Journal of Experimental Botany, Vol. 66, No. 19 pp. 5639–5650, 2015
doi:10.1093/jxb/erv298 Advance Access publication 15 June 2015
COMMENTARY
Implications of mutation of organelle genomes for organelle
function and evolution
John A. Raven*
Division of Plant Sciences, University of Dundee at the James Hutton Institute, Invergowrie, Dundee DD2 5DA, UK †
School of Plant Biology, University of Western Australia, M048, 35 Stirling Highway, Crawley, WA 6009, Australia
* To whom correspondence should be addressed. E-mail: [email protected]
† Permanent address of JAR.
Received 28 January 2015; Revised 28 April 2015; Accepted 22 May 2015
Editor: Christine Raines
Abstract
Organelle genomes undergo more variation, including that resulting from damage, than eukaryotic nuclear genomes,
or bacterial genomes, under the same conditions. Recent advances in characterizing the changes to genomes of
chloroplasts and mitochondria of Zea mays should, when applied more widely, help our understanding of how damage to organelle genomes relates to how organelle function is maintained through the life of individuals and in succeeding generations. Understanding of the degree of variation in the changes to organelle DNA and its repair among
photosynthetic organisms might help to explain the variations in the rate of nucleotide substitution among organelle
genomes. Further studies of organelle DNA variation, including that due to damage and its repair might also help us
to understand why the extent of DNA turnover in the organelles is so much greater than that in their bacterial (cyanobacteria for chloroplasts, proteobacteria for mitochondria) relatives with similar rates of production of DNA-damaging
reactive oxygen species. Finally, from the available data, even the longest-lived organelle-encoded proteins, and the
RNAs needed for their synthesis, are unlikely to maintain organelle function for much more than a week after the
complete loss of organelle DNA.
Key words: Chloroplast, DNA damage, DNA repair, evolution, mitochondrion, mutation, phylogeny, protein turnover, reactive
oxygen species.
Introduction
After significant resistance from many biologists, the
endosymbiotic origin of chloroplasts from cyanobacteria (Mereschowsky, 1905; Schmitz, 1982; Schimper, 1983;
Martin and Kowallik, 1999), and then the endosymbiotic
origin of mitochondria from (proteo)bacteria (Wallin, 1923),
is now almost universally accepted (O’Malley, 2015). One,
as yet, incompletely explained aspect of these energy-transducing organelles is the retention of an organelle genome in
all chloroplasts and in all mitochondria, apart from some
hydrogenosomes. The organelle genome houses a small
minority of the genes needed for organelle synthesis and function, the remainder having been transferred in evolution to
the nucleus with translation on cytosol ribosomes and import
of the resulting proteins into the organelle. The fraction of
the organelle proteome encoded by the organelle genome
has been estimated at 2.2% for Arabidopsis mitochondria
(Heazlewood et al., 2004) and 3.5–5.3% for Arabidopsis chloroplasts (Abdellah et al., 2000; Kleffman et al., 2004). Allen
© The Author 2015. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved.
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5640 | Raven
and Raven (1996), Martin et al. (1998), and Race et al. (1999)
discuss some of the reasons for the retention of genes in the
organelle (e.g. the control of gene expression by the organelle
redox status: Allen, 1993a, b; Allen and Raven, 1996; Allen,
2003; Allen et al., 2005) and for their relocation to the nucleus
(e.g. the damage to DNA by the reactive oxygen species and
other free radicals generated in the redox reactions of the
organelles: Allen and Raven, 1996).
Regardless of the reasons for the retention of genes in organelles, it is clear that changes in organelle DNA, including
that resulting from oxidative damage, is greater than in the
corresponding nuclear DNA, or bacterial or archaean DNA,
under as nearly similar conditions as possible. However, much
remains to be discovered about damage and other causes of
changes to organelle DNA and its significance for short-term
functioning of the organelles, their transmission to the next
generation, and evolution. Many of the impediments to our
understanding are technical, and Kumar et al. (2014) make
significant experimental advances that help to bring into better focus some aspects of what remains to be discovered.
Kumar et al. (2014) used two PCR-based techniques on chloroplast and mitochondrial genomes of developing Zea mays.
The novel method is ‘molecular integrity PCR’ (miPCR) that
quantifies long organelle DNA molecules that have no impediment to amplification and, hence, presumably, are functional
in vivo in transcription and thence translation. The standard
qPCR method gives the total number of copies, regardless of
whether they could be functional in vivo in transcription and
thus translation. They also measured ‘impediments’, including oxidative damage, using an in vitro DNA repair assay, as
a check on impediments estimated from the qPCR methods.
Part of the background to the work of Kumar et al. (2014)
is the reported loss of all genomes from organelles during nonreproductive development in some organisms; there are two to
many genomes per organelle early in cell development (Tables
1, 2). The first example to be reported is non-reproductive
Table 1. Copy number of mitochondrial and chloroplast genomes in Archeoplastida, i.e. with chloroplasts derived from a single primary
endosymbiosis
The general assumption is that the smallest unit detectable is the complete genome, or genome in process of replication, in the DAPI
experiments of Kuroiwa et al. (1981).
OrganismGlaucophyta
Rhodophyta
Cyanidiophyceae
Rhodophyta
Bangiophyceae
Rhodophyta
Florideophyceae
Chlorophyta
Prasinophyceae
Chlorophyta
Chlorophyceae
Chlorophyta
Trebouxiophyceae
Chlorophyta
Ulvophyceae
Chlorophyta
Charophyceae
Streptophyta
Embryophytes:Bryophyte (moss)
Streptophyta
Embryophytes: euphyllophyte:
Pteridophyte
Streptophyta
Embryophytes; euphyllophyte:
Glymnosperms
Streptophyta
Embryophytes: euphyllophyte
Angiosperms
Mitochondrial genome copy number
24
Chloroplast genome copy number
References
1
Kuroiwa et al. (1981)
1–9
15–26
Kuroiwa et al. (1981)
Hirakawa and Ishida (2014)
Kuroiwa et al. (1981)
5–50
Kuroiwa et al. (1981)
20–30
Kuroiwa et al. (1981)
2–80
Kuroiwa et al. (1981)
Hirakawa and Ishida (2014)
Kuroiwa et al. (1981)
2–50
0–156
15–32
Kuroiwa et al. (1981)
Lűttke (1988)
Kuroiwa et al. (1981)
Hirakawa and Ishida (2014)
Kuroiwa et al. (1981)
6–23
Kuroiwa et al. (1981)
8–16
Kuroiwa et al. (1981)
0–900
Lamppa et al. (1980)
Kuroiwa et al. (1981)
Bendich and Gauriloff (1984)
Bendich (1987)
Kuroiwa et al. (1990)
Rowan and Bendich (2009)
Preuten et al. (2010)
Wang et al. (2010)
10–300
0–2
Organelle genomes for organelle function and evolution | 5641
Table 2. Copy number of mitochondrial and chloroplast genomes in photosynthetic organisms with plastids from secondary or tertiary
endosymbiosis
Organism
Discicristates
Euglenophyta
Rhizarians
Chlorarachniophyta
Chromalveolates
Ochrophyta: Bacillariophyceae
Chromalveolates
Ochrophyta: Phaeophyceae
Chromalveolates
Ochrophyta: Raphidophyceae
Chromalveolates
Cryptophyta: Cryptophyceae
Mitochondrial genome copy number
18–40
24–43
Chloroplast genome copy number
References
20–34
Kuroiwa et al. (1981)
30–50
Hirakawa and Ishida (2014)
1a
Kuroiwa et al. (1981)
1a–16
Kuroiwa et al. (1981)
10–40
Ersland et al. (1981)
22–260
Kuroiwa et al. (1981)
Hirakawa and Ishida (2014)
a
While the general assumption is that the smallest unit detectable is the complete genome, or a genome in process of replication, in the DAPI
experiments of Kuroiwa et al. (1981), there are clear exceptions. In many of the Ochrophyta the chloroplasts genome was recognized as a
peripheral band within the chloroplast that was itself made up of the smallest detectable units (Kuroiwa et al., 1981).
development in the acellular green macroalga Acetabularia
(Dasycladales; Ulvophyceae; Chlorophyta) (Woodcock
and Bogorad, 1970; Coleman, 1979; Lüttke and Bonotto,
1981; Lüttke, 1988) using histochemical staining of DNA.
