribosome exit tunnel, thus confirming co-
translational action of cpSecA-dependent targeting in vivo (Zoschke and Barkan, 2015). Only recently,
SecA-mediated targeting in bacteria was demonstrated to also occur co-translationally (Huber et al.,
2017).
Co-translational interaction of PsbA and cpSRP54 has been suggested based on in vitro cross-linking
experiments (Nilsson and van Wijk, 2002; and references therein). However, Arabidopsis mutants lacking
cpSRP54 have very mild phenotypes, arguing against an essential role of cpSRP54 in membrane targeting
of PsbA or any other core subunit of the photosynthesis machinery (Amin et al., 1999; Tzvetkova-
Chevolleau et al., 2007). Nonetheless, cpFtsY, the essential chloroplast SRP receptor homologue that had
previously been implicated in PSII repair (Tzvetkova-Chevolleau et al., 2007; Asakura et al., 2008), was
shown to be associated with nascent PsbA in vitro suggesting a role in co-translational targeting (Walter
et al., 2015). Furthermore, the translocons cpSecY and ALB3 and the multifunctional protein Vipp1, all
of which are essential for thylakoid biogenesis (Sundberg et al., 1997; Roy and Barkan, 1998; Kroll et al.,
2001), appear to interact with nascent PsbA in vitro, suggesting their involvement in co-translational
membrane targeting of PsbA (Zhang et al., 2001; Walter et al., 2015). Recent in vitro data suggest that the
co-translational targeting of PetB (cytochrome b6) also involves the insertase ALB3 (Kroliczewski et al.,
2016).
The assembly of proteins into functional complexes often initiates co-translationally (e.g., Natan et al.,
2017). Biogenesis of the multiprotein complexes in the thylakoid membrane requires the tightly
25
coordinated action of multiple assembly factors that guide the association of plastid-encoded and nucleus-
encoded subunits (e.g., Schöttler et al., 2011; Nickelsen and Rengstl, 2013). Many plastid-encoded
subunits of these complexes are believed to be assembled co-translationally (Figure 2), because
unassembled free subunits are usually condemned to rapid degradation. However, only in few cases,
direct evidence for co-translational assembly has been obtained. The PSII assembly process is best
understood (e.g., Nickelsen and Rengstl, 2013), and several plastid-encoded core subunits of PSII have
been suggested to assemble co-translationally into the complex. During both de novo assembly and repair
of PSII, nascent PsbA subunits are integrated into an early assembly intermediate that contains PsbD
(Müller and Eichacker, 1999; Zhang et al., 1999). Whereas the first and second transmembrane domains
of PsbA only weakly interact with PsbD, a robust association is established after synthesis of the fourth
transmembrane domain (Zhang et al., 1999; Zhang and Aro, 2002). It is tempting to speculate that the
operon-like organization of chloroplast genes enhances the efficiency of co-translational targeting and
assembly (e.g., in the dicistronic psaA/B and psbD/C transcripts), as this was suggested for bacteria
(Natan et al., 2017).
In summary, a multitude of factors act co-translationally as “welcoming committee” for nascent
polypeptides and assist with the amazing metamorphosis of linear amino acid chains into functional
proteins and protein complexes in specific suborganellar locations (Figure 2). We are just beginning to
understand the complex interconnections of the diverse processes involved in co-translational protein
maturation, targeting and assembly, but it is becoming increasingly evident that the ribosome acts as a
central hub in the coordination of these processes.
OPERATIONAL CONTROL AND REGULATION OF CHLOROPLAST TRANSLATION
Upon stress and under changing environmental conditions, the thylakoid membrane system is adjusted to
achieve optimum photosynthetic performance and prevent photo-oxidative damage. These acclimation
responses require integration of multiple external and internal signals (Figure 2), and involve extensive
regulation of chloroplast translation (e.g., Nickelsen et al., 2014; Sun and Zerges, 2015).
Translational control versus regulation
The terms ‘translational control’ and ‘translational regulation’ are sometimes used synonymously.
However, factors controlling translation (e.g., the strength of a ribosome-binding site in the 5’UTR) do
not necessarily also regulate translation (i.e., dynamically change protein synthesis rates in response to
environmental stimuli or developmental programs).
26
The interplay of trans-acting protein factors and cis-acting RNA elements determines the translation
output of chloroplast genes. The functional involvement of these elements in translational control
typically is demonstrated by their genetic manipulation causing altered translational activity. However,
this does not necessarily imply a regulatory function in translation, resulting in a change in the protein
synthesis rate during adaptation processes. In other words, a given factor would have a regulatory
function if it were to become limiting for translation under specific conditions, thus altering protein
synthesis output. According to this definition, a true regulatory function of chloroplast translation factors
has been established only in very few cases in Chlamydomonas (see below). It was proposed that many of
the nucleus-encoded factors involved in chloroplast RNA metabolism simply suppress mutations that
accumulate in the (asexually reproducing) plastid genome over time (Maier et al., 2008; Lefebvre-
Legendre et al., 2014). Similarly, nucleus-encoded translation factors could be needed constitutively to fix
chloroplast mutations at the RNA level (e.g., by resolving secondary structures around ribosome-binding
sites to facilitate translation initiation). However, on an evolutionary time scale, factors that control
translation may also be recruited as true regulators that modulate translation in response to internal and
external triggers.
Translation plays a major role in the control and regulation of chloroplast gene expression
Translation is an extremely resource-consuming process due to the energy and nutrient demands involved
in the assembly of ribosomes, the synthesis of amino acids, the expression, processing and charging of
tRNAs, and the GTP-dependent reactions during initiation and elongation. Dividing bacterial cells use
~50 % of their energy for protein synthesis (Russell and Cook, 1995). Millar and co-workers calculated
the cellular energy budgets used for protein synthesis in Arabidopsis leaves (Li et al., 2017a). Their
estimate is that, dependent on the developmental stage, 13 – 38 % of the cellular ATP is used for protein
synthesis, with plastid translation accounting for ~70 % of the costs of cellular protein synthesis.
Synthesis of RbcL alone accounts for more than 15 % of the cellular ATP equivalents used for protein
synthesis (Li et al., 2017a). In view of the high costs of translation, the rate of protein synthesis is tightly
coordinated to the cellular demands in all domains of life. In addition, translational regulation has several
advantages. As translation is the final synthesis step in gene expression, its regulation (i) can mediate
rapid responses to internal or external stimuli, (ii) most directly affects the protein accumulation levels,
and (iii) can be readily exploited to enhance or attenuate changes in upstream steps of gene expression
(i.e., transcription and transcript accumulation). Importantly, translation can also be dynamically localized
to the site where the synthesized protein is needed, whereas transcription is largely bound to the position
of the genomic DNA. Finally, although in bacterial systems transcriptional co-regulation is easily
achieved by combining genes in operons, this genomic organization is rather static and, in contrast to
27
translationally co-regulated mRNAs (qualifying as ‘regulons’), cannot readily mediate dynamic responses
to different cues (Keene, 2007).
It is generally accepted that chloroplast gene expression is largely controlled and regulated at the post-
transcriptional and, especially, the translational levels, contrasting gene expression in cyanobacteria,
which is mainly transcriptionally regulated. Several lines of evidence led to this conclusion. First,
bacterial transcripts tend to be unstable (with half-lives in the range of minutes), enabling transcriptional
regulation (e.g., Pedersen and Reeh, 1978; Klug, 1993). By contrast, the half-lives of chloroplast
transcripts are in the range of hours or even days, thus disabling fast transcriptional responses (e.g.,
Mullet and Klein, 1987; Klaff and Gruissem, 1991). Second, although chloroplast gene clusters are
reminiscent of bacterial operons, they often are transcribed from different promoters (including operon-
internal promoters) and, frequently, the resulting primary transcripts are further processed into smaller
units (e.g., Barkan, 2011; Lyska et al., 2013; Börner et al., 2015). Also, chloroplast polycistronic
transcription units often comprise functionally unrelated genes (Sugiura, 1995), unlike bacterial operons,
where co-transcription serves to couple the expression of functionally related genes. Third, the translation
of some chloroplast mRNAs coding for core components of the photosynthesis machinery is induced by
light, whereas their mRNA levels remain virtually unaffected (e.g., Berry et al., 1988; Mühlbauer and
Eichacker, 1998). Fourth, translation of many chloroplast mRNAs was shown to be the rate-limiting step
in gene expression in Chlamydomonas (Eberhard et al., 2002), although the situation may be different in
angiosperms (Udy et al., 2012). Moreover, in a regulatory mechanism termed control by epistasy of
synthesis (CES), the translation rate of some subunits of photosynthetic protein complexes is regulated by
the presence or absence of their assembly partners (Choquet and Wollman, 2009; see also below). Fifth,
in recent years, a number of factors required for the translation of specific chloroplast reading frames
were discovered, which may be suggestive of extensive translational regulation (Table 1).
RNA cis-elements controlling translation
RNA sequence or structural elements that are located in cis, typically upstream of the reading frame, play
crucial roles in translational control. A major cis-acting sequence element for translation initiation in
chloroplasts is the SD sequence (Scharff et al., 2017; see above). Recent ribosome profiling studies have
suggested that, in bacteria and chloroplasts, SD-like sequences that are located within reading frames
cause programmed pausing of elongating ribosomes (Li et al., 2012; Zoschke et al., 2013a; Gawronski et
al., 2018). It has been proposed that SD-dependent pausing facilitates co-translational folding and
targeting of nascent polypeptides (Li et al., 2012; Fluman et al., 2014). Additional chloroplast cis-acting
sequence elements for translational control are the binding sites of trans-acting factors that stimulate
translation of specific reading frames (Table 1).
28
The degeneracy of the genetic code offers another elegant means of influencing the translation rate in cis.
Since an organism’s codon usage is usually adapted to the relative abundances of isoaccepting tRNAs, the
choice of synonymous codons can potentially control translation efficiency (e.g., Supek, 2016). Whether
or not this also applies to chloroplasts, is currently controversially discussed (e.g., Sugiura, 2014; Suzuki
and Morton, 2016; Gawronski et al., 2018). Although initial ribosome profiling studies could not confirm
a robust correlation between codon adaptation and translational speed/efficiency, recent methodological
and bioinformatic refinements uncovered the suspected relationship in several organisms (e.g., Dana and
Tuller, 2014; Nakahigashi et al., 2014).
In addition to primary sequence elements, features of mRNA 2D or 3D structure (or lack of structure) can
represent cis-elements that influence the translation process (e.g., Mauger et al., 2013). For example, the
lack of secondary structures at the start codon facilitates efficient SD-independent translation initiation in
bacteria and chloroplasts (Scharff et al., 2011; Scharff et al., 2017). Furthermore, translation initiation in
the chloroplast atpH and psbH mRNAs is stimulated by resolving secondary structures that mask
ribosome-binding site and start codon, respectively (Prikryl et al., 2011; Hammani et al., 2012). A recent
ribosome profiling study also revealed transcriptome-wide correlations between ribosome pausing and
secondary structures in chloroplast mRNAs (Gawronski et al., 2018).
Riboswitches are structured RNA elements that act as sensors for small molecules (metabolites).
Metabolite binding triggers a conformational switch that regulates transcription (typically by inducing
termination or antitermination) or translation (by exposing or sequestering the ribosome-binding site; e.g.,
Serganov and Nudler, 2013). Although riboswitches have not yet been identified in plastids, they have
been exploited as tools to control the translation of transgenes in chloroplast biotechnology (e.g.,
Verhounig et al., 2010). The use of improved algorithms for the computational prediction of riboswitches
(e.g., Philips et al., 2013) should help to clarify whether endogenous riboswitches exist in chloroplast
genomes.
Internal RNA structures within reading frames can adjust the translational speed by decelerating
elongating ribosomes in bacteria and chloroplasts (Mauger et al., 2013; Gawronski et al., 2018), a
mechanism that has been suggested to aid co-translational protein-complex maturation steps. Specific
translational pausing sites have been identified in the chloroplast psbA and atpA mRNAs (Kim et al.,
1991; Kim and Hollingsworth, 1992). However, although these pausing events appear to be influenced by
light or temperature (Kim et al., 1994a; Grennan and Ort, 2007), they could not be assigned to particular
sequences or structural elements of either the mRNA or the nascent polypeptide chain, nor could any
obvious molecular function directly be ascribed to them. In bacteria, structured RNA elements are also
29
involved in programmed frameshifting and translational coupling of neighboring reading frames on the
same transcript (Jackson et al., 2007; Mauger et al., 2013).
Cis-acting elements in the nascent peptide
Translational cis-elements can also be located in the nascent peptide. For example, translation of
consecutive proline codons is complicated due to the physicochemical properties of the secondary amino
acid proline, which make it a weak peptidyl donor and acceptor. Reading of consecutive proline triplets,
therefore, causes translational pausing (Artieri and Fraser, 2014; and references therein). Specific
elongation factors such as EF-P in bacteria facilitate the translation of consecutive proline codons
(Doerfel et al., 2013; Ude et al., 2013). EF-P is conserved in chloroplasts suggesting a similar function
(e.g., Manuell et al., 2007). More than two consecutive proline codons are usually avoided in proteins of
all organisms. For example, the entire tobacco chloroplast genome does not encode a single stretch of
three prolines.
