Journal of Experimental Botany, Vol. 65, No. 10, pp. 2563–2572, 2014
doi:10.1093/jxb/ert469 Advance Access publication 3 February, 2014
Opinion Paper
Cell cycling with the SEB: a personal view
John Bryant*
School of Biosciences, CLES, University of Exeter, Exeter EX4 4PS, UK
* To whom correspondence should be addressed. E-mail: [email protected]
Received 14 October 2013; Revised 2 December 2013; Accepted 5 December 2013
Abstract
This review, written from a personal perspective, traces firstly the development of plant cell cycle research from the
1970s onwards, with some focus on the work of the author and of Dr Dennis Francis. Secondly there is a discussion
of the support for and discussion of plant cell cycle research in the SEB, especially through the activities of the Cell
Cycle Group within the Society’s Cell Biology Section. In the main part of the review, selected aspects of DNA replication that have of been of special interest to the author are discussed. These are DNA polymerases and associated
proteins, pre-replication events, regulation of enzymes and other proteins, nature and activation of DNA replication
origins, and DNA endoreduplication. For all these topics, there is mention of the author’s own work, followed by a brief
synthesis of current understanding and a look to possible future developments.
Key words: Cell cycle, DNA polymerase, DNA replication, regulation, replication origins, SEB.
Introduction
This short review has two main themes. The first is the role of
the SEB in supporting and presenting work on the cell division cycle, with some mention of my work and the work of
Dr Dennis Francis. The second is to look at some specific topics within the general area of cell cycle research, giving a brief
historical overview and a look to the future. I have focused,
in particular, on topics concerned with DNA replication,
the research area that led me into the cell cycle. Because of
the personal perspective, the writing style is, in places, more
informal than usual.
During the 1970s there was relatively little research activity
on the plant cell cycle. A few research groups, including my
own, were investigating plant DNA polymerases (Castroviejo
et al., 1975; Robinson and Bryant, 1975; Tymonko and
Dunham, 1977; Stevens and Bryant, 1978) based on the view
that these would have a major role in the regulation of DNA
replication. There was also some interest in the regulation of
the cell cycle in relation to development and this included the
patterns of cell cycle arrest during cellular differentiation and
the regulation of both DNA replication and mitosis during
developmental transitions (Harland et al., 1973; Robinson
and Bryant, 1975). It also included the role of principal
control points (Van’t Hof, 1973) in the observed changes in
cell cycle dynamics in different phases of plant life or under
different in vitro conditions. Further, Jack Van’t Hof, whom
we regard as one of the ‘founding fathers’ of modern plant
cell cycle research, had started to apply the technique of
fibre autoradiography to plant cells (Van’t Hof, 1975; Van’t
Hof and Bjerknes, 1977) and had used this to demonstrate
the bidirectional movement of replication forks away from
individual origins of replication. This confirmed that plant
chromosomes were, like animal chromosomes, organized as
multiple replicons.
In 1978, Dennis Francis was appointed to a lectureship in
Plant Cell Biology in Cardiff where I was already a member of
the Faculty. He brought with him expertise in measuring cell
cycle activity, including the assessment of the lengths of the cell
cycle phases in flowering and in the dynamics of meristem formation in roots. I was already leading one of the few research
groups worldwide looking at the biochemistry of plant DNA
replication and had started to take an interest in the nature of
replication origins. I was also interested in the integration of
DNA replication within plant development. Thus began a collaboration which, although not the major part of the research
© The Author 2014. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved.
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2564 | Bryant
activity of either one of us, nevertheless continued sporadically for over 20 years (Bryant et al., 1981; Francis et al., 1985;
Silcock et al., 1990; Dambrauskas et al., 2003).
Alongside our joint research, I continued, after my move
to Exeter in 1985, to investigate the complexes of proteins
involved in DNA replication (Bryant et al., 1992) This led
eventually to assaying the expression of genes encoding some
of those proteins. Much of this later work was carried out in
collaboration with Dr Steve Aves (Hart et al., 2002). I also
became interested in the control of DNA endoreduplication,
a cell cycle variant that occurs in a wide range of plant cells.