Somewhat later, such a loss of chloroplast and mitochondrial
genomes was reported, mainly using the more quantitative
molecular genetic technique, in vegetative (somatic) cells
in the developing leaves of flowering plants (Bendich and
Gauriloff, 1984; Sodmergen et al., 1991; Rowan et al., 2002;
Oldenburg and Bendich, 2004; Shaver et al., 2006; Preuten
et al., 2010; Wang et al., 2010; Oldenburg et al., 2013), with
changes in the integrity of the organelle DNA when it is
retained (Oldenburg and Bendich, 2004; Shaver et al., 2006;
Oldenburg et al., 2013). These conclusions on decreasing content, and even complete loss, of organelle DNA, and changes
in organelle DNA integrity, during somatic cell development
have been cogently challenged (Li et al., 2006; Zotschke et al.,
2007). Possible means of reconciling the different conclusions
resulting from variations in the developmental stage of the
plant organ, and in experimental methods, have been suggested by Rowan and Bendich (2009) and outcomes of the
relevant novel techniques mentioned in the Introduction
(Kumar et al., 2014) are discussed in the following section.
Pending complete resolution of these experimental differences, and the differences in the results obtained, one contribution of the present work is to explore the possibility of continued
functioning of organelles after complete loss of their genomes.
Implications of advances in understanding
organelle DNA integrity and copy number
for understanding organelle genome
function, inheritance, and evolution
Organellar protein and DNA stability
There has been much emphasis on oxidative and UV damage
to DNA, and especially organelle DNA (Allen and Raven,
1996; Halliwell and Gutteridge, 2007) as part of causes of
variation in the DNA. The results of Kumar et al. (2014) are
used as an example of the effects of of photosynthetically
active radiation, with no specific UVB treatment, i.e. mainly
reactive oxygen effects. Of the treatments they used, Kumar
et al. (2014) found most damage, impediment, and in vitro
repair for both chloroplasts and mitochondria in Zea mays
plants grown in a 16/8 light/dark regime for 13 d, with less
damage and impediment in plants in the dark for 12 d followed by 1 d in the light, and even less in plants in the dark
for the full 13 d. The only difference between effects on chloroplasts and mitochondria occurs in light-grown plants. The
qPCR-derived copy number on chloroplasts grown in the
light is high, in agreement with measurements of damage,
degree of impediment, fraction of unimpeded DNA from
miPCR and in vitro repair, while in mitochondria the qPCR
copy number is low in the light-grown plants, contrasting
with the outcomes of the other methods.
Oxidative and UV damage occurs to proteins as well as to
DNA, and the relative roles of these of damage to the two
macromolecules in causing cell death in environmentally
challenged bacteria is, as discussed below, still a matter of
debate (Daly et al., 2010). For the maintenance of organelle
function over a significant fraction of the cell lifespan there is
a need for the replacement of organelle- and nuclear-encoded
proteins that have evaded the protective mechanisms (e.g.
non-enzymic free radical scavengers, superoxide dismutase,
ascorbate peroxidase) and have been damaged by oxidation
and UV. This protein replacement requires the presence of
organelle DNA and the RNAs and proteins associated with
transcription and translation, or of long-lived mRNA and
the RNAs and proteins associated with translation.
Organelle-encoded proteins clearly do turn over, as shown for
mitochondria of Saccharomyces cerevisae (de Jong et al., 2000;
data on wild type) and chloroplasts of algae and plants (Table 3a).
The half-lives for mitochondria-encoded proteins range from
185–415 h (Table 3a) and for chloroplast-encoded proteins range
from 1.5–415 h (Table 3b). If the faster rates of breakdown also
occurred in chloroplasts lacking DNA where, in the absence of
5642 | Raven
Table 3a. Half-lives of mitochondrial genome-encoded RNAs and proteins
Organism
t1/2 of RNA (h)
Arabidopsis thaliana
(11-fold range; no
absolute values)
t1/2of protein (h)
References
Giegé et al. (2000)
nad7 (complex I) 208
nad9 (complex I), 185
cox2 (complex IV), 185
atp8 (complex V) 238
ATP synthase (complex V),
mean for mitochondrial- and
nuclear-encoded subunits 415
Arabidopsis thaliana
Hordeum vulgare
Nelson et al. (2013)
Nelson et al. (2014)
Table 3b. Half-lives of chloroplast genome-encoded RNAs and proteins
Organism
t1/2 of RNA (h)
Spinacia oleracea
psbA mRNA 5 (young),
10 (mature)
rbcL mRNA 5 (young or mature)
tRNA(lys) unspliced precursor 3
16S rRNA 20–40
psbA mRNA 40
atpH mRNA, 11–45
psbC mRNA, 11–19
psbA mRNA 33
rbcL mRNA 5.2–14
Hordeum vulgare
Arabidopsis thaliana
Chlamydomonas
reinhardtii
Brassica napus
Chlamydomonas
reinhardtii
Ostreococcus taurii
Hordeum vulgare
long-lived mRNA (see below), there could be no replacement
of degraded protein, chloroplast function would be significantly
decreased after a day and would be very limited after a few days.
For mitochondria, there is also the maintenance of function after the loss of mitochondrial DNA. Using the qPCR
method (Kumar et al., 2014), Preuten et al. (2010) showed
that there were fewer copies of individual mitochondrial
t1/2 of protein(h)
Reference
Klaff and Gruisson (1991)
Kim et al. (1993)
Germain et al. (2012)
psbA gene product
(D1 protein), 1.5
psbA gene product
(D1 protein) 2
petA gene product
(cytochrome f) >21
petB gene product
(cytochrome b6) >21
petD gene product
(subunit IV of cytochrome
b6-f complex) 20
ATP synthase
components, 119–165
Core PSII proteins
(not psbA), 136–204
rbcL, 147)
Mean of chloroplastand nuclear-encoded
subunits of:
ATPsynthase, 116
Photosystem II
(excluding psbA), 116
Photosystem I
(excluding psaP) 415
psbA gene product
(D1 protein), 17
Schuster et al. (1988)
Sundberg et al. (1993)
Gong et al. (2001);
Mastrobuoni et al. (2012)
Martin et al. (2012
Nelson et al. (2014)
genes than of mitochondria in each mature cell. However,
respiration rates are independent of mitochondrial gene
number and, presumably, mitochondrial fission and fusion
‘share’ the genes among the mitochondria that can be
counted at a particular time (Preuten et al., 2010). This is less
likely for chloroplasts where fusion is very rare (CavalierSmith, 1970; Seguí-Simarro and Staehlin, 2009).
Organelle genomes for organelle function and evolution | 5643
Table 3c. Half-lives of nuclear-encoded chloroplast proteins
Organism
t1/2 of protein
Reference
Zea mays
Arabidopsis thaliana
Ostrecoccus tauri
Nicotiana tabaccum
Rubisco (rbcL+rbcS), 168 h
Simpson et al. (1981);
Piques et al. (2009)
Martin et al. (2012)
Krech et al. (2012)
Nicotiana tabaccum
Hordeum vulgare
rbcS 301 h
Nuclear and plastid-encoded
components of the PSI complex:
‘highly stable’
Nuclear and plastid-encoded
components of the cytochrome
b6-f complex: ‘lifetime at least
one week’
Mean of chloroplast- and
nuclear-encoded subunits of:
ATPsynthase, 116
Photosystem II (excluding
psbA), 116
Photosystem I (excluding
psaA) 415
psbA gene product (D1 protein,
chloroplast encoded), 17
Hojka et al. (2014)
Nelson et al. (2014)
Chloroplasts and mitochondria lacking DNA could retain
synthesis of organelle-encoded proteins if the mRNAs
for the proteins are long-lived and, of course, so would all
of the post-translational components synthesis apparatus.