A recent study also provided evidence for other specific peptide motifs (e.g., domains comprised of small
polar residues) inhibiting peptidyl transfer during elongation or peptide release during termination in
bacteria (Woolstenhulme et al., 2013). Furthermore, specific translational pausing events that are induced
by interactions of the prokaryotic ribosome with the nascent peptide have emerged as programmed
switches that regulate translation in response to small metabolites or the availability of protein
translocation factors (Ito et al., 2010). In chloroplasts, ribosome pausing events were correlated with
positively charged amino acids (Gawronski et al., 2018).
Protein factors that control translation in cis or trans
Proteins that bind the mRNA template, the ribosome or the nascent peptide chain can influence
translation. These proteins act either in an mRNA-specific manner or as more general factors that control
translation of many transcripts.
All mRNA-specific chloroplast translation factors characterized to date act in trans by binding the
5’UTRs of one (or a few) mRNAs and promoting translation of the downstream reading frame (see
above; Table 1). More generally acting trans-factors control the translation of several reading frames.
These factors are either genuine or auxiliary constituents of the ribosome, or bind the nascent peptide.
Hence these factors act in cis or trans. Cis-regulatory functions have been suggested for several ribosomal
proteins or ribosome-associated proteins such as PSRP2-6, pY and Rps1 (e.g., Yamaguchi and
Subramanian, 2003; Manuell et al., 2007; Sharma et al., 2010). In bacteria, Rps1 is involved in translation
initiation of specific transcripts and a similar function has been suggested for chloroplast Rps1 (see
30
above). Chloroplasts also contain an orthologue of the bacterial ribosome-associated protein pY that binds
70S ribosomes or 30S subunits under cold-shock conditions, thus inactivating translation and stabilizing
monomeric 70S ribosomes (Vila-Sanjurjo et al., 2004; and references therein). Similarly, it has been
proposed that plastid pY could be involved in the deactivation of translation and the stabilization of
chloroplast ribosomes in the dark (Sharma et al., 2010; and references therein). Also, the expression of
chloroplast translation elongation factors is regulated by light, suggesting their contribution to
translational regulation. However, direct evidence for a regulatory function of chloroplast elongation
factors Rps1 or pY is lacking.
In both bacteria and eukaryotes, the binding of specific protein factors can create subpools of specialized
ribosomes that selectively translate a specific set of mRNAs (e.g., Sauert et al., 2015). More systematic
and quantitative proteomic studies of chloroplast ribosomes will be required to determine whether
specialized ribosomes exist also in plastids.
Translation is regulated at the initiation and elongation levels
In theory, translation can be regulated at any of its three stages: initiation, elongation, or termination (e.g.,
Hershey et al., 2012). However, by far most common is the regulation of initiation. All chloroplast factors
characterized to date that promote translation of specific reading frames act at the level of initiation
(Table 1). However, in some cases, elongation also appears to be regulated. For example, elongation of
psaA, psaB, psbA and rbcL translation is regulated by light, and the synthesis of PsbA was suggested
additionally to depend on the availability of co-factors and assembly partners (e.g., Berry et al., 1988;
Klein et al., 1988b; Mullet et al., 1990; Kim et al., 1994a; Mühlbauer and Eichacker, 1998; Kim and
Mullet, 2003).
Light- and redox-dependent regulation of translation
Light regulation of translation ensures coordination of the major energy-producing process
(photosynthesis) with the major energy-consuming process (protein synthesis) in the chloroplast. In
addition, components of the photosynthetic apparatus (especially PsbA) are damaged by light,
necessitating specific repair synthesis.
The synthesis of several chloroplast proteins was shown to be activated at the translational level in
response to increased light intensity. Upon illumination, translation of rbcL and psbA is activated at the
level of elongation, a mechanism that was proposed to be regulated by the light-dependent generation of a
proton gradient across the thylakoid membrane, the ATP status of the chloroplast and/or redox signal(s)
generated by photosynthetic electron transfer (e.g., Taniguchi et al., 1993; Mühlbauer and Eichacker,
31
1998; Zhang et al., 1999). In line with regulation of elongation, the transfer of amaranth plants into the
dark caused a decline in RbcL protein synthesis that was not accompanied by the loss of rbcL mRNA
association with ribosomes (in polysomes), suggesting that elongating ribosomes paused (Berry et al.,
1988). In addition, psaA and psaB were shown to be light-regulated at the elongation level during de-
etiolation (Klein et al., 1988b). A potential mechanism for the general light-dependent activation of
chloroplast translation elongation could rely on pY (see above) or elongation factor Tu, which was
identified as redox-regulated (Schröter et al., 2010).
During de-etiolation, psbA translation is also stimulated at the level of initiation (e.g., Klein et al., 1988b;
Kim and Mullet, 1994). Evidence currently available suggests that psbA translation for PSII biogenesis is
regulated at the initiation level, whereas PsbA synthesis for PSII repair is regulated at the (presumably
faster responding) elongation level, a model supported also by data from Chlamydomonas (reviewed in
Nickelsen et al., 2014). In seed plants, psbA translation initiation is controlled by specific cis-element(s)
in the 5’UTR (Staub and Maliga, 1994; Hirose and Sugiura, 1996; Eibl et al., 1999) and by the trans-
factors HCF173 and HCF244, two putative oxidoreductases that promote psbA translation cooperatively
(Schult et al., 2007; Link et al., 2012). All of these elements may also play a role in the light-dependent
translation initiation of psbA.
Variations in light quantity and quality can cause imbalances in the activities of PSI and PSII, changing
the redox state of the chloroplast and potentially generating harmful reactive oxygen species (Pfalz et al.,
2012). The chloroplast redox state controls transcriptional and post-transcriptional adaptation responses in
the chloroplast (e.g., Allen and Pfannschmidt, 2000; Rochaix, 2007). In Chlamydomonas, a redox-
dependent regulatory mechanism has recently been suggested for the light-activated translation of psbD
(Schwarz et al., 2012). In this model, the RNA-binding proteins Nac2 and RBP40 form a disulfide bridge-
connected complex in the light which stabilizes psbD mRNA and activates its translation (Schwarz et al.,
2012; and references therein). In the dark, the complex is reduced and disassembles, and, as a result, PsbD
synthesis is inactivated. The electrons are likely provided by the NADPH-dependent thioredoxin
reductase C, an enzyme shown to reduce disulfide bonds in the dark by utilizing NADPH generated in the
oxidative pentose phosphate pathway (Schwarz et al., 2012).
Plants perceive light independently of photosynthesis by photoreceptors that are located outside of
chloroplasts. However, light-induced regulation of translation was also observed in isolated chloroplasts,
arguing against a crucial role of photoreceptors (Sun and Zerges, 2015).
Altogether, our current knowledge about light-induced translational regulation in seed plant chloroplasts
is restricted to very few photosynthesis-related genes, with psbA translation being best studied. However,
32
even for psbA, the underlying mechanisms and the mode of action of the factors involved are not well
understood.
Developmental regulation of translation
Developmental regulation of plastid gene expression is crucial for the differentiation and interconversion
of plastid types. For example, conversion of proplastids to chloroplasts requires establishment of the
thylakoid system and must be tightly coordinated with cell division and the emergence of photosynthetic
tissues from meristems (Jarvis and Lopez-Juez, 2013). The mechanisms involved in chloroplast
differentiation were identified in classical experiments that analyzed the process of de-etiolation. During
de-etiolation, a rapid rise in translational activity was observed by pulse labeling experiments for several
plastid mRNAs encoding subunits of PSI (psaA/B), PSII (psbA/B/C/D) and Rubisco (rbcL; e.g., Klein and
Mullet, 1986, 1987; Kim and Mullet, 2003; Kleffmann et al., 2007). The initial steep increase in
translation of these mRNAs was followed by a slow decline as chloroplast differentiation continued in the
light. Polysome analyses demonstrated that psaA/B, rbcL and psbA transcripts are found in polysomes
throughout the chloroplast differentiation process, suggesting that the regulation of their translation
occurs, at least in part, at the elongation level (Klein et al., 1988b). This conclusion was further
substantiated by the observation that psaA, psbA and rbcL translation initiation remains unaltered during
light-dependent chloroplast differentiation (Kim et al., 1994b; Kim and Mullet, 2003).
There is also evidence for developmentally activated translation initiation of plastid mRNAs. Initiation of
psbA translation is induced during de-etiolation (Eichacker et al., 1992; Kim and Mullet, 1994), and this
induction is likely controlled by cis-elements in the psbA 5’UTR (Staub and Maliga, 1994; and references
therein). It was further proposed that the light-induced synthesis of chlorophyll controls both the
accumulation and the translation of chlorophyll-binding apoproteins of PSI and PSII during de-etiolation
(e.g., Eichacker et al., 1992). However, this conclusion was mainly based on the observed lack of
accumulation of chlorophyll-binding apoproteins in the absence of chlorophyll, as determined by protein
immunoblotting and pulse labeling experiments in vivo and in vitro (e.g., Klein et al., 1988a; Klein et al.,
1988b; Eichacker et al., 1990; Eichacker et al., 1992). Although pulse labeling can reveal protein
synthesis rates, it cannot unambiguously distinguish between the absence of synthesis and rapid
degradation of newly synthesized proteins (especially not for proteins with high turnover rates, such as
PsbA). A recent ribosome profiling analysis of plastid translation in a maize mutant with knocked-out
chlorophyll synthesis showed that the synthesis of plastid-encoded chlorophyll-binding apoproteins is
virtually unaltered in the absence of chlorophyll (Zoschke et al., 2017). Even the co-translational
thylakoid membrane engagement of nascent apoproteins was shown to be independent of chlorophyll
33
synthesis (Zoschke et al., 2017). However, apoproteins were shown to undergo rapid degradation in the
absence of chlorophyll (e.g., Mullet et al., 1990; Kim et al., 1994b; Eichacker et al., 1996).
The above described de-etiolation of seedlings represents a dramatic change in plant development and
reflects the situation during germination, when light-dependent and developmental programs operate
simultaneously. Another useful experimental system to study translational regulation during chloroplast
differentiation is provided by the longitudinal developmental gradient of young leaves in grasses such as
maize, rice or barley. At the leaf base, meristematic, proplastid-containing tissue is found, followed by a
developmental gradient towards the leaf tip which includes etioplasts, differentiating chloroplasts
(etiochloroplasts) and fully differentiated chloroplasts (e.g., Leech et al., 1973). This natural
developmental gradient has been exploited to systematically study the dynamic changes in gene
expression at the level of transcript and protein accumulation (e.g., Barkan, 1989; Baumgartner et al.,
1989; Cahoon et al., 2004; Li et al., 2010; Majeran et al., 2010).
Recently, ribosome profiling enabled the transcriptome-wide examination of plastid translation in the
longitudinal developmental gradient of maize leaves (Chotewutmontri and Barkan, 2016). This
pioneering study comprehensively determined the relative contributions of changing transcript levels and
translational activity to the protein synthesis output (Chotewutmontri and Barkan, 2016). The data
revealed that the synthesis rates of most plastid-encoded proteins increases early in development and
drops later, once the photosynthetic machinery is set up. Strikingly, PsbA and PetD, two proteins that do
not follow this general rule and, instead, display increasing synthesis levels that peak in the latest
examined developmental stage, represent the subunits with the highest turnover rates in their complexes
(Li et al., 2017a), likely explaining their continued production.
In general, the regulation of plastid gene expression during leaf development in maize is achieved by
changes in transcript levels that are often tuned by changes in translation efficiency (Chotewutmontri and
Barkan, 2016). Remarkably, the developmental dynamics in protein synthesis that was discovered
revealed two major regulons that are defined by changes in RNA accumulation and translation efficiency:
early in development, proteins needed for chloroplast gene expression have the highest synthesis output
(e.g., RNA polymerase subunits and ribosomal proteins), whereas in later developmental stages, proteins
required for photosynthesis are extensively synthesized (i.e., subunits of PSII, Cyt b6f, PSI, ATP synthase,
and NDH complexes). To achieve this developmental pattern in plastid protein synthesis, translational
regulation is especially important for polycistronic transcription units that encode proteins of both
regulons. Indeed, in these transcripts (e.g., the psaA/psaB/rps14 transcription unit encoding two PSI
34
subunits and a ribosomal protein), strong differential translational regulation was observed
(Chotewutmontri and Barkan, 2016).
Unfortunately, none of the factors involved in the complex developmental regulation of chloroplast
translation has been identified so far. It would be particularly exciting to reveal the mechanisms
underlying the translational switch between these two major regulons during leaf development. A possible
explanation could lie in the known developmental switch in the usage of the two distinct types of RNA
polymerases during chloroplast differentiation (Börner et al., 2015). In theory, the two polymerase
activities could produce transcripts coding for identical proteins but differing in their translational
competence. Alternatively, RNA-binding proteins that facilitate translation (see above and Table 1) could
mediate the developmental regulation of translation.