My interest in the nature of DNA replication origins has continued to the present day, although all the efforts of my group
to isolate bona fide origins came to nothing. In the meantime,
Dennis Francis, after a visit to Jack Van’t Hof’s laboratory
to learn autoradiography, continued his study of cell cycle
dynamics in a variety of situations including stress responses,
floral induction, and seed development (the latter also involving an interest in DNA endoreduplication). He was also
interested in the relationship between DNA C-value and cell
cycle dynamics (Kidd et al., 1987) and he studied variations,
both inter- and intra-specific, in replicon spacing and in fork
rates (Francis and Bennett, 1982; Ormrod and Francis, 1986;
Kidd et al., 1989). Both of these were the subject of discussion whenever we met. Towards the end of his active research
career, Dennis became very interested in the regulation of the
G2–M transition which led him to work with the plant homologues of cdc25 and wee1 (Orchard et al., 2005; Sorrell et al.,
2005; Francis, 2008, 2011). Some of this later work was carried out in collaboration with Dr Hilary Rogers (Spadafora
et al., 2012; Cook et al., 2013).
The SEB and the cell cycle
However, it is within the activities of the Society for
Experimental Biology that the collaboration between myself
and Dennis Francis proved most fruitful. In 1984, taking
advantage of the Society’s meeting in Cardiff, we ran an international symposium on The Cell Division Cycle in Plants. The
proceedings were published by Cambridge University Press
as No. 26 of the SEB Seminar Series (Bryant and Francis,
1985). The symposium was very enthusiastically received and
from it arose some scientific friendships and collaborations
which lasted for many years.
With regard to the SEB itself, the symposium led to a
renewal of interest in the cell division cycle and to the establishment of the Cell Cycle Group within the Cell Biology
Section. Over a period of more than 20 years, the Group
ran sessions on the cell cycle at the Annual Main meeting
every two to three years; these sessions often included speakers working on a variety of organisms, not just plants. In
addition, the Group put on sessions on topics such as DNA
repair. This eclectic approach was typified by the 2006 SEB
Symposium, The Eukaryotic Cell Cycle (Bryant and Francis,
2008) and by the Group’s co-sponsorship of two other symposia, Programmed Cell Death in Animals and Plants (Bryant
et al., 2000a) and The Nuclear Envelope (Evans et al., 2004).
However, the Group did, from time-to-time, run plant-only
sessions; thus the plant cell cycle sessions held within the 1992
Annual Main Meeting led to the publication of Molecular
and Cell Biology of the Plant Cell Cycle (Ormrod and Francis,
1993). A special issue of The Journal of Experimental Botany
(Vol. 54, No. 380) came from sessions on Plant Reproductive
Biology held in conjunction with the Plant Developmental
Biology Group at the Annual Main Meeting in Swansea in
2002. More recently, a special issue of Annals of Botany (Vol.
107, No. 7) resulted from the sessions held during the Annual
Main Meeting at Prague in 2010. The current special issue of
The Journal of Experimental Botany, based on sessions held
at the 2013 Annual Main Meeting in Valencia, follows in that
tradition.
Finally, I note that the Group has nominated, on behalf
of the Cell Biology Section, three plenary speakers at the
Annual Main Meeting, namely Ron Laskey (Bidder Lecturer,
York, 1998), Linda Hanley-Bowdoin (Woolhouse Lecturer,
Swansea, 2002), and Kim Nasmyth (Bidder Lecturer,
Glasgow, 2009).