Retention of the RNA and protein components needed for
protein synthesis can be compromised by oxidative and UV
damage, as is the case for organelle DNA and the organelleencoded proteins involved in functions other than protein
synthesis. There seem to be no data on absolute rather than
relative half-lives of mitochondria-encoded RNAs of plants
or algae (Table 3a). The half-lives of chloroplast-encoded
RNAs range from 3 h to 45 h (Table 3b). These values cannot be directly related, in the case of mRNAs, to the rate of
decline in the synthesis of chloroplast-encoded proteins since
this protein synthesis is translationally controlled (Fromm
et al., 1985; Inamine et al., 1985; Hosler et al., 1989; Danon,
1997; Gilham et al., 1997; Eberhard et al., 2002, but see Udy
et al., 2012), as is that of mitochondria-encoded proteins
(Gilham et al., 1997; Holec et al., 2006). This means that an
organelle mRNA might go through several decay half-lives
before the amount of transcript becomes limiting for the
rate of protein synthesis. Even with a several-fold increase
in the effective (for protein synthesis) half-life of mRNA, if
the faster rates of breakdown of mRNA and all of the posttranslational components synthesis apparatus also occurred
in flowering plant chloroplasts lacking DNA, chloroplast
function would be significantly decreased after several days
and would be very limited after a week or two. The longest
measured chloroplast mRNA half-life (Table 3b) would give
a half-life of chloroplast proteins similar to the highest values
for protein turnover (Table 3b).
There seem to be no relevant data to determine if the damage to organelle-encoded proteins is more important than
damage to the organelle genome for terminally differentiated
organelles (i.e. not contributing to the next generation) in
multicellular organisms. Daly et al. (2010) have suggested, and
provided evidence, that, even in the unicellular Deinococcus,
damage to proteins is more significant than damage to DNA
in determining survival under extreme conditions (see below).
A parallel example of sustaining protein content in organelles despite damage to proteins and a lack of the encoding
DNA is found for kleptoplastids. These are chloroplasts that
have been retained within their cells by phagotrophs feeding
on algae and remain photosynthetically functional for days
or even months (Raven et al., 2009). In most cases, the nuclei
of the food alga are not retained (e.g. in the relatively wellinvestigated sacoglossan marine gastropods), so that there is
no possibility of replacing damaged nuclear-encoded chloroplast proteins unless there has been horizontal gene transfer from the algal nuclei to the phagotroph nucleus (Raven
et al., 2009). While there have been claims of such horizontal
gene transfer (Raven et al., 2009), the very thorough work
of Wägele et al. (2011) showed that this was very unlikely, so
the enigma of maintenance of the nuclear-encoded fraction
of the chloroplast proteome of kleptoplastids remains. Even
the capacity to express the chloroplast-encoded gene psbA
(encoding the rapidly turning over D1 protein of photosystem II) requires a quality control gene (ftsH) that is nuclearencoded in streptophytes (Charophyta plus Embryophyta)
(de Vries et al., 2013). However, the phylogenetic analysis of
de Vries et al. (2013) shows that ftsH is chloroplast-encoded
in the food algae Acetabularia acetabulum and Vaucheria
litorea, providing the longest-functioning kleptoplastids (in
the sacoglossans Elysia chlorotica and Elysia timida, respectively), so that quality control of D1 can be maintained. This
important finding still does not explain how the many other
nuclear-encoded gene products essential for photosynthesis,
either directly as catalysts of photosynthetic reactions or as
requirements for expression of chloroplast-encoded proteins
(Zerges et al., 1997; Barkan and Goldschidt-Clermont, 2000)
can be maintained in long-lived kleptoplastids. A further
question arises from the use of Acetabularia chloroplasts as
long-lived kleptoplastids, since, during vegetative growth,
some Acetabularia chloroplasts apparently lack genomes
(Lűttke, 1988).
A further aspect of the dependence of kleptoplastids
on the nuclear genome of the source alga concerns the
fate of the glycolate resulting from Rubisco oxygenase
and phosphoglycolate phosphatase in kleptoplastids. The
production of glycolate cannot be suppressed by expression in the sacoglossan of inorganic carbon concentrating mechanisms (CCMs) from the host alga, since these
CCMs typically have extraplastidial components encoded
by algal nuclear genes (Falkowski and Raven, 2007; Raven
et al., 2012, 2014), and so would not be acquired with the
kleptoplastids. In the absence of a CCM (i.e. diffusive CO2
entry) glycolate is produced but cannot be metabolized
since all but the initial and final steps of the photorespiratory carbon oxidation cycle are extraplastidial (Falkowski
and Raven, 2007; Raven et al., 2012, 2014); the intermediate
steps would not be acquired with the kleptoplastids. This
has serious carbon balance and energetic consequences
(Table 5.5 of Raven, 1984).
5644 | Raven
Turning at last to measured half-lives of nucleus-encoded
chloroplast proteins, Table 3c gives a range from 116–301 h
for the most directly measured values, with the possibility of
some even longer half-lives. However, there is also the possibility of nuclear-encoded proteins with shorter half-lives that
would restrict the functional life of a kleptoplastid. While the
t1/2 data in Table 3c do not readily explain the longest-lived
kleptoplastids, it should be remembered that some mammalian erythrocytes function for months in an enucleate state.
Gillooly et al. (2012) showed that, when the two determinants of erythrocyte life-span, i.e. temperature at which the
erythrocytes function, and body mass of the animal in which
they occur, are allowed for, there is no difference in life-span
between enucleate mammalian erythrocytes and the erythrocytes of fish, amphibians, reptiles, and birds that are nucleate throughout their life, have functional mitochondria (Stier
et al., 2013), and retain transcriptional activity (Moreira
et al., 2011). While the habitat of a mammalian erythrocyte
is significantly different from that of plastids, the erythrocyte
data do show how long (over 104 h: Gillooly et al., 2012) enucleate cells can function despite the absence of protein synthesis. Raven (1991) pointed out that mature enucleate sieve
tube elements would lack the capacity to replace damaged
proteins and also suggested that low O2 concentrations in the
sieve tube elements and the frequent occurrence of a lignified
sheath round the vascular tissue would restrict damage from
oxidative and UVB damage, respectively. However, the need
for such protection is diminished by the demonstration of
symplasmic protein movement from the nucleate companion
cells to the enucleate sieve tube elements (Fisher et al., 1992;
Oparka and Turgeon, 1999).
Is there a protected germline organelle genome in
plants?
The copy number of the mitochondrial and, in particular,
the chloroplast genome has been measured with DNA dye
and molecular genetic methods (Tables 1, 2). These techniques generally showed several or many copies of the plastid
genome and (in algae) the mitochondrial genome, although
no genomes could be detected in some plastids of an alga,
and some mitochondria and chloroplasts of flowering plants
(Tables 1, 2). The consequences of the absence of DNA from
some organelles in vegetative growth are considered below.
Reproduction and the means by which at least one organelle
master genome, or its equivalent as subgenomic fragments,
is inherited, is explored here (Bendich, 1987, 1993, 1996;
Takanashi et al., 2010; Udy et al., 2012).
Where there are multiple copies of the organelle genome in
each organelle and/or many organelles per cell, then the probability is that each progeny cell at cell division will have at least
one organelle genome (Bendich, 1987; Udy et al., 2012). This
seems to be the case for chloroplasts (Tables 1, 2), although in
some algae and the embryophytic hornworts and some cells
of Selaginella there is only one chloroplast per cell and so they
must rely on the first alternative, i.e. the presence of multiple
genome copies in each organelle, in cell division. One or other
(or both) alternative means of ensuring organelle genome
propagation also apply to algal mitochondria (Tables 1, 2).
However, for flowering plant mitochondrial inheritance studies of egg cells of Arabidopsis, Wang et al. (2010) reported
that 67% of egg cell mitochondria contained no DNA, and
the 33% with DNA had an average of 109 kb DNA rather
than the 366.9 kb for a mitochondrial master genome, i.e. the
master genome is shared among mitochondria. This is also
the case for Antirrhinium majus and Nicotiana tabacum, but
not for Pelargonium zonale (Kuroiwa et al., 1990; Wang et al.,
2010). The reported extent of the loss of functional organelle
DNA during vegetative development in flowering plants correlates with the potential for plant regeneration from somatic
cells (Shaver et al., 2006).