Translational autoregulation and feedback regulation
Control of protein synthesis by translational autoregulation or feedback regulation represents an elegant
means of robustly fine-tuning protein synthesis levels to changing cellular requirements. In the
chloroplast of Chlamydomonas, the translation of specific subunits of the four photosynthetic complexes
(PSII, Cyt b6f, PSI, ATP synthase) was shown to be feedback-regulated by the assembly state of the
respective complex, a process termed “control by epistasy of synthesis” (CES; Choquet and Wollman,
2009). CES regulation ensures the stoichiometric production of chloroplast subunits that reside in
oligomeric complexes according to the requirements of their sequential assembly (Choquet and Wollman,
2009). The best-studied case mechanistically is the CES involved in petA gene expression. PetA is a
subunit of the cytochrome b6f complex, whose synthesis is strongly reduced in the absence of its assembly
partners PetB (cytochrome b6) and PetD (subunit IV; Kuras and Wollman, 1994). The protein factors
MCA1 and TCA1 were shown to be cooperatively associated with the 5’UTR of the petA mRNA, where
they stabilize the transcript (MCA1) and promote translation initiation (TCA1; Wostrikoff et al., 2001;
Loiselay et al., 2008). Boulouis et al. discovered a regulation mechanism in which a C-terminal domain of
PetA that is exposed only in the unassembled PetA protein binds to MCA1 and causes its proteolytic
degradation (Boulouis et al., 2011). This leads, in turn, to down-regulation of petA gene expression at
both the level of transcript accumulation and the translational level. Conversely, if the assembly partners
PetB and PetD are available, PetA is assembled into the Cyt b6f complex, the C-terminal domain is
inaccessible, and hence, MCA1 remains stable and facilitates petA expression in complex with TCA1. In
this way, the CES mechanism ensures the adequate synthesis of PetA according to the availability of its
assembly partners in the Cyt b6f complex.
35
In chloroplasts of seed plants, evidence for a similar feedback regulation mechanism has been obtained in
only a single case. The synthesis of the plastid-encoded large subunit of Rubisco (RbcL) in tobacco is
adjusted to that of its nucleus-encoded assembly partner, the small subunit RBCS, by a mechanism that is
similar to the CES-regulated rbcL expression in Chlamydomonas (Rodermel et al., 1988; Khrebtukova
and Spreitzer, 1996; Rodermel et al., 1996). Furthermore, evidence was provided that unassembled RbcL
represses its own translation, possibly through direct RNA binding (Yosef et al., 2004; Wostrikoff and
Stern, 2007). In a systematic genetic approach, the essential plastid-encoded Cyt b6f subunits PetA, PetB
and PetD were knocked out to disrupt complex assembly and test for potentially conserved CES
regulatory mechanisms in tobacco (Monde et al., 2000). In the mutants, only a mild effect on translation
of the polycistronic petA transcript was detected by polysome analyses, suggesting that the CES
mechanism is not conserved in seed plants (Monde et al., 2000). This may not be entirely surprising,
given that the regulatory trans-factors involved, MCA1 and TCA1, are not found in seed plants.
Moreover, a ribosome profiling analysis of two maize mutants with defective AtpB synthesis did not
show a substantial effect on atpA translation (Zoschke et al., 2013a). This suggests that also the AtpB-
dependent translation of atpA, the only trans-activating CES mechanism observed in Chlamydomonas
(Drapier et al., 2007), is not conserved in seed plants. However, in Arabidopsis, it was observed that
mutation of the chloroplast RNA-binding protein HCF107 that facilitates psbH translation also causes a
reduction in PsbB (CP47) synthesis (Felder et al., 2001). Remarkably, when psbH expression was rescued
by introducing a psbH gene copy into the nuclear genome of an hcf107 mutant, accumulation of the PsbB
protein was rescued as well, despite the absence of functional HCF107 protein (Levey et al., 2014). A
possible explanation for this observation is that PsbH accumulation is needed for PsbB synthesis, pointing
to a potential CES mechanism in PSII biogenesis (Levey et al., 2014). Interestingly, recent transcriptome-
wide analyses of translation in maize and Arabidopsis chloroplasts revealed that most proteins are
synthesized in the amounts that correspond to (i) their steady state stoichiometry as subunits of protein
complexes, and (ii) the abundancy of the respective protein complex (Chotewutmontri and Barkan, 2016;
Lukoszek et al., 2016). This suggests that precise regulation of gene expression is more important than
post-translational proteolytic adjustments of subunit stoichiometry. However, several alternative
regulatory mechanisms can explain this behavior, and it remains to be examined whether or not CES is
involved.
In many organisms, auto-regulated proteins have been observed to comprise an intrinsic RNA-binding
activity. Prominent examples include several ribosomal proteins, the initiation factor IF3 and the β
subunit of the bacterial RNA polymerase (Choquet and Wollman, 2009). The RNA-binding motifs allow
these proteins to bind the 5’UTR of their own mRNA and inhibit translation initiation whenever their
36
assembly partners are not available, thus generating a negative regulatory feedback loop. The chloroplast
genome codes for numerous RNA-binding proteins (e.g., orthologues of bacterial RNA polymerase
subunits and ribosomal proteins). Hence, it is tempting to speculate that some of them may also auto-
regulate their own synthesis. A plastid-encoded protein that has been shown to associate with its own
RNA in vivo is the putative splicing factor MatK (Zoschke et al., 2010). The maturase MatK is encoded in
the intron of the trnK gene. MatK binds to the trnK transcript, most likely assisting in splicing (Zoschke
et al., 2010; and references therein). Notably, a MatK-related bacterial maturase is translationally auto-
regulated (Singh et al., 2002), and an auto-regulatory mechanism was also proposed for the expression of
plastid MatK (Hertel et al., 2013).
In summary, whether autoregulation and/or feedback regulation of translation are common in seed plant
chloroplasts, or whether these regulatory mechanisms are specific to Chlamydomonas, remains to be
investigated. Pronounced differences between microalgae and seed plants may not be surprising, if one
considers the dissimilar evolutionary forces that act on regulatory circuits in a unicellular organism with
one chloroplast per cell compared to multicellular organisms with numerous chloroplasts per cell.
Translational regulation in response to other internal and external triggers
Temperature is a major abiotic signal influencing translational regulation. In bacteria, cold-stress
conditions induce a general block of translation that is mediated by the pY protein. However, the
synthesis of a small subset of proteins, such as RNA helicases, IF1-3 and trigger factor, is induced under
these conditions (Barria et al., 2013). In chloroplasts, changes in temperature alter the speed of all
enzymatic reactions (including the Calvin-Benson-Bassham cycle) but, unless extreme, they have
virtually no impact on the light reactions of photosynthesis. Consequently, temperature changes can cause
dramatic imbalances in the redox homoeostasis of photosynthesis (e.g., Crosatti et al., 2013). It has long
been known that the chloroplast translation capacity is crucial to plant adaptation to chilling stress. This
became evident by the discovery of numerous mutants with impaired chloroplast translation that exhibit
cold-sensitive phenotypes (e.g., Barkan, 1993; Rogalski et al., 2008; Liu et al., 2010).
A recent ribosome profiling study comprehensively examined chloroplast translational regulation upon
temperature stress (Lukoszek et al., 2016). Ignatova and co-workers observed in Arabidopsis that elevated
temperature causes dramatic changes in the protein synthesis output of subunits of photosynthetic
complexes (Lukoszek et al., 2016). This regulation was driven by both changes in transcript levels and
translational regulation. Interestingly, in several cases, unbalanced changes in protein synthesis rates were
observed for different subunits within a complex, in that the altered synthesis rates did not reflect the
subunit stoichiometry. This finding may suggest an increased relevance of control at the level of protein
37
turnover of at least some subunits during heat stress (Lukoszek et al., 2016). In sum, although it is well
established that plastid translation is critical for acclimation to changing temperature conditions, the
molecular mechanisms and the factors involved are currently unknown.
A special case of internal specialization of gene expression is the chloroplast dimorphism in C4 plants. In
C4 species such as maize, light reactions and carbon reactions of photosynthesis are partitioned between
mesophyll cells and bundle sheath cells. The division of labor between the two cell types is reflected by
specialized chloroplast types, referred to as chloroplast dimorphism. At the molecular level, bundle sheath
chloroplasts have a high content of Rubisco and NDH complex, whereas mesophyll chloroplasts are
enriched in PSII (e.g., Majeran et al., 2010). This specialization is achieved by differential gene
expression (e.g., Sharpe et al., 2011; and references therein). Recently, transcript accumulation and
translation have been comprehensively analyzed in bundle sheath and mesophyll chloroplasts by
ribosome profiling (Chotewutmontri and Barkan, 2016). This study confirmed that most of the observed
differences in protein accumulation can be explained by differential protein synthesis which in turn is
predominantly achieved by differential expression at the RNA level. However, the synthesis of some
proteins such as the PSII core subunits PsbA/B/C/D is additionally tuned by translational regulation. The
underlying regulatory mechanisms are unknown. The study showed that the synthesis output of Rubisco,
PSII, PSI and NDH complex subunits reflects well their protein accumulation in the different chloroplast
types. This is not the case for the subunits of the ATP synthase and the Cyt b6f complex, suggesting more
extensive post-translational adjustments in these complexes.
Internal metabolic signals can also control translation (e.g. via riboswitches in bacteria; Serganov and
Nudler, 2013). Several recent discoveries in Chlamydomonas have established interesting links between
chloroplast metabolism and gene expression (Bohne and Nickelsen, 2017). DLA2, a subunit of the
chloroplast pyruvate dehydrogenase complex, was shown to have RNA-binding activity which enables its
moonlighting function in translational regulation of the psbA mRNA (Bohne et al., 2013). Considering its
functions in metabolism and translational regulation, DLA2 is anticipated to coordinate fatty acid and
protein syntheses during thylakoid biogenesis (Bohne et al., 2013). The phylogenetic conservation of the
DLA2 amino acid sequence and RNA-binding properties suggests that it performs a similar dual function
in seed plants (Bohne et al., 2013). A moonlighting function was also suggested for RbcL, which was
shown to bind RNA (Yosef et al., 2004; Cohen et al., 2006), and recently has been proposed to be
involved in the redox stress-induced localization of oxidized chloroplast RNAs in Chlamydomonas,
independent of its function in the Rubisco complex (Zhan et al., 2015). Moreover, recent in vitro data
suggest that PsbD protein synthesis in Chlamydomonas is metabolically controlled (Schwarz et al., 2012;
38
see above). Whether similar regulatory connections between metabolism and chloroplast translation exist
in seed plants remains to be investigated.
In bacteria, stress-induced regulation of gene expression is triggered by the so-called stringent response.
Upon nutrient deprivation and other stresses, bacteria produce the effector molecule (p)ppGpp (guanosine
tetraphosphate/pentaphosphate, also known as alarmone) that coordinates numerous cellular responses,
including broad changes in transcription and translation (Gaca et al., 2015). Chloroplasts also harbor the
enzymes needed for ppGpp synthesis (and degradation). Plastids produce alarmone not only in response
to stress, but also to orchestrate chloroplast and nuclear gene expression (reviewed, e.g., in Field, 2017).
ppGpp has been shown to influence transcription, translation, and many other metabolic and
physiological processes in plants (e.g., Field, 2017). In an in vitro chloroplast translation system from pea,
ppGpp inhibited protein synthesis, suggesting conservation of the bacterial mode of ppGpp-mediated
translational repression by interaction with IF2 and/or EF-G (Nomura et al., 2012). Furthermore,
overaccumulation of ppGpp in vivo caused a dramatic reduction in the accumulation of many plastid-
encoded proteins, mRNAs, tRNA and rRNAs, suggesting an overall reduced translation capacity (e.g.,
Sugliani et al., 2016). Whether the translational activity of specific chloroplast mRNAs is regulated by
ppGpp and how this contributes to ppGpp-induced changes in chloroplast gene expression still needs to
be determined.
In summary, considerable progress has been made in describing changes in translation in response to light
and developmental signals, but comparably little is known about the mechanisms and factors that adjust
plastid protein synthesis to other internal and external cues that affect protein homeostasis.
OUTLOOK
Over the last decades, extensive research on the structure and function of the translational apparatus of
bacteria and chloroplasts has revealed many similarities but also substantial differences. Unfortunately,
our knowledge about genuine regulatory processes, the underlying molecular mechanisms and the factors
involved is still limited. For example, the influence of internal and external triggers such as light,
temperature, nutrient availability, osmotic status, redox state, phytohormones, alarmone, and diurnal and
circadian rhythms on chloroplast gene expression has so far not been comprehensively examined at the
translational level. Likewise, investigation of the interconnection of chloroplast translation with protein
synthesis in the cytosol and the mitochondria is likely to provide fresh insights into intracellular signaling
and crosstalk in response to changing environmental conditions and developmental programs. Last but
not least, the detailed molecular function of RNA cis-elements and proteinaceous trans-factors in the
regulation of chloroplast translation is mostly unknown. The exciting advent of chloroplast ribosome
39
profiling and other in vivo ribonomic methods, such as the comprehensive examination of RNA
secondary structures or the mapping of binding sites for RNA-binding proteins, can be anticipated to
address many of these open questions in the biology of plant organelles in the future.
ACKNOWLEDGEMENTS
We apologize to the authors of numerous papers we were unable to cite due to space constraints. We
thank Alice Barkan (University of Oregon) for helpful suggestions on the manuscript. We acknowledge
the generous sharing of unpublished data by Alice Barkan, Christian Schmitz-Linneweber (Humboldt
University of Berlin), Karin Meierhoff (Heinrich Heine University Düsseldorf), and Mark Aurel Schöttler
(Max Planck Institute of Molecular Plant Physiology). Research on plastid translation in the authors’
laboratories is funded by the Max Planck Society, the Deutsche Forschungsgemeinschaft (DFG; grant ZO
302/4-1 to R.Z., SFB-TRR 175 to R.Z. and R.B.), the German Academic Exchange Service (DAAD;
project ID 57387429 to R.Z.), and the European Research Council (ERC) under the European Union’s
Horizon 2020 research and innovation programme (ERC-ADG-2014; grant agreement No 669982 to
R.B.).
AUTHOR CONTRIBUTIONS
RZ and RB wrote the paper.
REFERENCES
Adachi, Y., Kuroda, H., Yukawa, Y., and Sugiura, M. (2012). Translation of partially overlapping psbD-psbC mRNAs in chloroplasts: the role of 5'-processing and translational coupling. Nucleic Acids Res 40, 3152-3158.
Ahlert, D., Ruf, S., and Bock, R. (2003). Plastid protein synthesis is required for plant development in tobacco. Proc Natl Acad Sci U S A 100, 15730-15735.
Ahmed, T., Yin, Z., and Bhushan, S. (2016). Cryo-EM structure of the large subunit of the spinach chloroplast ribosome. Sci Rep 6, 35793.
Akkaya, M.S., and Breitenberger, C.A. (1992). Light regulation of protein synthesis factor EF-G in pea chloroplasts. Plant Mol Biol 20, 791-800.
Albrecht, V., Ingenfeld, A., and Apel, K. (2006). Characterization of the snowy cotyledon 1 mutant of Arabidopsis thaliana: the impact of chloroplast elongation factor G on chloroplast development and plant vitality. Plant Mol Biol 60, 507-518.
Alkatib, S., Scharff, L.B., Rogalski, M., Fleischmann, T.T., Matthes, A., Seeger, S., Schöttler, M.A., Ruf, S., and Bock, R. (2012). The contributions of wobbling and superwobbling to the reading of the genetic code. PLoS Genet 8, e1003076.
Allen, J.F. (2015). Why chloroplasts and mitochondria retain their own genomes and genetic systems: Colocation for redox regulation of gene expression. Proc Natl Acad Sci U S A 112, 10231-10238.
Allen, J.F., and Pfannschmidt, T. (2000). Balancing the two photosystems: photosynthetic electron transfer governs transcription of reaction centre genes in chloroplasts. Philos Trans R Soc Lond B Biol Sci 355, 1351-1359.
Allen, J.F., de Paula, W.B., Puthiyaveetil, S., and Nield, J. (2011). A structural phylogenetic map for chloroplast photosynthesis. Trends Plant Sci 16, 645-655.
40
Amin, P., Sy, D.A., Pilgrim, M.L., Parry, D.H., Nussaume, L., and Hoffman, N.E. (1999). Arabidopsis mutants lacking the 43- and 54-kilodalton subunits of the chloroplast signal recognition particle have distinct phenotypes. Plant Physiol 121, 61-70.
Artieri, C.G., and Fraser, H.B. (2014). Accounting for biases in riboprofiling data indicates a major role for proline in stalling translation. Genome Res 24, 2011-2021.
Asakura, Y., Kikuchi, S., and Nakai, M. (2008). Non-identical contributions of two membrane-bound cpSRP components, cpFtsY and Alb3, to thylakoid biogenesis. Plant J 56, 1007-1017.
Barbrook, A.C., Howe, C.J., and Purton, S. (2006). Why are plastid genomes retained in non-photosynthetic organisms? Trends Plant Sci 11, 101-108.
Barkan, A. (1988). Proteins encoded by a complex chloroplast transcription unit are each translated from both monocistronic and polycistronic mRNAs. EMBO J 7, 2637-2644.
Barkan, A. (1989). Tissue-dependent plastid RNA splicing in maize: transcripts from four plastid genes are predominantly unspliced in leaf meristems and roots. The Plant Cell 1, 437-445.
Barkan, A. (1993). Nuclear mutants of maize with defects in chloroplast polysome assembly have altered chloroplast RNA metabolism. The Plant Cell 5, 389-402.
Barkan, A. (1998). Approaches to investigating nuclear genes that function in chloroplast biogenesis in land plants. Methods Enzymol 297, 38-57.
Barkan, A. (2011). Expression of plastid genes: organelle-specific elaborations on a prokaryotic scaffold. Plant Physiol 155, 1520-1532.
Barkan, A., and Small, I. (2014). Pentatricopeptide repeat proteins in plants. Annu Rev Plant Biol 65, 415-442.
Barkan, A., Walker, M., Nolasco, M., and Johnson, D. (1994). A nuclear mutation in maize blocks the processing and translation of several chloroplast mRNAs and provides evidence for the differential translation of alternative mRNA forms. EMBO J 13, 3170-3181.
Barria, C., Malecki, M., and Arraiano, C.M. (2013). Bacterial adaptation to cold. Microbiology 159, 2437-2443.
Baumgartner, B.J., Rapp, J.C., and Mullet, J.E. (1989). Plastid Transcription Activity and DNA Copy Number Increase Early in Barley Chloroplast Development. Plant Physiol 89, 1011-1018.
Beligni, M.V., Yamaguchi, K., and Mayfield, S.P. (2004). The translational apparatus of Chlamydomonas reinhardtii chloroplast. Photosynth Res 82, 315-325.
Berry, J.O., Carr, J.P., and Klessig, D.F. (1988). mRNAs encoding ribulose-1,5-bisphosphate carboxylase remain bound to polysomes but are not translated in amaranth seedlings transferred to darkness. Proc Natl Acad Sci U S A 85, 4190-4419.
Bhadula, S.K., Elthon, T.E., Habben, J.E., Helentjaris, T.G., Jiao, S., and Ristic, Z. (2001). Heat-stress induced synthesis of chloroplast protein synthesis elongation factor (EF-Tu) in a heat-tolerant maize line. Planta 212, 359-366.
Bhushan, S., Gartmann, M., Halic, M., Armache, J.P., Jarasch, A., Mielke, T., Berninghausen, O., Wilson, D.N., and Beckmann, R. (2010). α-Helical nascent polypeptide chains visualized within distinct regions of the ribosomal exit tunnel. Nat Struct Mol Biol 17, 313-317.
Bieri, P., Leibundgut, M., Saurer, M., Boehringer, D., and Ban, N. (2017). The complete structure of the chloroplast 70S ribosome in complex with translation factor pY. EMBO J 36, 475-486.
Bock, R. (2007). Cell and Molecular Biology of Plastids. (Berlin: Springer Verlag). Bock, R., and Timmis, J.N. (2008). Reconstructing evolution: gene transfer from plastids to the nucleus.
Bioessays 30, 556-566. Bock, R., Kössel, H., and Maliga, P. (1994). Introduction of a heterologous editing site into the tobacco
plastid genome: the lack of RNA editing leads to a mutant phenotype. EMBO J 13, 4623-4628. Boeck, R., and Kolakofsky, D. (1994). Positions +5 and +6 can be major determinants of the efficiency of
non-AUG initiation codons for protein synthesis. EMBO J 13, 3608-3617. Boehm, M., Romero, E., Reisinger, V., Yu, J., Komenda, J., Eichacker, L.A., Dekker, J.P., and Nixon,
P.J. (2011). Investigating the early stages of photosystem II assembly in Synechocystis sp. PCC 6803: isolation of CP47 and CP43 complexes. J Biol Chem 286, 14812-14819.
Bohne, A.V., and Nickelsen, J. (2017). Metabolic Control of Chloroplast Gene Expression: An Emerging Theme. Mol Plant 10, 1-3.
41
Bohne, A.V., Schwarz, C., Schottkowski, M., Lidschreiber, M., Piotrowski, M., Zerges, W., and Nickelsen, J. (2013). Reciprocal regulation of protein synthesis and carbon metabolism for thylakoid membrane biogenesis. PLoS Biol 11, e1001482.
Börner, T., Aleynikova, A.Y., Zubo, Y.O., and Kusnetsov, V.V. (2015). Chloroplast RNA polymerases: Role in chloroplast biogenesis. Biochim Biophys Acta 1847, 761-769.
Boulouis, A., Raynaud, C., Bujaldon, S., Aznar, A., Wollman, F.A., and Choquet, Y. (2011). The nucleus-encoded trans-acting factor MCA1 plays a critical role in the regulation of cytochrome f synthesis in Chlamydomonas chloroplasts. The Plant Cell 23, 333-349.
Breiman, A., Fieulaine, S., Meinnel, T., and Giglione, C. (2016). The intriguing realm of protein biogenesis: Facing the green co-translational protein maturation networks. Biochim Biophys Acta 1864, 531-550.
Breitenberger, C.A., Graves, M.C., and Spremulli, L.L. (1979). Evidence for the nuclear location of the gene for chloroplast elongation factor G. Arch Biochem Biophys 194, 265-270.
Bubunenko, M.G., Schmidt, J., and Subramanian, A.R. (1994). Protein substitution in chloroplast ribosome evolution. A eukaryotic cytosolic protein has replaced its organelle homologue (L23) in spinach. J Mol Biol 240, 28-41.
Buckingham, R.H., Grentzmann, G., and Kisselev, L. (1997). Polypeptide chain release factors. Mol Microbiol 24, 449-456.
Cahoon, A.B., Harris, F.M., and Stern, D.B. (2004). Analysis of developing maize plastids reveals two mRNA stability classes correlating with RNA polymerase type. EMBO Rep 5, 801-806.
Cai, W., Okuda, K., Peng, L., and Shikanai, T. (2011). PROTON GRADIENT REGULATION 3 recognizes multiple targets with limited similarity and mediates translation and RNA stabilization in plastids. Plant J 67, 318-327.
Campos, F., Garcia-Gomez, B.I., Solorzano, R.M., Salazar, E., Estevez, J., Leon, P., Alvarez-Buylla, E.R., and Covarrubias, A.A. (2001). A cDNA for nuclear-encoded chloroplast translational initiationfactor 2 from a higher plant is able to complement an infB Escherichia coli null mutant. J Biol Chem 276,28388-28394.
Celedon, J.M., and Cline, K. (2013). Intra-plastid protein trafficking: how plant cells adapted prokaryotic mechanisms to the eukaryotic condition. Biochim Biophys Acta 1833, 341-351.
Choquet, Y., and Wollman, F.A. (2009). The CES process. In Chlamydomonas Source Book, E. Harris, D.B. Stern, and G. Whitman, eds (New York: Academic Press/Elsevier), pp. 1027–1064.
Chotewutmontri, P., and Barkan, A. (2016). Dynamics of Chloroplast Translation during Chloroplast Differentiation in Maize. PLoS Genet 12, e1006106.
Cohen, I., Sapir, Y., and Shapira, M. (2006). A conserved mechanism controls translation of Rubisco large subunit in different photosynthetic organisms. Plant Physiol 141, 1089-1097.
Crosatti, C., Rizza, F., Badeck, F.W., Mazzucotelli, E., and Cattivelli, L. (2013). Harden the chloroplast to protect the plant. Physiol Plant 147, 55-63.
Dana, A., and Tuller, T. (2014). The effect of tRNA levels on decoding times of mRNA codons. Nucleic Acids Res 42, 9171-9181.
Doerfel, L.K., Wohlgemuth, I., Kothe, C., Peske, F., Urlaub, H., and Rodnina, M.V. (2013). EF-P is essential for rapid synthesis of proteins containing consecutive proline residues. Science 339, 85-88.
Drapier, D., Rimbault, B., Vallon, O., Wollman, F.A., and Choquet, Y. (2007). Intertwined translational regulations set uneven stoichiometry of chloroplast ATP synthase subunits. EMBO J 26, 3581-3591.
Drechsel, O., and Bock, R. (2011). Selection of Shine-Dalgarno sequences in plastids. Nucleic Acids Res 39, 1427-1438.
Eberhard, S., Drapier, D., and Wollman, F.A. (2002). Searching limiting steps in the expression of chloroplast-encoded proteins: relations between gene copy number, transcription, transcript abundance and translation rate in the chloroplast of Chlamydomonas reinhardtii. Plant J 31, 149-160.
Eibl, C., Zou, Z., Beck, a., Kim, M., Mullet, J., and Koop, H.U. (1999). In vivo analysis of plastid psbA, rbcL and rpl32 UTR elements by chloroplast transformation: tobacco plastid gene expression is controlled by modulation of transcript levels and translation efficiency. Plant J 19, 333-345.
Eichacker, L., Paulsen, H., and Rüdiger, W. (1992). Synthesis of chlorophyll a regulates translation of chlorophyll a apoproteins P700, CP47, CP43 and D2 in barley etioplasts. Eur J Biochem 205, 17-24.