Presumably the integrity of the master genomes (or their
equivalent) could be because they have either been subject to
fewer DNA changes, including that caused by damage, than
occurs in leaf cells, or have had relatively complete repair so
that their genomes are again in the master genome format;
investigation is needed of the structural variation and recovery history of the organelle genomes between reproductive
events. Embryophytes do not appear to have mechanisms
similar to those in metazoans maintaining relatively structurally unchanged germ line mitochondria (Ivanov, 2007;
de Paula et al., 2013). The ‘quiescent centre’ of flowering
plant roots (Clowes, 1958; Dubrovsky and Barlow, 2015)
has aspects of ‘stemness’, for example, a smaller electrical
potential difference at the inner membrane of the mitochondria and low activities of tricarboxylic acid cycle enzymes
(Jiang et al., 2006; de Paula et al., 2013), and retains organelle DNA turnover (Fujie et al., 1993). However, the cells
in the root apex do not contribute to the production of the
organs of sexual reproduction. For plant shoot apices, the
selection of the least damaged mitochondria for incorporation into reproductive cells does not fit with the suggestion
of Seguí-Simmaro and Staehlin (2009) of homogenization
of mitochondrial DNA involving mitochondrial fusion and
reticulation.
Unicellular organisms have even less possibility of maintaining a germ line of organelles, especially if, as in the
smallest known eukaryote, the picoplanktonic marine prasinophycean green alga Ostreococcus, there is only one chloroplast and one mitochondrion per cell (Blanc-Mathieu
et al., 2013). This means that there are no organelles that are
inactive in redox reaction and thus not as prone to oxidative
damage as those that are active in redox reactions (de Paula
et al., 2013). Although there seem to be no data on organelle
genome copy number for Ostreococcus, other very small
algal cells (e.g. the cyanidiophycean red alga Cyanidioschyzon
merolae) have multiple copies of the genome in each organelle
(Table 1; Hirakawa and Ishida, 2014). Is it possible that one
(or more) of these copies represent a germ line while the rest
do not?
Also unicellular (or, better, acellular) is the large-celled
green macroalga Acetabularia for which genome occurrence in
chloroplasts has been investigated by dye staining (Woodcock
and Bogorad, 1970; Coleman, 1979; Lüttke and Bonotto,
1981; Lüttke, 1988). This work on Acetabularia shows that
young vegetative cells have genomes in all their chloroplasts;
Organelle genomes for organelle function and evolution | 5645
as the alga continues vegetative growth an increasing fraction of the chloroplasts lack genomes, but that all chloroplasts in reproductive cells have genomes. There seem to be
no data on the extent of oxidative or photochemical damage
to DNA during vegetative growth, and during reproduction,
in Acetabularia.
Turning to variations in organelle function, there are
significant genotypic differences in mitochondrial function
among genotypes of Ostreococcus, as indicated by rhodamine as a measure of the proton motive force (proton
electrochemical potential difference) across the inner mitochondrial membrane (Schaum and Collins, 2014), so any
‘germ line’ mechanism to maintain mitochondrial function
as indicated by rhodamine does not work uniformly among
genotypes. Some kind of germ line-like mechanism for vegetative cell division in diatoms may be involved in the results
of Laney et al. (2012). Diatoms have overlapping frustules
comprising the bipartite cell wall with larger (epitheca) and
smaller (hypotheca) components. The new frustule components at vegetative cell division occur within the parental
frustule components. The frustule forming inside the original epitheca forms a hypotheca that is the same size as the
parental hypotheca, to produce a cell the same size as the
parental cell. The parental hypotheca becomes the epitheca
of the new cell, with a smaller new hypotheca formed inside
it. Laney et al. (2012) showed that the ‘younger’ (smaller)
cell had a greater specific growth rate than the ‘older’
(large) cell, to a greater extent than expected from the general specific growth rate–cell volume relationship of algae
(see Fig. 3 of Finkel et al., 2010). Since the nuclei of the
young and old cells should be essentially identical, could
it be that the younger cells have inherited organelles that
are less damaged at the nucleic acid and/or protein level?
This possibility involves the sorting of more and less damaged organelles at cell division such as is known to occur in
maintaining ‘stemness’ in mammalian stem cells (Katajisto
et al., 2015).
Structural variation is not correlated with the rate of
evolution of organelle genomes
An inevitable outcome of DNA damage and some other
forms of structural variation (Kumar et al., 2014) is chemical
alteration to nucleotides; without repair this would, at best,
give rise to inherited nucleotide change in the genome and,
at worst, the absence of replication or translation of the gene
involved. In view of the technical and interpretation problems
with the estimation of damage and other causes of variation
to organelle DNA discussed by Kumar et al. (2014), it is difficult to obtain comparative data on the variation, either gross
(without allowance for repair) or net (allowing for repair),
among oxygenic photosynthetic organisms. There are, however, data on the rate of genetic change in one or more of the
mitochondrial, chloroplastm and nuclear genomes.
Early comparisons for flowering plants (Wolfe et al., 1987)
showed that silent (synonymous) substitutions in mitochondrial DNA are less than one-third those of chloroplast DNA
which, in turn, is half of that in nuclear DNA. The mean
silent substitution rate in plant mitochondrial DNA is one
or two orders of magnitude less than that of metazoan
mitochondrial DNA (Brown et al., 1979), while the rates for
plant and metazoan nuclear DNA are similar (Wolfe et al.,
1987). The rates of non-synonymous substitutions in plant
mitochondria, chloroplasts, and nuclei are about an order
of magnitude less than those of the corresponding synonymous substitutions (Wolfe et al., 1987: Table 4). Later work
showed very high and variable mitochondrial substitution
rates in Plantago spp. (Cho et al., 2004) and some other
flowering plants (summarized by Smith et al., 2014a, b), and
extremely low substitution rates in the mitochondrial DNA
of Liriodendron and possibly even lower rates in the closely
related Magnolia (Richardson et al., 2013). The synonymous
substitution rate in the mitochondrial genome of flowering plants varies about 5000-fold (Richardson et al., 2013:
Table 4). Of the plants examined by Richardson et al. (2013)
Liriodendron and Magnolia had the lowest synonymous substitution rates in the chloroplast genome (though not as low
as in mitochondria of these magnolids), with a 50-fold range
in rates among flowering plants (Table 4).
Despite the great variability in flowering plant mitochondrial genome synonymous substitution rates, with a much
smaller range for plant chloroplast genomes, the general case
for seed plants is for mitochondrial substitution rates to be
less than the chloroplast rates, which are, in turn, less than the
nuclear rates (Smith et al., 2014a, b), as concluded by Wolfe
et al. (1987). Subsequent work on metazoans shows that the
conclusion of Wolfe et al. (1987), which was based largely
on vertebrates, as to the rate of synonymous substitution in
mitochondrial genomes relative to the nuclear genome rate
applies not just to the Deuterozoa (including vertebrates),
but also the Ecdysozoa (e.g. arthropods), Lophotrochozoa
(e.g. annelids and molluscs), platyhelminthes, and the medusozoan Cnidaria (Hellberg, 2006). Lower mitochondrial substitution rates are found in anthozoan Cnidaria and Porifera,
resembling the rates in flowering plants and fungi (Hellberg,
2006). I hesitate to relate the similarity of substitution rates
of Cnidaria, Porifera, flowering plants, and fungi to a shared
sedentary habit.
The situation with respect to the ranking of substitution
rates among the three genomes is different in algae (Smith
et al., 2014a, b: Table 4). Smith et al. (2014a, b) examined
the synonymous substitution rates in the three genomes of
Cyanophora paradoxa, a member of the basal (Glaucophyta)
clade of the Archaeplastida (=Plantae sensu lato), and showed
that the mitochondrial substitution rate was higher than both
that in the chloroplasts and the similar rate in the nucleus.