42
Eichacker, L.A., Helfrich, M., Rüdiger, W., and Müller, B. (1996). Stabilization of chlorophyll a-binding apoproteins P700, CP47, CP43, D2, and D1 by chlorophyll a or Zn-pheophytin a. J Biol Chem 271, 32174-32179.
Eichacker, L.A., Soll, J., Lauterbach, P., Rüdiger, W., Klein, R.R., and Mullet, J.E. (1990). In vitro synthesis of chlorophyll a in the dark triggers accumulation of chlorophyll a apoproteins in barley etioplasts. J Biol Chem 265, 13566-13571.
Esposito, D., Fey, J.P., Eberhard, S., Hicks, A.J., and Stern, D.B. (2003). In vivo evidence for the prokaryotic model of extended codon-anticodon interaction in translation initiation. EMBO J 22, 651-656.
Fargo, D.C., Zhang, M., Gillham, N.W., and Boynton, J.E. (1998). Shine-Dalgarno-like sequences are not required for translation of chloroplast mRNAs in Chlamydomonas reinhardtii chloroplasts or in Escherichia coli. Mol Gen Genet 257, 271-282.
Felder, S., Meierhoff, K., Sane, A.P., Meurer, J., Driemel, C., Plucken, H., Klaff, P., Stein, B., Bechtold, N., and Westhoff, P. (2001). The nucleus-encoded HCF107 gene of Arabidopsis provides a link between intercistronic RNA processing and the accumulation of translation-competent psbH transcripts in chloroplasts. Plant Cell 13, 2127-2141.
Field, B. (2017). Green magic: regulation of the chloroplast stress response by (p)ppGpp in plants and algae. J Exp Bot.
Fluman, N., Navon, S., Bibi, E., and Pilpel, Y. (2014). mRNA-programmed translation pauses in the targeting of E. coli membrane proteins. Elife 3.
Fox, L., Erion, J., Tarnowski, J., Spremulli, L., Brot, N., and Weissbach, H. (1980). Euglena gracilis chloroplast EF-Ts. Evidence that it is a nuclear-coded gene product. J Biol Chem 255, 6018-6019.
Franzetti, B., Carol, P., and Mache, R. (1992). Characterization and RNA-binding properties of a chloroplast S1-like ribosomal protein. J Biol Chem 267, 19075-19081.
Friemann, A., and Hachtel, W. (1988). Chloroplast messenger RNAs of free and thylakoid-bound polysomes from Vicia faba L. Planta 175, 50-59.
Gaca, A.O., Colomer-Winter, C., and Lemos, J.A. (2015). Many means to a common end: the intricacies of (p)ppGpp metabolism and its control of bacterial homeostasis. J Bacteriol 197, 1146-1156.
Gawronski, P., Jensen, P.E., Karpinski, S., Leister, D., and Scharff, L.B. (2018). Plastid ribosome pausing is induced by multiple features and is linked to protein complex assembly. Plant Physiol.
Germain, A., Hotto, A.M., Barkan, A., and Stern, D.B. (2013). RNA processing and decay in plastids. Wiley Interdiscip Rev RNA 4, 295-316.
Giglione, C., Fieulaine, S., and Meinnel, T. (2015). N-terminal protein modifications: Bringing back into play the ribosome. Biochimie 114, 134-146.
Gloge, F., Becker, A.H., Kramer, G., and Bukau, B. (2014). Co-translational mechanisms of protein maturation. Curr Opin Struct Biol 24, 24-33.
Graf, M., Arenz, S., Huter, P., Dönhöfer, A., Nováček, J., and Wilson, D.N. (2016). Cryo-EM structure of the spinach chloroplast ribosome reveals the location of plastid-specific ribosomal proteins and extensions. Nucleic Acids Res.
Green, B.R. (2011). Chloroplast genomes of photosynthetic eukaryotes. Plant J 66, 34-44. Grennan, A.K., and Ort, D.R. (2007). Cool temperatures interfere with D1 synthesis in tomato by causing
ribosomal pausing. Photosynth Res 94, 375-385. Hammani, K., Cook, W.B., and Barkan, A. (2012). RNA binding and RNA remodeling activities of the
half-a-tetratricopeptide (HAT) protein HCF107 underlie its effects on gene expression. Proc Natl Acad Sci U S A 109, 5651-5656.
Hammani, K., Bonnard, G., Bouchoucha, A., Gobert, A., Pinker, F., Salinas, T., and Giege, P. (2014). Helical repeats modular proteins are major players for organelle gene expression. Biochimie 100, 141-150.
Hashimoto, M., Endo, T., Peltier, G., Tasaka, M., and Shikanai, T. (2003). A nucleus-encoded factor, CRR2, is essential for the expression of chloroplast ndhB in Arabidopsis. Plant J 36, 541-549.
Hershey, J.W., Sonenberg, N., and Mathews, M.B. (2012). Principles of translational control: an overview. Cold Spring Harb Perspect Biol 4.
Hertel, S., Zoschke, R., Neumann, L., Qu, Y., Axmann, I.M., and Schmitz-Linneweber, C. (2013). Multiple checkpoints for the expression of the chloroplast-encoded splicing factor MatK. Plant Physiol 163, 1686-1698.
43
Hirose, T., and Sugiura, M. (1996). Cis-acting elements and trans-acting factors for accurate translation of chloroplast psbA mRNAs: development of an in vitro translation system from tobacco chloroplasts. EMBO J 15, 1687-1695.
Hirose, T., and Sugiura, M. (1997). Both RNA editing and RNA cleavage are required for translation of tobacco chloroplast ndhD mRNA: a possible regulatory mechanism for the expression of a chloroplast operon consisting of functionally unrelated genes. EMBO J 16, 6804-6811.
Hirose, T., and Sugiura, M. (2004). Functional Shine-Dalgarno-like sequences for translational initiation of chloroplast mRNAs. Plant Cell Physiol 45, 114-117.
Hirose, T., Kusumegi, T., and Sugiura, M. (1998). Translation of tobacco chloroplast rps14 mRNA depends on a Shine-Dalgarno-like sequence in the 5'-untranslated region but not on internal RNA editing in the coding region. FEBS Lett 430, 257-260.
Hirose, T., Ideue, T., Wakasugi, T., and Sugiura, M. (1999). The chloroplast infA gene with a functional UUG initiation codon. FEBS Lett 445, 169-172.
Hoch, B., Maier, R.M., Appel, K., Igloi, G.L., and Kössel, H. (1991). Editing of a chloroplast mRNA by creation of an initiation codon. Nature 353, 178-180.
Holtkamp, W., Kokic, G., Jäger, M., Mittelstaet, J., Komar, A.A., and Rodnina, M.V. (2015). Cotranslational protein folding on the ribosome monitored in real time. Science 350, 1104-1107.
Huber, D., Jamshad, M., Hanmer, R., Schibich, D., Döring, K., Marcomini, I., Kramer, G., and Bukau, B. (2017). SecA Cotranslationally Interacts with Nascent Substrate Proteins In Vivo. J Bacteriol 199.
Ibhaya, D., and Jagendorf, A.T. (1984). Synthesis of subunit III of CF0 by thylakoid-bound polysomes from pea chloroplasts. Plant Mol Biol 3, 277-280.
Ichinose, M., and Sugita, M. (2016). RNA Editing and Its Molecular Mechanism in Plant Organelles. Genes (Basel) 8.
Ingolia, N.T. (2016). Ribosome Footprint Profiling of Translation throughout the Genome. Cell 165, 22-33. Ingolia, N.T., Ghaemmaghami, S., Newman, J.R., and Weissman, J.S. (2009). Genome-wide analysis in
vivo of translation with nucleotide resolution using ribosome profiling. Science 324, 218-223. Ito, K., Chiba, S., and Pogliano, K. (2010). Divergent stalling sequences sense and control cellular
physiology. Biochem Biophys Res Commun 393, 1-5. Jackson, R.J., Kaminski, A., and Pöyry, T.A.A. (2007). Coupled termination-reinitiation events in mRNA
translation. In Translational Control in Biology and Medicine, M.B. Mathews, N. Sonenberg, and J.W.B. Hershey, eds (New York: Cold Spring Harbor Laboratory Press), pp. 197-223.
Jan, C.H., Williams, C.C., and Weissman, J.S. (2014). Principles of ER cotranslational translocation revealed by proximity-specific ribosome profiling. Science 346, 1257521.
Jarvis, P., and Lopez-Juez, E. (2013). Biogenesis and homeostasis of chloroplasts and other plastids. Nat Rev Mol Cell Biol 14, 787-802.
Kahlau, S., and Bock, R. (2008). Plastid transcriptomics and translatomics of tomato fruit development and chloroplast-to-chromoplast differentiation: chromoplast gene expression largely serves the production of a single protein. Plant Cell 20, 856-874.
Kallia-Raftopoulos, S., and Kalpaxis, D.L. (1999). Slow sequential conformational changes in Escherichia coli ribosomes induced by lincomycin: kinetic evidence. Mol Pharmacol 56, 1042-1046.
Keene, J.D. (2007). RNA regulons: coordination of post-transcriptional events. Nat Rev Genet 8, 533-543. Khrebtukova, I., and Spreitzer, R.J. (1996). Elimination of the Chlamydomonas gene family that encodes
the small subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase. Proc Natl Acad Sci U S A 93, 13689-13693.
Kiel, M.C., Kaji, H., and Kaji, A. (2007). Ribosome recycling: An essential process of protein synthesis. Biochem Mol Biol Educ 35, 40-44.
Kim, J., and Mullet, J.E. (1994). Ribosome-binding sites on chloroplast rbcL and psbA mRNAs and light-induced initiation of D1 translation. Plant Mol Biol 25, 437-448.
Kim, J., and Mullet, J.E. (2003). A mechanism for light-induced translation of the rbcL mRNA encoding the large subunit of ribulose-1,5-bisphosphate carboxylase in barley chloroplasts. Plant Cell Physiol 44, 491-499.
Kim, J., Klein, P.G., and Mullet, J.E. (1991). Ribosomes pause at specific sites during synthesis of membrane-bound chloroplast reaction center protein D1. J Biol Chem 266, 14931-14938.
44
Kim, J., Klein, P.G., and Mullet, J.E. (1994a). Synthesis and turnover of photosystem II reaction center protein D1. Ribosome pausing increases during chloroplast development. J Biol Chem 269, 17918-17923.
Kim, J., Eichacker, L.A., Rudiger, W., and Mullet, J.E. (1994b). Chlorophyll regulates accumulation of the plastid-encoded chlorophyll proteins P700 and D1 by increasing apoprotein stability. Plant Physiol 104, 907-916.
Kim, J.K., and Hollingsworth, M.J. (1992). Localization of in vivo ribosome pause sites. Anal Biochem 206, 183-188.
Klaff, P., and Gruissem, W. (1991). Changes in Chloroplast mRNA Stability during Leaf Development. Plant Cell 3, 517-529.
Kleffmann, T., von Zychlinski, A., Russenberger, D., Hirsch-Hoffmann, M., Gehrig, P., Gruissem, W., and Baginsky, S. (2007). Proteome dynamics during plastid differentiation in rice. Plant Physiol 143, 912-923.
Klein, R.R., and Mullet, J.E. (1986). Regulation of chloroplast-encoded chlorophyll-binding protein translation during higher plant chloroplast biogenesis. J Biol Chem 261, 11138-11145.
Klein, R.R., and Mullet, J.E. (1987). Control of gene expression during higher plant chloroplast biogenesis. Protein synthesis and transcript levels of psbA, psaA-psaB, and rbcL in dark-grown and illuminated barley seedlings. J Biol Chem 262, 4341-4348.
Klein, R.R., Gamble, P.E., and Mullet, J.E. (1988a). Light-Dependent Accumulation of Radiolabeled Plastid-Encoded Chlorophyll a-Apoproteins Requires Chlorophyll a: I. Analysis of Chlorophyll-Deficient Mutants and Phytochrome Involvement. Plant Physiol 88, 1246-1256.
Klein, R.R., Mason, H.S., and Mullet, J.E. (1988b). Light-regulated translation of chloroplast proteins. I. Transcripts of psaA-psaB, psbA, and rbcL are associated with polysomes in dark-grown and illuminated barley seedlings. J Cell Biol 106, 289-301.
Kleine, T., and Leister, D. (2016). Retrograde signaling: Organelles go networking. Biochim Biophys Acta 1857, 1313-1325.
Klug, G. (1993). The role of mRNA degradation in the regulated expression of bacterial photosynthesis genes. Mol Microbiol 9, 1-7.
Kössel, H., Natt, E., Strittmatter, G., Fritzche, E., Gozdzicka-Jozefiak, A., and Przybyl, D. (1985). Structure and expression of rRNA operons from plastids of higher plants. In Molecular form and function of the plant genome, L. van Vloten-Doting, G. Groot, and T. Hall, eds (Plenum Publishing Corporation), pp. 183-198.
Kroliczewski, J., Piskozub, M., Bartoszewski, R., and Kroliczewska, B. (2016). ALB3 Insertase Mediates Cytochrome b6 Co-translational Import into the Thylakoid Membrane. Sci Rep 6, 34557.
Kroll, D., Meierhoff, K., Bechtold, N., Kinoshita, M., Westphal, S., Vothknecht, U.C., Soll, J., and Westhoff, P. (2001). VIPP1, a nuclear gene of Arabidopsis thaliana essential for thylakoid membrane formation. Proc Natl Acad Sci U S A 98, 4238-4242.
Kupsch, C., Ruwe, H., Gusewski, S., Tillich, M., Small, I., and Schmitz-Linneweber, C. (2012). Arabidopsis chloroplast RNA binding proteins CP31A and CP29A associate with large transcript pools and confer cold stress tolerance by influencing multiple chloroplast RNA processing steps. Plant Cell 24, 4266-4280.
Kuras, R., and Wollman, F.A. (1994). The assembly of cytochrome b6/f complexes: an approach using genetic transformation of the green alga Chlamydomonas reinhardtii. EMBO J 13, 1019-1027.
Kuroda, H., Suzuki, H., Kusumegi, T., Hirose, T., Yukawa, Y., and Sugiura, M. (2007). Translation of psbC mRNAs starts from the downstream GUG, not the upstream AUG, and requires the extended Shine-Dalgarno sequence in tobacco chloroplasts. Plant Cell Physiol 48, 1374-1378.
Laalami, S., Zig, L., and Putzer, H. (2014). Initiation of mRNA decay in bacteria. Cell Mol Life Sci 71, 1799-1828.
Leech, R.M., Rumsby, M.G., and Thomson, W.W. (1973). Plastid differentiation, acyl lipid, and fatty acid changes in developing green maize leaves. Plant Physiol 52, 240-245.
Lefebvre-Legendre, L., Merendino, L., Rivier, C., and Goldschmidt-Clermont, M. (2014). On the complexity of chloroplast RNA metabolism: psaA trans-splicing can be bypassed in Chlamydomonas. Mol Biol Evol 31, 2697-2707.
45
Legen, J., Ruf, S., Kroop, X., Wang, G., Barkan, A., Bock, R., and Schmitz-Linneweber, C. (2018). Stabilization and translation of synthetic operon-derived mRNAs in chloroplasts by sequences representing PPR protein binding sites. Plant J.
Lehniger, M.K., Finster, S., Melonek, J., Oetke, S., Krupinska, K., and Schmitz-Linneweber, C. (2017). Global RNA association with the transcriptionally active chromosome of chloroplasts. Plant Mol Biol 95, 303-311.
Levey, T., Westhoff, P., and Meierhoff, K. (2014). Expression of a nuclear-encoded psbH gene complements the plastidic RNA processing defect in the PSII mutant hcf107 in Arabidopsis thaliana. Plant J 80, 292-304.
Li, G.W., Oh, E., and Weissman, J.S. (2012). The anti-Shine-Dalgarno sequence drives translational pausing and codon choice in bacteria. Nature 484, 538-541.
Li, L., Nelson, C.J., Trösch, J., Castleden, I., Huang, S., and Millar, A.H. (2017a). Protein Degradation Rate in Arabidopsis thaliana Leaf Growth and Development. Plant Cell 29, 207-228.
Li, P., Ponnala, L., Gandotra, N., Wang, L., Si, Y., Tausta, S.L., Kebrom, T.H., Provart, N., Patel, R., Myers, C.R., Reidel, E.J., Turgeon, R., Liu, P., Sun, Q., Nelson, T., and Brutnell, T.P. (2010). The developmental dynamics of the maize leaf transcriptome. Nat Genet 42, 1060-1067.
Li, Y., Martin, J.R., Aldama, G.A., Fernandez, D.E., and Cline, K. (2017b). Identification of Putative Substrates of SEC2, a Chloroplast Inner Envelope Translocase. Plant Physiol 173, 2121-2137.
Link, S., Engelmann, K., Meierhoff, K., and Westhoff, P. (2012). The atypical short-chain dehydrogenases HCF173 and HCF244 are jointly involved in translational initiation of the psbA mRNA of Arabidopsis. Plant Physiol 160, 2202-2218.
Liu, X., Rodermel, S.R., and Yu, F. (2010). A var2 leaf variegation suppressor locus, SUPPRESSOR OF VARIEGATION3, encodes a putative chloroplast translation elongation factor that is important for chloroplast development in the cold. BMC Plant Biol 10, 287.
Loiselay, C., Gumpel, N.J., Girard-Bascou, J., Watson, A.T., Purton, S., Wollman, F.A., and Choquet, Y. (2008). Molecular identification and function of cis- and trans-acting determinants for petA transcriptstability in Chlamydomonas reinhardtii chloroplasts. Mol Cell Biol 28, 5529-5542.
Lukoszek, R., Feist, P., and Ignatova, Z. (2016). Insights into the adaptive response of Arabidopsis thaliana to prolonged thermal stress by ribosomal profiling and RNA-Seq. BMC Plant Biol 16, 221.
Lyska, D., Meierhoff, K., and Westhoff, P. (2013). How to build functional thylakoid membranes: from plastid transcription to protein complex assembly. Planta 237, 413-428.
Maier, U.G., Zauner, S., Woehle, C., Bolte, K., Hempel, F., Allen, J.F., and Martin, W.F. (2013). Massively convergent evolution for ribosomal protein gene content in plastid and mitochondrial genomes. Genome Biol Evol 5, 2318-2329.
Maier, U.G., Bozarth, A., Funk, H.T., Zauner, S., Rensing, S.A., Schmitz-Linneweber, C., Börner, T., and Tillich, M. (2008). Complex chloroplast RNA metabolism: just debugging the genetic programme? BMC Biol 6, 36.
Majeran, W., Friso, G., Asakura, Y., Qu, X., Huang, M., Ponnala, L., Watkins, K.P., Barkan, A., and van Wijk, K.J. (2011). Nucleoid-enriched proteomes in developing plastids and chloroplasts from maize leaves: a new conceptual framework for nucleoid functions. Plant Physiol 158, 156-189.
Majeran, W., Friso, G., Ponnala, L., Connolly, B., Huang, M., Reidel, E., Zhang, C., Asakura, Y., Bhuiyan, N.H., Sun, Q., Turgeon, R., and van Wijk, K.J. (2010). Structural and metabolic transitions of C4 leaf development and differentiation defined by microscopy and quantitative proteomics in maize. Plant Cell 22, 3509-3542.
Makarova, O.V., Makarov, E.M., Sousa, R., and Dreyfus, M. (1995). Transcribing of Escherichia coli genes with mutant T7 RNA polymerases: stability of lacZ mRNA inversely correlates with polymerase speed. Proc Natl Acad Sci U S A 92, 12250-12254.
Manuell, A.L., Quispe, J., and Mayfield, S.P. (2007). Structure of the chloroplast ribosome: novel domains for translation regulation. PLoS Biol 5, e209.
Margulies, M.M., Tiffany, H.L., and Hattori, T. (1987). Photosystem I reaction center polypeptides of spinach are synthesized on thylakoid-bound ribosomes. Arch Biochem Biophys 254, 454-461.
Martin Avila, E., Gisby, M.F., and Day, A. (2016). Seamless editing of the chloroplast genome in plants. BMC Plant Biol 16, 168.
46
Martin, W., Rujan, T., Richly, E., Hansen, A., Cornelsen, S., Lins, T., Leister, D., Stoebe, B., Hasegawa, M., and Penny, D. (2002). Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid phylogeny and thousands of cyanobacterial genes in the nucleus. Proc Natl Acad Sci U S A 99, 12246-12251.
Mauger, D.M., Siegfried, N.A., and Weeks, K.M. (2013). The genetic code as expressed through relationships between mRNA structure and protein function. FEBS Lett 587, 1180-1188.
McCormac, D.J., and Barkan, A. (1999). A nuclear gene in maize required for the translation of the chloroplast atpB/E mRNA. Plant Cell 11, 1709-1716.
McGary, K., and Nudler, E. (2013). RNA polymerase and the ribosome: the close relationship. Curr Opin Microbiol 16, 112-117.
Meierhoff, K., Felder, S., Nakamura, T., Bechtold, N., and Schuster, G. (2003). HCF152, an Arabidopsis RNA binding pentatricopeptide repeat protein involved in the processing of chloroplast psbB-psbT-psbH-petB-petD RNAs. Plant Cell 15, 1480-1495.
Meurer, J., Lezhneva, L., Amann, K., Godel, M., Bezhani, S., Sherameti, I., and Oelmuller, R. (2002). A peptide chain release factor 2 affects the stability of UGA-containing transcripts in Arabidopsis chloroplasts. Plant Cell 14, 3255-3269.
Millen, R.S., Olmstead, R.G., Adams, K.L., Palmer, J.D., Lao, N.T., Heggie, L., Kavanagh, T.A., Hibberd, J.M., Gray, J.C., Morden, C.W., Calie, P.J., Jermiin, L.S., and Wolfe, K.H. (2001). Many parallel losses of infA from chloroplast DNA during angiosperm evolution with multiple independent transfers to the nucleus. Plant Cell 13, 645-658.
Miura, E., Kato, Y., Matsushima, R., Albrecht, V., Laalami, S., and Sakamoto, W. (2007). The balance between protein synthesis and degradation in chloroplasts determines leaf variegation in Arabidopsis yellow variegated mutants. Plant Cell 19, 1313-1328.
Monde, R.A., Zito, F., Olive, J., Wollman, F.A., and Stern, D.B. (2000). Post-transcriptional defects in tobacco chloroplast mutants lacking the cytochrome b6/f complex. Plant J 21, 61-72.
Moreno, J.C., Tiller, N., Diez, M., Karcher, D., Tillich, M., Schöttler, M.A., and Bock, R. (2017). Generation and characterization of a collection of knock-down lines for the chloroplast Clp protease complex in tobacco. J Exp Bot.
Motohashi, R., Yamazaki, T., Myouga, F., Ito, T., Ito, K., Satou, M., Kobayashi, M., Nagata, N., Yoshida, S., Nagashima, A., Tanaka, K., Takahashi, S., and Shinozaki, K. (2007). Chloroplast ribosome release factor 1 (AtcpRF1) is essential for chloroplast development. Plant Mol Biol 64, 481-497.
Mühlbauer, S.K., and Eichacker, L.A. (1998). Light-dependent formation of the photosynthetic proton gradient regulates translation elongation in chloroplasts. J Biol Chem 273, 20935-209340.
Müller, B., and Eichacker, L.A. (1999). Assembly of the D1 precursor in monomeric photosystem II reaction center precomplexes precedes chlorophyll a-triggered accumulation of reaction center II in barley etioplasts. Plant Cell 11, 2365-2377.
Mullet, J.E., and Klein, R.R. (1987). Transcription and RNA stability are important determinants of higher plant chloroplast RNA levels. EMBO J 6, 1571-1579.
Mullet, J.E., Klein, P.G., and Klein, R.R. (1990). Chlorophyll regulates accumulation of the plastid-encoded chlorophyll apoproteins CP43 and D1 by increasing apoprotein stability. Proc Natl Acad Sci U S A 87, 4038-4042.
Nakagawa, S., Niimura, Y., and Gojobori, T. (2017). Comparative genomic analysis of translation initiation mechanisms for genes lacking the Shine-Dalgarno sequence in prokaryotes. Nucleic Acids Res 45, 3922-3931.
Nakahigashi, K., Takai, Y., Shiwa, Y., Wada, M., Honma, M., Yoshikawa, H., Tomita, M., Kanai, A., and Mori, H. (2014). Effect of codon adaptation on codon-level and gene-level translation efficiency in vivo. BMC Genomics 15, 1115.
Nakamura, T., Ohta, M., Sugiura, M., and Sugita, M. (2001). Chloroplast ribonucleoproteins function as a stabilizing factor of ribosome-free mRNAs in the stroma. J Biol Chem 276, 147-152.
Natan, E., Wells, J.N., Teichmann, S.A., and Marsh, J.A. (2017). Regulation, evolution and consequences of cotranslational protein complex assembly. Curr Opin Struct Biol 42, 90-97.
47
Nesbit, A.D., Whippo, C., Hangarter, R.P., and Kehoe, D.M. (2015). Translation initiation factor 3 families: what are their roles in regulating cyanobacterial and chloroplast gene expression? Photosynth Res 126, 147-159.
Nickelsen, J., and Rengstl, B. (2013). Photosystem II assembly: from cyanobacteria to plants. Annu Rev Plant Biol 64, 609-635.
Nickelsen, J., Bohne, A.-V., and Westhoff, P. (2014). Chloroplast Gene Expression—Translation. In Plastid Biology, S.M. Theg and F.-A. Wollman, eds (Springer New York), pp. 49-78.
Nierhaus, K.H., and Wittmann, H.G. (1980). Ribosomal function and its inhibition by antibiotics in prokaryotes. Naturwissenschaften 67, 234-250.
Nilsson, R., and van Wijk, K.J. (2002). Transient interaction of cpSRP54 with elongating nascent chains of the chloroplast-encoded D1 protein; 'cpSRP54 caught in the act'. FEBS Lett 524, 127-133.
Nomura, Y., Takabayashi, T., Kuroda, H., Yukawa, Y., Sattasuk, K., Akita, M., Nozawa, A., and Tozawa, Y. (2012). ppGpp inhibits peptide elongation cycle of chloroplast translation system in vitro. Plant Mol Biol 78, 185-196.