Smith et al. (2014a, b) review literature showing a similar pattern in another clade (Rhodophyta) of the Archaeplastida
(Table 4). For the Haptophyta, an algal clade with secondarily endosymbiotic chloroplasts derived from the Rhodophyta,
the mitochondrial rate is much higher than the chloroplast
rate, although here the nuclear rate is rather higher than that
of the chloroplast (Table 4; Smith et al., 2014a, b). For the
other clade of the Archaeoplastida, the Chlorophyta (including an algal member of the Streptophyta), the rates of substitution for the three genomes are closely similar, i.e. still
5646 | Raven
Table 4. Phylogenetic comparison of the rates of synonymous substitution in mitochondrial and chloroplast genomes relative to the rate
in the corresponding nuclear genome
Organism
Archaeoplastida
Glaucophyta
Cyanophora
Archaeplastida
Rhodophyta
Porphyra
Archaeplastida
Chlorophyta s.l.
Chlamydomonasa
Mesostigmab
Archaeplastida
Seed plants
Gymnosperms
Archaeplastida
Seed plants
Angiosperms
Ochrophyta
Raphidophyceae
Olisthodiscus
Haptophyta
Phaeocystis
Cryptophyta
Guillardia
Chlorarachniophyta
Bigelowia
Mitochondrial genome substitution rate
relative to nucleus=1
Chloroplast genome substitution rate
relative to nucleus=1
References
4.4
0.83
Smith et al. (2014a)
2.8
0.29
Smith et al. (2014a, b)
0.83
0.60
0.83
0.40
Smith et al. (2014a, b)
Hirakawa & Ishida (2014)
0.25
0.5
Smith et al. (2014a, b)
0.06c
0.19d
Smith et al. (2014a,b)
Ersland et al. (1981)
2.8
0.29
Smith et al. (2014a, b)
Hirakawa and Ishida (2014)
Hirakawa and Ishida (2014)
a
Chlorophyta sensu stricto.
Streptophyta sensu stricto.
c
5000 fold range (Richardson et al., 2013)
d
50 fold range (Richardson et al., 2013)
b
different from the seed plants (embryophyte members of the
Streptophyta) with the mitochondrial rate generally less than
that of the chloroplast, and even less than that for the nucleus
(summarized by Smith et al., 2014a, b: Table 4).
Although it is unfortunate that there are no comparable
rates of DNA damage for the organelle genomes for which
nucleotide substitution rates are available, it seems unlikely
that the substitution rates reflect differences in the rate of
DNA damage.
Why are the rates of DNA turnover in organelles much
higher than in their bacterial ancestors despite similar
rates of production of oxidants?
Despite the remaining technical problems (Kumar et al.,
2014), there is an obvious contrast between the large potential extent of oxidative DNA damage in organelles and the
bacterial ancestors of the organelles that have similar rates
(perhaps slightly lower for bacteria) of redox reaction per unit
volume to that of the organelles, and the much lower extent
of DNA turnover in the bacteria. There are very limited data
on the DNA turnover rates of organelles: organelle genomes
of Euglena gracilis turn over, 1.5–1.6 times per generation,
with minimal turnover of the nuclear genome (Manning and
Richards, 1972; Richards and Ryan, 1974). The minimal
turnover of the Euglena nuclear genome is similar to that
of the Proteobacteria (from which the mitochondrion was
derived) and of the Cyanobacteria (from which the chloroplast was derived) (Bridges, 1997; Foster, 1999; Shirkey et al.,
2003; Rajeev et al., 2013). The extant relatives of the organelle ancestors presumably have similar volume-based rates
of oxidant production to those of the respective organelles:
the rates may be lower in the free-living Cyanobacteria and
Proteobacteria since they have to fulfil more non-bioenergetic
cellular functions than is the case for the organelles.
It may be more relevant to ask why organelle DNA is so
prone to structural change, with substantial DNA turnover
presumably based on the synthesis of unchanged DNA to
replace changed DNA, rather than why their ancestors with
similar exposure to conditions that damage DNA have much
less DNA turnover. Even in extreme environmental conditions there is little oxidant damage to cyanobacterial DNA
(Shirkey et al., 2003; Rajeev et al., 2013). Bendich (2007)
point out that bacterial and archaeal genomes, like organelle
genomes, do not generally consist entirely, or even largely,
of ‘completed’ genomes, but contains a range of genomes
sizes reflecting (among other things) different extents of
replication of the complete genome. Furthermore, circular
Organelle genomes for organelle function and evolution | 5647
chromosomes are by no means universal among archaeal,
bacterial, and organelle genomes (Bendich and Smith, 1990;
Bendich, 2004, 2007; Shaver et al., 2008). The similarities in
the occurrence of incomplete genomes in Archaea, Bacteria,
and organelles clearly do not explain the differences in the
rates of DNA turnover between organelle genomes and the
genomes of Bacteria and Archaea.
Deinococcus radiodurans is a bacterium with extreme resistance to treatments that damage DNA and proteins, i.e. ionizing (including gamma) radiation, desiccation, oxidation,
and agents such as mitomycin C (Makarova et al., 2001;
Daly et al., 2010; Slade and Radman, 2011). Deinococcus is
closely related to Thermus, but otherwise is not closely related
to other known Bacteria (Makarova et al., 2001). While it is
not a close relative to the Proteobacteria from which mitochondria arose, Deinococcus is, like most interpretations of
the functioning of the ancestral Proteobacteria giving rise
to mitochondria, an aerobic chemo-organotroph. Mutant
studies show that the characteristics of Deinococcus that permit resistance to extreme environments are not required for
normal growth under non-extreme conditions (Makarova
et al., 2001; Daly et al., 2010; Slade and Radman, 2011).
Much of the emphasis on resistance to extreme conditions
by Deinococcus has emphasized damage to DNA (Makarova
et al., 2001), with such considerations as the minimum number (not more than five) of copies of the genome in each cell
that are needed for resistance of extreme conditions. However,
Daly et al. (2010) point out that the number of DNA doublestrand breaks caused by ionizing radiation in Deinococcus
are similar to the number that occur in (for example) meiosis in eukaryotes without the need for special methods of
DNA repair. Daly et al. (2010) emphasize the protection of
protein from oxidative damage in Deinococcus, and have partially characterized a low molecular mass extract containing
manganese, phosphate, and amino acids that acts to protect
proteins from oxidative damage, for example, by removing
hydroxyl radicals. However, this seems to have no direct relation to mechanisms protecting proteins from oxidative damage in mitochondria and chloroplasts.
Conclusions
Organelle genomes are subject to variation, including that
resulting from damage from reactive oxygen species, than
are eukaryotic nuclear genomes or bacterial genomes under
the same conditions. There have been, and still are, technical
problems in characterizing this damage and other variation
and its repair. Recent advances (Kumar et al., 2014) in the
methodology for characterizing the variation of genomes of
chloroplasts and mitochondria of Zea mays should, when
applied more widely, help us to understand how variation
in, including damage to, organelle genomes relates to maintenance of genome function during the life of individuals
and of organelle genetic integrity in individuals and in succeeding generations. Understanding of the extent of variation in organelle DNA sequence and structure, and its repair,
among photosynthetic organisms might also help to explain
the variations in the rate of inherited nucleotide substitution
among organelle genomes, and its relation to inherited nucleotide substitution rate in nuclear genomes. Further studies of
organelle DNA damage and repair might also help to understand why the extent of DNA turnover in the organelles is so
much greater than that in their bacterial (cyanobacteria for
chloroplasts, proteobacteria for mitochondria) relatives with
similar rates of production of DNA-damaging reactive oxygen species. Finally, we would hope for a better understanding of the relation of organelle DNA damage to organelle
protein damage and its repair; present data suggest that, after
the loss of functional organelle DNA encoding a given protein, neither the lifetime of that protein nor of the RNA species needed to replace it using mRNA produced before DNA
loss, can account for organelle function requiring that protein
for much longer than a week.
Acknowledgements
Discussions with John Allen, John Beardall, Jason Bragg, Sinéad Collins,
Paul Falkowski, Zoe Finkel, Kevin Flynn, Mario Giordano, Stephen
Hubbard, Andy Knoll, Hans Lambers, Tony Larkum, Antonietta Quigg,
Patricia Sanchez-Baracaldo, and Elisa Schaum have been very helpful.