Pedersen, S., and Reeh, S. (1978). Functional mRNA half lives in E. coli. Mol Gen Genet 166, 329-336. Peled-Zehavi, H., and Danon, A. (2007). Translation and translational regulation in chloroplasts. In Cell and
Molecular Biology of Plastids, R. Bock, ed (Berlin: Springer Verlag), pp. 249-281. Pesaresi, P., Varotto, C., Meurer, J., Jahns, P., Salamini, F., and Leister, D. (2001). Knock-out of the
plastid ribosomal protein L11 in Arabidopsis: effects on mRNA translation and photosynthesis. Plant J 27, 179-189.
Pfalz, J., and Pfannschmidt, T. (2013). Essential nucleoid proteins in early chloroplast development. Trends Plant Sci 18, 186-194.
Pfalz, J., Bayraktar, O.A., Prikryl, J., and Barkan, A. (2009). Site-specific binding of a PPR protein defines and stabilizes 5' and 3' mRNA termini in chloroplasts. EMBO J 28, 2042-2052.
Pfalz, J., Liere, K., Kandlbinder, A., Dietz, K.J., and Oelmuller, R. (2006). pTAC2, -6, and -12 are components of the transcriptionally active plastid chromosome that are required for plastid gene expression. Plant Cell 18, 176-197.
Pfalz, J., Liebers, M., Hirth, M., Grubler, B., Holtzegel, U., Schröter, Y., Dietzel, L., and Pfannschmidt, T. (2012). Environmental control of plant nuclear gene expression by chloroplast redox signals. FrontPlant Sci 3, 257.
Philips, A., Milanowska, K., Lach, G., and Bujnicki, J.M. (2013). LigandRNA: computational predictor of RNA-ligand interactions. RNA 19, 1605-1616.
Pinker, F., Bonnard, G., Gobert, A., Gutmann, B., Hammani, K., Sauter, C., Gegenheimer, P.A., and Giegé, P. (2013). PPR proteins shed a new light on RNase P biology. RNA Biol 10, 1457-1468.
Pribil, M., Labs, M., and Leister, D. (2014). Structure and dynamics of thylakoids in land plants. J Exp Bot 65, 1955-1972.
Prikryl, J., Rojas, M., Schuster, G., and Barkan, A. (2011). Mechanism of RNA stabilization and translational activation by a pentatricopeptide repeat protein. Proc Natl Acad Sci U S A 108, 415-420.
Qu, X., Lancaster, L., Noller, H.F., Bustamante, C., and Tinoco, I., Jr. (2012). Ribosomal protein S1 unwinds double-stranded RNA in multiple steps. Proc Natl Acad Sci U S A 109, 14458-14463.
Reuveni, S., Ehrenberg, M., and Paulsson, J. (2017). Ribosomes are optimized for autocatalytic production. Nature 547, 293-297.
Ries, F., Carius, Y., Rohr, M., Gries, K., Keller, S., Lancaster, C.R.D., and Willmund, F. (2017). Structural and molecular comparison of bacterial and eukaryotic trigger factors. Sci Rep 7, 10680.
Rochaix, J.D. (2007). Role of thylakoid protein kinases in photosynthetic acclimation. FEBS Lett 581, 2768-2775.
Rodermel, S., Haley, J., Jiang, C.Z., Tsai, C.H., and Bogorad, L. (1996). A mechanism for intergenomic integration: abundance of ribulose bisphosphate carboxylase small-subunit protein influences the translation of the large-subunit mRNA. Proc Natl Acad Sci U S A 93, 3881-3885.
Rodermel, S.R., Abbott, M.S., and Bogorad, L. (1988). Nuclear-organelle interactions: nuclear antisense gene inhibits ribulose bisphosphate carboxylase enzyme levels in transformed tobacco plants. Cell 55, 673-681.
48
Rogalski, M., Schöttler, M.A., Thiele, W., Schulze, W.X., and Bock, R. (2008). Rpl33, a nonessential plastid-encoded ribosomal protein in tobacco, is required under cold stress conditions. Plant Cell 20, 2221-2237.
Röhl, T., and van Wijk, K.J. (2001). In vitro reconstitution of insertion and processing of cytochrome f in a homologous chloroplast translation system. J Biol Chem 276, 35465-35472.
Rolland, N., Janosi, L., Block, M.A., Shuda, M., Teyssier, E., Miege, C., Cheniclet, C., Carde, J.P., Kaji, A., and Joyard, J. (1999). Plant ribosome recycling factor homologue is a chloroplastic protein and is bactericidal in escherichia coli carrying temperature-sensitive ribosome recycling factor. Proc Natl Acad Sci U S A 96, 5464-5469.
Rose, R.J., and Lindbeck, A.G.C. (1982). Morphological Studies on the Transcription of Spinach Chloroplast DNA. Zeitschrift für Pflanzenphysiologie 106, 129-137.
Rott, M., Martins, N.F., Thiele, W., Lein, W., Bock, R., Kramer, D.M., and Schöttler, M.A. (2011). ATP synthase repression in tobacco restricts photosynthetic electron transport, CO2 assimilation, and plant growth by overacidification of the thylakoid lumen. Plant Cell 23, 304-321.
Roy, L.M., and Barkan, A. (1998). A SecY homologue is required for the elaboration of the chloroplast thylakoid membrane and for normal chloroplast gene expression. The Journal of cell biology 141, 385-395.
Ruf, M., and Kössel, H. (1988). Occurrence and spacing of ribosome recognition sites in mRNAs of chloroplasts from higher plants. FEBS Letters 240, 41-44.
Russell, J.B., and Cook, G.M. (1995). Energetics of bacterial growth: balance of anabolic and catabolic reactions. Microbiol Rev 59, 48-62.
Ruwe, H., Kupsch, C., Teubner, M., and Schmitz-Linneweber, C. (2011). The RNA-recognition motif in chloroplasts. J Plant Physiol 168, 1361-1371.
Sauert, M., Temmel, H., and Moll, I. (2015). Heterogeneity of the translational machinery: Variations on a common theme. Biochimie 114, 39-47.
Scharff, L.B., Childs, L., Walther, D., and Bock, R. (2011). Local absence of secondary structure permits translation of mRNAs that lack ribosome-binding sites. PLoS Genet 7, e1002155.
Scharff, L.B., Ehrnthaler, M., Janowski, M., Childs, L.H., Hasse, C., Gremmels, J., Ruf, S., Zoschke, R., and Bock, R. (2017). Shine-Dalgarno Sequences Play an Essential Role in the Translation of Plastid mRNAs in Tobacco. Plant Cell.
Schmitz-Linneweber, C., Tillich, M., Herrmann, R.G., and Maier, R.M. (2001). Heterologous, splicing-dependent RNA editing in chloroplasts: allotetraploidy provides trans-factors. EMBO J 20, 4874-4883.
Schmitz-Linneweber, C., Kushnir, S., Babiychuk, E., Poltnigg, P., Herrmann, R.G., and Maier, R.M. (2005b). Pigment deficiency in nightshade/tobacco cybrids is caused by the failure to edit the plastid ATPase alpha-subunit mRNA. Plant Cell 17, 1815-1828.
Schöttler, M.A., Albus, C.A., and Bock, R. (2011). Photosystem I: its biogenesis and function in higher plants. J Plant Physiol 168, 1452-1461.
Schöttler, M.A., Toth, S.Z., Boulouis, A., and Kahlau, S. (2015). Photosynthetic complex stoichiometry dynamics in higher plants: biogenesis, function, and turnover of ATP synthase and the cytochrome b6f complex. J Exp Bot 66, 2373-2400.
Schröter, Y., Steiner, S., Matthai, K., and Pfannschmidt, T. (2010). Analysis of oligomeric protein complexes in the chloroplast sub-proteome of nucleic acid-binding proteins from mustard reveals potential redox regulators of plastid gene expression. Proteomics 10, 2191-2204.
Schult, K., Meierhoff, K., Paradies, S., Töller, T., Wolff, P., and Westhoff, P. (2007). The nuclear-encoded factor HCF173 is involved in the initiation of translation of the psbA mRNA in Arabidopsis thaliana. Plant Cell 19, 1329-1346.
Schünemann, D. (2007). Mechanisms of protein import into thylakoids of chloroplasts. Biol Chem 388, 907-915.
Schwarz, C., Bohne, A.V., Wang, F., Cejudo, F.J., and Nickelsen, J. (2012). An intermolecular disulfide-based light switch for chloroplast psbD gene expression in Chlamydomonas reinhardtii. Plant J 72, 378-389.
Serganov, A., and Nudler, E. (2013). A decade of riboswitches. Cell 152, 17-24.
49
Shajani, Z., Sykes, M.T., and Williamson, J.R. (2011). Assembly of bacterial ribosomes. Annu Rev Biochem 80, 501-526.
Sharma, M.R., Wilson, D.N., Datta, P.P., Barat, C., Schluenzen, F., Fucini, P., and Agrawal, R.K. (2007). Cryo-EM study of the spinach chloroplast ribosome reveals the structural and functional roles of plastid-specific ribosomal proteins. Proc Natl Acad Sci U S A 104, 19315-19320.
Sharma, M.R., Dönhöfer, A., Barat, C., Marquez, V., Datta, P.P., Fucini, P., Wilson, D.N., and Agrawal, R.K. (2010). PSRP1 is not a ribosomal protein, but a ribosome-binding factor that is recycled by the ribosome-recycling factor (RRF) and elongation factor G (EF-G). J Biol Chem 285, 4006-4014.
Sharpe, R.M., Mahajan, A., Takacs, E.M., Stern, D.B., and Cahoon, A.B. (2011). Developmental and cell type characterization of bundle sheath and mesophyll chloroplast transcript abundance in maize. Curr Genet 57, 89-102.
Shi, Z., Fujii, K., Kovary, K.M., Genuth, N.R., Rost, H.L., Teruel, M.N., and Barna, M. (2017). Heterogeneous Ribosomes Preferentially Translate Distinct Subpools of mRNAs Genome-wide. Mol Cell 67, 71-83 e77.
Shine, J., and Dalgarno, L. (1974). The 3'-terminal sequence of Escherichia coli 16S ribosomal RNA: complementarity to nonsense triplets and ribosome binding sites. Proc Natl Acad Sci U S A 71, 1342-1346.
Shteiman-Kotler, A., and Schuster, G. (2000). RNA-binding characteristics of the chloroplast S1-like ribosomal protein CS1. Nucleic Acids Res 28, 3310-3315.
Sijben-Müller, G., Hallick, R.B., Alt, J., Westhoff, P., and Herrmann, R.G. (1986). Spinach plastid genes coding for initiation factor IF-1, ribosomal protein S11 and RNA polymerase alpha-subunit. Nucleic Acids Res 14, 1029-1044.
Singh, B.N., Mishra, R.N., Agarwal, P.K., Goswami, M., Nair, S., Sopory, S.K., and Reddy, M.K. (2004). A pea chloroplast translation elongation factor that is regulated by abiotic factors. Biochem Biophys Res Commun 320, 523-530.
Singh, R.N., Saldanha, R.J., D'Souza, L.M., and Lambowitz, A.M. (2002). Binding of a group II intron-encoded reverse transcriptase/maturase to its high affinity intron RNA binding site involves sequence-specific recognition and autoregulates translation. J Mol Biol 318, 287-303.
Sosso, D., Canut, M., Gendrot, G., Dedieu, A., Chambrier, P., Barkan, A., Consonni, G., and Rogowsky, P.M. (2012). PPR8522 encodes a chloroplast-targeted pentatricopeptide repeat protein necessary for maizeembryogenesis and vegetative development. J Exp Bot 63, 5843-5857.
Sreedharan, S.P., Beck, C.M., and Spremulli, L.L. (1985). Euglena gracilis chloroplast elongation factor Tu. Purification and initial characterization. J Biol Chem 260, 3126-3131.
Staub, J.M., and Maliga, P. (1994). Translation of psbA mRNA is regulated by light via the 5'-untranslated region in tobacco plastids. Plant J 6, 547-553.
Staub, J.M., and Maliga, P. (1995). Expression of a chimeric uidA gene indicates that polycistronic mRNAs are efficiently translated in tobacco plastids. Plant J 7, 845-848.
Stern, D.B., Harris, E.H., and Witman, G.B., eds. (2009). The Chlamydomonas Sourcebook (Second Edition); (London: Academic Press).
Stoppel, R., Lezhneva, L., Schwenkert, S., Torabi, S., Felder, S., Meierhoff, K., Westhoff, P., and Meurer, J. (2011). Recruitment of a ribosomal release factor for light- and stress-dependent regulation of petB transcript stability in Arabidopsis chloroplasts. Plant Cell 23, 2680-2695.
Sugiura, M. (1995). The chloroplast genome. Essays Biochem 30, 49-57. Sugiura, M. (2014). Plastid mRNA translation. Methods Mol Biol 1132, 73-91. Sugliani, M., Abdelkefi, H., Ke, H., Bouveret, E., Robaglia, C., Caffarri, S., and Field, B. (2016). An
Ancient Bacterial Signaling Pathway Regulates Chloroplast Function to Influence Growth and Development in Arabidopsis. Plant Cell 28, 661-679.