Comments by two anonymous reviewers were very helpful in improving the
manuscript.
References
Abdellah F, Salamina F, Lesiter D. 2000. A prediction of the size and
evolutionary origin of the proteome of Arabidopsis. Trends in Plant Science
5, 141–142.
Allen JF. 1993a. Redox control of gene expression and function of
chloroplast genomes: an hypothesis. Photosynthesis Research 36,
95–102.
Allen JF. 1993b. Control of gene expression by redox potential and
the requirement for chloroplast and mitochondrial genomes. Journal of
Theoretical Biology 165, 609–631.
Allen JF. 2003. The function of genomes in bioenergetics organelles.
Philosophical Transactions of the Royal Society B 358, 19–38.
Allen, JF, Puthiyaveetil S, Strőm J, Allen CF. 2005. Energy transduction
anchors genes in organelles. Bioessays 27, 426–435.
Allen JF, Raven JA. 1996. Free-radical-induced mutation vs. redox
regulation: costs and benefits of genes in organelles. Journal of Molecular
Evolution 42, 482–492.
Barkan A, Goldschmidt-Clermont M. 2000. Participation of nuclear
genes in chloroplast gene expression. Biochemie 82, 559–572.
Bendich AJ. 1987. Why do chloroplasts and mitochondria contain so
many copies of their genome? Bioessays 6, 279–282.
Bendich AJ. 1993. Reaching for the ring: the study of mitochondrial
genome structure. Current Genetics 24, 279–280.
Bendich AJ. 1996, Structural analysis of mitochondrial DNA from fungi
and plants using moving pictures and pulsed-field gel electrophoresis.
Journal of Molecular Biology 255, 564–588.
Bendich AJ. 2004. Circular chloroplast chromosomes: the grand illusion.
The Plant Cell 16, 1661–1666.
Bendich AJ. 2007. The size and form of chromosomes are constant in
the nucleus, but highly variable in bacteria, mitochondria, and chloroplasts.
Bioessays 29, 474–483.
Bendich AJ, Gauriloff LP. 1984. Morphometric analysis of cucurbit
mitochondria: the relationship between chondriome volume and DNA
content. Protoplasma 119, 1–7.
Bendich AJ, Smith SB. 1990. Moving pictures and pulse-field gel
electrophoresis show linear DNA molecules from chloroplasts and
mitochondria. Current Genetics 17, 421–425.
5648 | Raven
Blanc-Mathieu R, Sanchez-Ferandin S, Eyre-Walker A, Piganeau
G. 2013. Organellar inheritance in the green lineage: insights from
Ostreococcus tauri. Genome Biology and Evolution 5, 1503–1511.
Bridges BA. 1997. DNA turnover and mutation in resting cells. Bioessays
19, 347–352.
Brown WM, George Jr M, Wilson AC. 1979. Rapid evolution of animal
mitochondrial DNA. Proceedings of the National Academy of Sciences,
USA 76, 1967–1971.
Cavalier-Smith T. 1970. Electron microscopic evidence for chloroplast
fusion in zygotes of Chlamydomonas reinhardii. Nature 228, 333–335.
Cho Y, Mower JP, Qiu Y-L, Palmer JD. 2004. Mitochondrial substitution
rates are extraordinarily elevated and variable in a genus of flowering
plants. Proceedings of the National Academy of Sciences, USA 101,
17741–17746.
Clowes FAL. 1958. Development of quiescent centres in roots meristems.
New Phytologist 57, 85–88.
Coleman AW. 1979. Use of the fluorochrome 4’6-diamido-2-phenylindole
in genetic and developmental studies of chloroplast DNA. Journal of Cell
Biology 82, 299–305.
Daly MJ, Gaidamakova EK, Matrosova VY, Kiang JG, Fukumoto R,
Leed DY, Wehr NB, Viteri GA, Berlett BS, Levine RL. 2010. Smallmolecule antioxidant protein shields in Deinococcus radiodurans. PLoS
ONE 5, e12570.
Danon A. 1997. Translational regulation in the chloroplast. Plant
Physiology 115, 1293–1298.
De Jong L, Elzinga SDJ, McCammon MT, Grivell LA, van der Spek
H. 2000. Increased synthesis and decreaed stability of mitochondrial
translatmion products in yeast as a result of loss of mitochondrial (NAD+)dependent isocitrate dehydrogenase. FEBS Letters 483, 62–66.
De Paula WBM, Lucas CM, Agip A-NA, Vizcay-Barrena G, Allen JF.
2013. Energy, ageing, fidelity, and sex: oocyte mitochondrial DNA as a
potential genetic template. Philosophical Transactions of the Royal Society
of London B 368, 2012063.
de Vries J, Habicht J, Woehle C, Huang C, Christa G, Wägele H,
Nickelsen J, Martin WF, Gould SB. 2013. Is ftsH the key to plastid
longevity in the sacoglossan slugs? Genome Biology and Evolution 5,
2540–2548.
Dubrovsky JG, Barlow PW. 2015. The origins of the quiescent centre
concept. New Phytologist 206, 493–496.
Eberhard S, Finazzi G, Wollman F-A. 2002. Searching limiting steps
in the expression of chloroplast-encoded proteins: relation between gene
copy number, transcription, transcript abundance, and translation rate
in the chloroplast of Chlamydomonas reinhardtii. The Plant Journal 31,
149–160.
Ersland DR, Aldrich J, Cattolico RA. 1981. Kinetic complexity,
homogeneity, and copy number of chloroplast DNAs for the marine alga
Olisthodiscus luteus. Plant Physiology 68, 1468–1473.
Falkowski PG, Raven JA. 2007. Aquatic photosynthesis ,2nd edn.
Princeton; Princeton University Press.
Finkel ZV, Beardall J, Flynn KJ, Quigg A, Rees TAV, Raven JA. 2010.
Phytoplankton in a changing world: cell size and elemental stoichiometry.
Journal of Plankton Research 32, 119–137.
Fisher DB, Wu Y, Ku MSB. 1992. Turnover of soluble proteins in the
wheat sieve tube. Plant Physiology 100, 1433–1441.
Foster PL. 1999. Mechanisms of stationary phase mutation: a decade of
adaptive mutation. Annual Review of Genetics 33, 57–88.
Fromm H, Devic M, Fluhr R, Edelman M. 1985. Control of psbA gene
expression: in mature Spirodela chloroplast light regulation of 32 kDa protein
synthesis is independent of transcript level. EMBO Journal 4, 291–295.
Fujie M, Kuroiwa H, Suzuki T, Kawano S, Kuroiwa T. 1993. Organelle
DNA synthesis in the quiescent centre of Arabidopsis thaliana (Col.).
Journal of Experimental Botany 44, 689–693.
Germain A, Kim SH, Gutierrez R, Stern DB. 2012. Ribonuclease II
preserves chloroplast RNA homeostasis by increasing mRNA decay rates,
and cooperates with polynucleotide phosphorylase in 3’ end maturation.
The Plant Journal 72, 960–971.
Giegé P, Hoffmann M, Binder S, Brennicke A. 2000. RNA degradation
buffers assymetries of transcription in Arabidopsis mitochondria. EMBO
Reports 1, 164–170.
Gilham NW, Boynton J, Rochaix JD. 1997. Translational regulation
of gene expression in chloroplasts and mitochondria. Annual Review of
Genetics 28, 71–93.
Gillooly JF, Hayward A, Hou C, Burleigh JG. 2012. Explaining
differences in the lifespan and replicative capacity of cells: a general model
and comparative analysis of vertebrates. Proceedings of the Royal Society
B 279, 3976–3980.
Gong XS, Chung S, Fernández-Velasco JG. 2001. Electron
transfer and stability of the cytochrome b6f complex in a small domain
deletion mutant of cytochrome f. Journal of Biological Chemistry 276,
24365–24371.
Halliwell B, Gutteridge JMC. 2007. Free radicals in biology and
medicine , 4th edn. Oxford: Oxford University Press.
Heazlewood JL, Tonti-Filippini JS, Gout AM, Day DA, Whelan J,
Millar AH. 2004. Experimental analysis of the Arabidopsis mitochondrial
proteome highlights signalling and regulatory components, provides
assessment of targeting prediction programs, and indicates plant-specific
mitochondrial proteins. The Plant Cell 16, 241–256.
Hellberg ME. 2006. No variation and low synonymous substitution rates in
coral mtDNA despite high nuclear variation. BMC Molecular Biology 6, 24.
Hirakawa Y, Ishida K-Y. 2014. Polyploidy of endosymbiotically derived
genomes in complex algae. Genome Biology and Evolution 6, 974–980.
Hojka M, Theil W, Tóth SZ, Lein W, Bock R, Schöttler MA. 2014.
Inducible repression of nuclear-encoded subunits of the cytochrome b6-f
complex in tobacco reveals an extraordinarily long lifetime of the complex.
Plant Physiology 165, 1632–1646.
Holec S, Langer H, Kühn K, Alioua M, Börner T, Gagliardi D. 2006.
Relaxed transcription in Arabidopsis mitochondria is counterbalanced
by RNA stability control mediated by polyadenylation and polynucleotide
phosphorylase. Molecular and Cellular Biology 26, 2869–2876.
Hosler JP, Wurtz EA, Harris EH, Gillham NW, Boynton JE. 1989.
Relationship between gene dosage and gene expression in the chloroplast
of Chlamydomonas reinhardtii. Plant Physiology 91, 648–655.
Inamine G, Nash B, Weissbach H, Brot N. 1985. Light regulation of the
synthesis of the large subunit of ribulose-1.5-bisphosphate carboxylase
in peas: evidence for translational control. Proceedings of the National
Academy of Sciences, USA 82, 5690–5694.
Ivanov VB. 2007. Stem cells in the root and the problem of stem cells in
plants. Russian Journal of Developmental Biology 38, 338–349.
Jiang K, Ballinger T, Li D, Zhang S, Feldman L. 2006. A role for
mitochondria in the establishment and maintenance of the maize root
quiescent center. Plant Physiology 140, 1118–1125.
Katajisto P, Döhla J, Chaffer CL, Pentinmikko N, Marjanovic N,
Iqbal S, Koncu R, Chen W, Weinberg RA, Sabatini DM. 2015.
Asymmetric apportioning of aged mitochondria between daughter cells is
required for stemness. Science 348, 340–343.
Kim MY, Christopher DA, Mullet JA. 1993. Direct evidence for selective
modulation of pdbA, rpoA, rbcL, and 16S-RNA stability during barley
chloroplast debelopment. Plant Molecular Biology 22, 447–463.
Klaff P, Gruisson W. 1991. Changes in chloroplast mRNA stability during
leaf development. The Plant Cell 3, 517–529.
Kleffman T, Rusenberger D, von Zychlinski A, Christopher W,
Sjölander W, Bagicnky S. 2004. The Arabidiopsis thaliana chloroplast
proteome reveals pathway abundance and reveal protein functions.
Current Biology 14, 354–362.
Krech K, Ruf S, Masduki FF, Thiele W, Bednarczyk D, Albus CA,
Tiller N, Hasse C, Schöttler MA, Bock R. 2012. Thy plastid genomeencoded Ycf4 protein functioins as a non-essential assembly factor for
photosystem I in higher plants. Plant Physiology 159, 579–591.
Kumar RA, Oldenburg DJ, Bendich AJ. 2014.Changes in DNA
damage, molecular integrity, and copy number for plastid DNA and
mitochondrial DNA during maize development. Journal of Experimental
Botany 65, 6425–6439.
Kuroiwa T, Suzuki T, Ogawa K, Kawano S. 1981. The chloroplast
nucleus: distribution, number, size, and shape, and a model for the
multiplication of the chloroplast genome during chloroplast development.
Plant and Cell Physiology 22, 381–396.
Kuroiwa T, Kuroiwa H, Mita T, Fujie M. 1990. Fluorescence microscopy
study of the formation of giant mitochondrial nuclei in the young ovules of
Pelargonium zonale. Protoplasma 158, 191–194.
Organelle genomes for organelle function and evolution | 5649
Lamppa GK, Elliot LV, Bendich AJ. 1980. Changes in chloroplast
number during pea leaf development. Planta 148, 437–443.
Laney SR, Olsen RJ, Sosik HM. 2012. Diatoms favour their younger
daughters. Limnology and Oceanography 57, 1572–1578.
Li W, Ruf S, Bock R. 2006. Constancy of organellar genome copy
numbers during leaf development in higher plants. Molecular Genetics and
Genomics 275, 185–192.
Lűttke A. 1988. The lack of chloroplast DNA in Acetabularia mediterranea
(acetabulum) (Chlorophyceae): a reinvestigation. Journal of Phycology 24,
173–180.
Lűttke A, Bonotto S. 1981. Chloroplast DNA of Acetabularia
mediterranea: cell cycle related changes in distribution. Planta 153,
536–542.
Makarova KS, Aravind L, Wolf YI, Tatusov RL, Minton KW, Koonin
EV, Daly MJ. 2001. Genome of the extremely radiation-resistant
bacterium Deinococcus radiodurans viewed from the perspective of
comparative genomics. Microbial Molecular Biology Reviews 65, 44–79.
Manning JE, Richards OC. 1972. Synthesis and turnover of Euglena
gracilis nuclear and chloroplast deoxynucleic acid. Biochemistry 11,
2036–2043.
Martin S, Mungapati VS, Salvo-chirnside E, Kerr LE, Le Bihan T.
2012. Proteome turnover in the green alga Ostreococcus tauri by timecourse, 15N metabolic labelling mass spectrometry. Journal of Proteome
Research 11, 476–486.
Martin W, Kowallik KV. 1999. Annotated English translation
of Mereschowsky’s, 1905 paper ‘Uber Natur und Ursprung der
Chromatophoren in Pflanzenreiche’. European Journal of Phycology 34,
287–295.
Martin W, Stoebe B, Gorymekin V, Henesmann S, Hasegawa M,
Kowallik KV. 1998. Gene transfer to the nucleus and the evolution of
chloroplasts. Nature 393, 162–165.
Mastrobuoni G, Igang S, Pietzke M, Wenzel M, Assmus HE, Schulze
WX, Kempa S. 2012. Proteome dynamics and early salt stress response
of the photosynthetic organism Chlamydomonas reinhardtii. BMC
Genomics 13, 215.
Mereschowsky C. 1905. Uber Natur und Ursprung der Chromatophoren
in Pflanzenreiche. Biologische Centralblatte 25, 583–694. [English
translation by Martin and Kowallik, 1999.]
Moreira D, Roher N, Ribas L, et al. 2011. RNA-Seq reveals integrated
immune response in nucleated erythrocytes. PLoS ONE 6, e26998.
Nelson CJ, Alexova R, Jacoby RP, Millar AH. 2014. Proteins with high
turnover rate in barley leaves estimated by proteome analysis combined
with in planta isotope labelling. Plant Physiology 166, 91–108.
Nelson CJ, Li L, Jacoby RP, Miller AH. 2013. Degradation rates of
mitochondrial proteins in Arabidopsis thaliana cells. Journal of Proteome
Research 12, 3449–3459.
Oldenburg DJ, Bendich AJ. 2004. Changes in the structure of
DNA molecules and the amount of DNA per plastid during chloroplast
development in maize. Journal of Molecular Biology 344, 1311–1330.
Oldenburg DJ, Kumar RA, Bendich AJ. 2013. The amount and integrity
of mtDNA in maize with development. Planta 237, 603–617.
O’Malley MA. 2015. Endosymbiosis and its implications for evolutionary
theory. Proceedings of the National Academy of Sciences, USA (in press)
doi:, 10.1073/pnas.1421389112.
Oparka K, Turgeon R. 1999. Sieve elements and companion cells: traffic
control centers of the phloem. The Plant Cell 11, 739–750.
Piques M, Schulze WX, Höhne M, Usadel B, Gibon Y, Rohwer J, Stitt
M. 2009. Ribosome and transcript copy numbers, polysome occupancy
and enzyme dynamics in Arabidopsis. Molecular Systems Biology 5, 314.