Sun, Y., and Zerges, W. (2015). Translational regulation in chloroplasts for development and homeostasis. Biochim Biophys Acta 1847, 809-820.
Sundberg, E., Slagter, J.G., Fridborg, I., Cleary, S.P., Robinson, C., and Coupland, G. (1997). ALBINO3, an Arabidopsis nuclear gene essential for chloroplast differentiation, encodes a chloroplast protein that shows homology to proteins present in bacterial membranes and yeast mitochondria. Plant Cell 9, 717-730.
50
Supek, F. (2016). The Code of Silence: Widespread Associations Between Synonymous Codon Biases and Gene Function. J Mol Evol 82, 65-73.
Suzuki, H., and Morton, B.R. (2016). Codon Adaptation of Plastid Genes. PLoS One 11, e0154306. Taniguchi, M., Kuroda, H., and Satoh, K. (1993). ATP-dependent protein synthesis in isolated pea
chloroplasts. Evidence for accumulation of a translation intermediate of the D1 protein. FEBS Lett 317, 57-61.
Teubner, M., Fuß, J., Kühn, K., Krause, K., and Schmitz-Linneweber, C. (2017). The RNA recognition motif protein CP33A is a global ligand of chloroplast mRNAs and is essential for plastid biogenesis and plant development. Plant J 89, 472-485.
Tiller, N., and Bock, R. (2014). The translational apparatus of plastids and its role in plant development. Mol Plant 7, 1105-1120.
Tiller, N., Weingartner, M., Thiele, W., Maximova, E., Schöttler, M.A., and Bock, R. (2012). The plastid-specific ribosomal proteins of Arabidopsis thaliana can be divided into non-essential proteins and genuine ribosomal proteins. Plant J 69, 302-316.
Timmis, J.N., Ayliffe, M.A., Huang, C.Y., and Martin, W. (2004). Endosymbiotic gene transfer: organelle genomes forge eukaryotic chromosomes. Nat Rev Genet 5, 123-135.
Trösch, R., Mühlhaus, T., Schroda, M., and Willmund, F. (2015). ATP-dependent molecular chaperones in plastids--More complex than expected. Biochim Biophys Acta 1847, 872-888.
Tzvetkova-Chevolleau, T., Hutin, C., Noël, L.D., Goforth, R., Carde, J.P., Caffarri, S., Sinning, I., Groves, M., Teulon, J.M., Hoffman, N.E., Henry, R., Havaux, M., and Nussaume, L. (2007). Canonical signal recognition particle components can be bypassed for posttranslational protein targeting in chloroplasts. Plant Cell 19, 1635-1648.
Ude, S., Lassak, J., Starosta, A.L., Kraxenberger, T., Wilson, D.N., and Jung, K. (2013). Translation elongation factor EF-P alleviates ribosome stalling at polyproline stretches. Science 339, 82-85.
Udy, D.B., Belcher, S., Williams-Carrier, R., Gualberto, J.M., and Barkan, A. (2012). Effects of reduced chloroplast gene copy number on chloroplast gene expression in maize. Plant Physiol 160, 1420-1431.
van Wijk, K.J., Andersson, B., and Aro, E.M. (1996). Kinetic resolution of the incorporation of the D1 protein into photosystem II and localization of assembly intermediates in thylakoid membranes of spinach chloroplasts. J Biol Chem 271, 9627-9636.
Verhounig, A., Karcher, D., and Bock, R. (2010). Inducible gene expression from the plastid genome by a synthetic riboswitch. Proc Natl Acad Sci U S A 107, 6204-6209.
Vila-Sanjurjo, A., Schuwirth, B.S., Hau, C.W., and Cate, J.H. (2004). Structural basis for the control of translation initiation during stress. Nat Struct Mol Biol 11, 1054-1059.
Voelker, R., Mendel-Hartvig, J., and Barkan, A. (1997). Transposon-disruption of a maize nuclear gene, tha1, encoding a chloroplast SecA homologue: in vivo role of cp-SecA in thylakoid protein targeting. Genetics 145, 467-478.
Walter, B., Hristou, A., Nowaczyk, M.M., and Schünemann, D. (2015). In vitro reconstitution of co-translational D1 insertion reveals a role of the cpSec-Alb3 translocase and Vipp1 in photosystem II biogenesis. Biochem J 468, 315-324.
Walter, M., Piepenburg, K., Schöttler, M.A., Petersen, K., Kahlau, S., Tiller, N., Drechsel, O., Weingartner, M., Kudla, J., and Bock, R. (2010). Knockout of the plastid RNase E leads to defective RNA processing and chloroplast ribosome deficiency. Plant J 64, 851-863.
Whitfeld, P.R., Leaver, C.J., Bottomley, W., and Atchison, B.A. (1978). Low-molecular-weight (4.5S) ribonucleic acid in higher-plant chloroplast ribosomes. Biochemical Journal 175, 1103-1112.
Williams, C.C., Jan, C.H., and Weissman, J.S. (2014). Targeting and plasticity of mitochondrial proteins revealed by proximity-specific ribosome profiling. Science 346, 748-751.
Wolin, S.L., and Walter, P. (1988). Ribosome pausing and stacking during translation of a eukaryotic mRNA. EMBO J 7, 3559-3569.
Woolstenhulme, C.J., Parajuli, S., Healey, D.W., Valverde, D.P., Petersen, E.N., Starosta, A.L., Guydosh, N.R., Johnson, W.E., Wilson, D.N., and Buskirk, A.R. (2013). Nascent peptides that block protein synthesis in bacteria. Proc Natl Acad Sci U S A 110, E878-887.
Wostrikoff, K., and Stern, D. (2007). Rubisco large-subunit translation is autoregulated in response to its assembly state in tobacco chloroplasts. Proc Natl Acad Sci U S A 104, 6466-6471.
51
Wostrikoff, K., Choquet, Y., Wollman, F.A., and Girard-Bascou, J. (2001). TCA1, a single nuclear-encoded translational activator specific for petA mRNA in Chlamydomonas reinhardtii chloroplast. Genetics 159, 119-132.
Yamaguchi, K., and Subramanian, A.R. (2000). The plastid ribosomal proteins. Identification of all the proteins in the 50 S subunit of an organelle ribosome (chloroplast). J Biol Chem 275, 28466-28482.
Yamaguchi, K., and Subramanian, A.R. (2003). Proteomic identification of all plastid-specific ribosomal proteins in higher plant chloroplast 30S ribosomal subunit. Eur J Biochem 270, 190-205.
Yamaguchi, K., von Knoblauch, K., and Subramanian, A.R. (2000). The plastid ribosomal proteins. Identification of all the proteins in the 30 S subunit of an organelle ribosome (chloroplast). J Biol Chem 275, 28455-28465.
Yamamoto, H., Wittek, D., Gupta, R., Qin, B., Ueda, T., Krause, R., Yamamoto, K., Albrecht, R., Pech, M., and Nierhaus, K.H. (2016). 70S-scanning initiation is a novel and frequent initiation mode of ribosomal translation in bacteria. Proc Natl Acad Sci U S A 113, E1180-1189.
Yamamoto, T., Burke, J., Autz, G., and Jagendorf, A.T. (1981). Bound Ribosomes of Pea Chloroplast Thylakoid Membranes: Location and Release in Vitro by High Salt, Puromycin, and RNase. Plant Physiol 67, 940-949.
Yamazaki, H., Tasaka, M., and Shikanai, T. (2004). PPR motifs of the nucleus-encoded factor, PGR3, function in the selective and distinct steps of chloroplast gene expression in Arabidopsis. Plant J 38, 152-163.
Yosef, I., Irihimovitch, V., Knopf, J.A., Cohen, I., Orr-Dahan, I., Nahum, E., Keasar, C., and Shapira, M. (2004). RNA binding activity of the ribulose-1,5-bisphosphate carboxylase/oxygenase large subunitfrom Chlamydomonas reinhardtii. J Biol Chem 279, 10148-10156.
Yukawa, M., and Sugiura, M. (2008). Termination codon-dependent translation of partially overlapping ndhC-ndhK transcripts in chloroplasts. Proc Natl Acad Sci U S A 105, 19550-19554.
Yukawa, M., Kuroda, H., and Sugiura, M. (2007). A new in vitro translation system for non-radioactive assay from tobacco chloroplasts: effect of pre-mRNA processing on translation in vitro. Plant J 49, 367-376.
Zhan, Y., Dhaliwal, J.S., Adjibade, P., Uniacke, J., Mazroui, R., and Zerges, W. (2015). Localized control of oxidized RNA. J Cell Sci 128, 4210-4219.
Zhang, L., and Aro, E.M. (2002). Synthesis, membrane insertion and assembly of the chloroplast-encoded D1 protein into photosystem II. FEBS Lett 512, 13-18.
Zhang, L., Paakkarinen, V., van Wijk, K.J., and Aro, E.M. (1999). Co-translational assembly of the D1 protein into photosystem II. J Biol Chem 274, 16062-16067.
Zhang, L., Paakkarinen, V., Suorsa, M., and Aro, E.M. (2001). A SecY homologue is involved in chloroplast-encoded D1 protein biogenesis. J Biol Chem 276, 37809-37814.
Zheng, M., Liu, X., Liang, S., Fu, S., Qi, Y., Zhao, J., Shao, J., An, L., and Yu, F. (2016). Chloroplast Translation Initiation Factors Regulate Leaf Variegation and Development. Plant Physiol 172, 1117-1130.
Zhou, F., Karcher, D., and Bock, R. (2007). Identification of a plastid intercistronic expression element (IEE) facilitating the expression of stable translatable monocistronic mRNAs from operons. Plant J 52, 961-972.
Ziehe, D., Dünschede, B., and Schünemann, D. (2017). From bacteria to chloroplasts: evolution of the chloroplast SRP system. Biol Chem, 653-661.
Zito, F., Kuras, R., Choquet, Y., Kössel, H., and Wollman, F.A. (1997). Mutations of cytochrome b6 in Chlamydomonas reinhardtii disclose the functional significance for a proline to leucine conversion by petB editing in maize and tobacco. Plant Mol Biol 33, 79-86.
Zoschke, R., and Barkan, A. (2015). Genome-wide analysis of thylakoid-bound ribosomes in maize reveals principles of cotranslational targeting to the thylakoid membrane. Proc Natl Acad Sci U S A 112, E1678-E1687.
Zoschke, R., Watkins, K.P., and Barkan, A. (2013a). A rapid ribosome profiling method elucidates chloroplast ribosome behavior in vivo. Plant Cell 25, 2265-2275.
Zoschke, R., Chotewutmontri, P., and Barkan, A. (2017). Translation and Co-translational Membrane Engagement of Plastid-encoded Chlorophyll-binding Proteins Are Not Influenced by Chlorophyll Availability in Maize. Front Plant Sci 8, 385.
52
Zoschke, R., Watkins, K.P., Miranda, R.G., and Barkan, A. (2016). The PPR-SMR protein PPR53 enhances the stability and translation of specific chloroplast RNAs in maize. Plant J 85, 594-606.
Zoschke, R., Qu, Y., Zubo, Y.O., Börner, T., and Schmitz-Linneweber, C. (2013b). Mutation of the pentatricopeptide repeat-SMR protein SVR7 impairs accumulation and translation of chloroplast ATP synthase subunits in Arabidopsis thaliana. J Plant Res 126, 403-414.
Zoschke, R., Nakamura, M., Liere, K., Sugiura, M., Börner, T., and Schmitz-Linneweber, C. (2010). An organellar maturase associates with multiple group II introns. Proc Natl Acad Sci U S A 107, 3245-3250.
Zoschke, R., Kroeger, T., Belcher, S., Schöttler, M.A., Barkan, A., and Schmitz-Linneweber, C. (2012). The pentatricopeptide repeat-SMR protein ATP4 promotes translation of the chloroplast atpB/E mRNA. Plant J 72, 547-558.
Zybailov, B., Rutschow, H., Friso, G., Rudella, A., Emanuelsson, O., Sun, Q., and van Wijk, K.J. (2008). Sorting signals, N-terminal modifications and abundance of the chloroplast proteome. PLoS One 3, e1994.
DOI 10.1105/tpc.18.00016; originally published online April 2, 2018;Plant CellReimo Zoschke and Ralph Bock
RegulationChloroplast Translation: Structural and Functional Organization, Operational Control and
This information is current as of July 16, 2018
Permissions https://www.copyright.com/ccc/openurl.do?sid=pd_hw1532298X&issn=1532298X&WT.mc_id=pd_hw1532298X
eTOCs http://www.plantcell.org/cgi/alerts/ctmain
Sign up for eTOCs at:
CiteTrack Alerts http://www.plantcell.org/cgi/alerts/ctmain
Sign up for CiteTrack Alerts at:
Subscription Information http://www.aspb.org/publications/subscriptions.cfm
is available at:Plant Physiology and The Plant CellSubscription Information for
ADVANCING THE SCIENCE OF PLANT BIOLOGY © American Society of Plant Biologists