Preuten T, Cineu E, Fuchs J, Zoschke R, Liere K, Börner T. 2010.
Fewer genes than organelles: extremely low and variable gene copy numbers
in mitochondria of somatic plant cells. The Plant Journal 64, 948–959.
Race HL, Herrmann RE, Martin W. 1999. Why have organelles retained
genomes? Trends in Genetics 16, 164–370.
Rajeev L, Nunes da Rocha U, Klitgord N, et al. 2013. Dynamic
cyanobacterial in a desert biological soil crust. The ISME Journal 7,
2178–2191.
Raven JA. 1984. Energetics and transport in aquatic plants . New York:
AR Liss.
Raven JA. 1991. Long-term functioning of enucleate sieve elements:
possible mechanisms of damage avoidance and damage repair. Plant, Cell
and Environment 14, 139–146.
Raven JA, Beardall J, Giordano M. 2014. Energy costs of carbon
dioxide concentrating mechanisms in aquatic organisms. Photosynthesis
Research 121, 111–124.
Raven JA, Beardall J, Flynn KJ, Maberly SC. 2009. Phagotrophy in the
origins of photosynthesis in eukaryotes and as a complementary mode of
nutrition in phototrophs: relation to Darwin’s insectivorous plants. Journal
of Experimental Botany 60, 3975–3987.
Raven JA, Giordano M, Beardall J, Maberly SC. 2012. Algal evolution
in relation to atmospheric CO2: carboxylases, carbon-concentrating
mechanisms and carbon oxidation cycles. Philosophical Transactions of
the Royal Society B 367, 493–507.
Richards OC, Ryan RS. 1974. Synthesis and turnover of Euglena gracilis
mitochondrial DNA. Journal of Molecular Biology 82, 57–75.
Richardson A, Rice DW, Alverson AJ, Young GJ, Palmer JD. 2013
The ‘fossilized’ mitochondrial genome of Liriodendron tulipifera: ancestral
gene content and order, ancestral editing sites, and extraordinarily low
mutational rates. BMC Biology 11, 29.
Rowan BA, Bendich AJ. 2009. The loss of DNA from chloroplasts
as leaves mature; fact or artefact? Journal of Experimental Botany 60,
3005–3010.
Rowan BA, Oldenburg DJ, Bendich AJ. 2002. The demise of
chloroplast DNA in Arabidopsis. Current Genetics 46, 176–181.
Schaum CE, Collins S. 2014. Plasticity predicts evolution in a marine
alga. Proceedings of the Royal Society of London B 281, 201486.
Schimper AFG. 1883. Ueber die Entwiselung der Chlorophyllkōrper
und Farbkōrper. Flora oder Allgemeine Botanische Zeitung 41, 105–114,
121–131, 137–146, 153–162.
Schmitz FAJ. 1882. Der Chromatophoren der Algen. Vergleichende
Untersuchungen über Bau and Entwicklung der Chlorophyllkōrper und der
analogen Farbstoffkōrper der Algen. Bonn Verlag von Max Coghen & Sohn
(Dr. Cohen).
Schuster G, Timberg R, Ohad I. 1988. Turnover of thylakoid
photosystem II proteins during photoinhibition of Chlamydomonas
reinhardtii. European Journal of Biochemistry 177, 403–410.
Seguí-Simarro JM, Staehlin LA. 2009. Mitochondrial reticulation in
shoot apical meristem cells of Arabidopsis provides a mechanism of
homogenisation of mtDNA prior to gamete formation. Plant Signalling and
Behaviour 4, 168–171.
Shaver JM, Oldenburg DJ, Bendich AJ. 2006. Changes in chloroplast
DNA during development in tobacco, Medicago, pea, and maize. Planta
224, 72–82.
Shaver AM, Oldenberg, DJ, Bendich AJ. 2008. The structure of
chloroplast DNA molecules and the effect of light in the amount of
chloroplast DNA during development of Medicago trunculata. Plant
Physiology 146, 1064–1074.
Shirkey B, McMaster NJ, Smith SC, Wright DJ, Rodriguez H, Jaruga
P, Birincioglu M, Helm RF, Potts M. 2003. Genomic DNA of Nostoc
commune (Cyanobacteria) becomes covalently modified during longterm (decades) desiccation but is protected from oxidative damage and
degradation. Nucleic Acids Research 31, 2995–300.
Simpson E, Cooke RJ, Davies DD. 1981. Measurement of protein
degradation in leaves of Zea mays using [3H] acetic anhydride and tritiated
water. Plant Physiology 67, 1214–1219.
Slade D, Radman M. 2011. Oxidative stress resistance in
Deinococcus radiodurans. Microbial and Molecular Biology Reviews 75,
133–191.
Smith DR, Jackson CJ, Reyes-Prieto A. 2014a. Nucleotide substitution
rate of the glaucophyte Cyanophora suggest an ancestrally lower mutation
rate in plastid vs mitochondrial DNA for the Archaeplastida. Molecular
Phylogenetics and Evolution 79, 80–184.
Smith DR, Arrigo KR, Aldekamp A-C, Allen AE. 2014b. Massive
differences in synonymous substitution among mitochondrial, plastid,
and nuclear genes of Phaeocystis algae. Molecular Phylogenetics and
Evolution 71, 36–40.
Sodmergen, Kawano S, Tabo S, Kuroiwa T. 1991. Degradation of
chloroplast DNA in the second leaves of rice (Oryza sativa) before leaf
yellowing. Protoplasma 160, 89–98.
5650 | Raven
Stier A, Bize P, Schull Q, Zall J, Singh F, Geny R, Gros F, Royer C,
Messeminn S, Criscuolo F. 2013. Avian erythrocytes have functional
mitochondria, opening novel perspectives for birds as animal models in the
study of aging. Frontiers in Zoology 10, 33.
Sundberg C, McCaffery S, Anderson JM. 1993. Turnover of
Photosystem II D1 protein in higher plants under photoinhibitory and
nonphotoinhibitory irradiance. Journal of Biological Chemistry 268,
25476–25482.
Takanashi H. Ohnishi T, Mogi M, Okamoto T, Arimura S-I, Tsutsumi
N. 2010. Studies of mitochondrial morphology and DNA content in the rice
egg cell. Current Genetics 56, 33–41.
Udy DB. Belcher S, Williams-Carrier R, Gualberto JM, Barkan A.
2012. Effects of reduced chloroplast gene copy number on chloroplast
gene expression in maize. Plant Physiology 160, 1420–1431.
Wägele H, Deusch O, Händeler K, et al. 2011. Transcriptomic evidence
that longevity of acquired plastids in the photosynthetic slugs Elysia timida
and Plakobranchus ocellatus does not entail lateral gene transfer of algal
nuclear genes. Molecular Biology and Evolution 28, 699–706.
Wallin JE. 1923. The mitochondria problem. American Naturalist 57, 255–261.
Wang D-Y, Zhang Q, Liu Y, Lin Z-F, Zhang S-X, Sun M-X,
Sodmergen. 2010. The levels of male gametic mitochondrial DNA are
highly regulated in angiosperms with regard to mitochondrial inheritance.
The Plant Cell 22, 2402–2416.
Wolfe KH, Li W-H, Sharp PM. 1987. Rates of nucleotide substititution
vary greatly between plant mitochondrial, chloroplast and nuclear DNA.
Proceedings of the National Academy of Sciences USA 84, 9054–9058.
Woodcock CF, Bogorad L. 1970. Evidence for variation in the quantity of
DNA among plastids of Acetabularia. Journal of Cell Biology 44, 361–375.
Zerges W, Girard-Bascou J, Rochaix JD. 1997. Translation of the
chloroplast psbC mRNA is controlled by interactions between its 5’ leader
and the nuclear loci TBC1 and TBC3 in Chlamydomonas reinhardii.
Molecular and Cell Biology 17, 3440–3448.
Zotschke R, Liere K, Börner T. 2007. From seedling to mature plant:
Arabidopsis plastidial genome copy number, RNA accumulation and
transcription are differentially regulated during leaf development. The Plant
Journal 50, 710–722.