Selected topics in DNA replication
Proteins involved in DNA replication
In the 1970s and 1980s, much of my work in this area, some of
it carried out in collaboration with Dr Valgene Dunham, then
at the State University of New York Fredonia and later at the
University of Western Kentucky (Dunham and Bryant, 1986)
focused on the DNA polymerases. How many different types
were there, did their activities change in relation to phases of
plant development and if so, could anything about their roles
be inferred? (reviewed by Bryant and Dunham, 1988). Leaving
aside the polymerase(s) of chloroplasts and mitochondria,
we reached the conclusion that activity of a ‘high-molecularweight’ DNA polymerase was correlated with DNA replication.
In all respects that we able to assay, this strongly resembled the
DNA polymerase-α of mammalian cells (Stevens and Bryant,
1978; Bryant and Dunham, 1988). There was also strong evidence for a low-molecular-weight DNA polymerase that was
tightly bound to chromatin (Tymonko and Dunham, 1977;
Stevens et al., 1978); in its properties this strongly resembled
DNA polymerase-β, a repair-associated polymerase in animal
cells. The distribution of its activity in plants certainly suggested that it was not involved with replication and thus it was
suggested that this was also a repair enzyme. It needs to be said
that, although the existence of plant DNA polymerase-α was
accepted by all of the small number of groups working with
plant polymerases, the existence of polymerase-β-like enzyme
was disputed (as discussed in Bryant and Dunham, 1988;
Bryant, 2010). The difference of opinion lasted until relatively
recently and was resolved when annotation of sequenced plant
genomes (MIPS, 2008) revealed the existence of a gene encoding DNA polymerase-β (Bryant, 2010), although some investigators still deny its existence.
From this point, this research went four ways. The first
was the growing realization that replication of nuclear DNA
Cell cycling with the SEB: a personal view | 2565
involved multiple DNA polymerases. Our current understanding (reviewed in Bryant, 2010) is that DNA polymerase-α with
its associated primase activity lays down and slightly extends
the primers. On the lagging strand, primer extension to form
Okazaki fragments is completed by DNA polymerase-δ
with its processivity factor, PCNA. (Note that PCNA has a
number of other roles in DNA replication; see, for example,
Raynaud et al., 2006.) On the leading strand, primers are
extended by DNA polymerase-ε. The latter enzyme is also
involved in DNA stress checkpoints and possibly in some
types of DNA repair. The list of polymerases has grown to
15 in mammals. Although there are still some differences of
opinion about the suite of polymerases possessed by plants,
my view (Bryant, 2010) is that plants possess 11 of the 15
mammalian polymerases, including DNA polymerase-β (see
above). One of the mammalian enzymes absent from plants
is DNA polymerase-γ (despite earlier claims—see review in
Bryant and Dunham, 1988) which replicates mitochondrial
DNA. Instead, plants possess two enzymes that appear to be
involved in organellar DNA synthesis (reviewed in Bryant
2010; see also Moriyama et al., 2011).
Relevant to our understanding of the cell cycle in plant
development are the wider-ranging effects of mutations in
DNA polymerase genes. Thus, Arabidopsis INCURVATA2
actually encodes the catalytic sub-unit of DNA polymerase-α
(Barrero et al., 2007). Although strong mutant alleles are
lethal, weaker alleles cause distinct developmental aberrations: early flowering, leaf incurvature, and homeotic transformations of floral organs. Similarly, the TILTED allele is
a weak mutation in one of two genes that code for the catalytic subunit of DNA polymerase-ε. The mutants show a 35%
increase in the length of the cell cycle and a change in cell patterning during embryo development (Jenik et al., 2005). Both
these examples show an interrelation between DNA replication and development, possibly acting via epigenetic patterning (Barrero et al., 2007; Sanchez et al., 2008). Another gene
that affects the function of DNA polymerase-ε is Abo4-1,
first detected by virtue of its mutant phenotype presenting as
an over-sensitivity to ABA (Dona et al., 2013). This may be
related to the role of the polymerase in the activation of the
stress checkpoint. It certainly shows again that the influence
of these polymerases goes far beyond their basic catalytic
activity, reaching out into aspects of development.
The second path along which this research developed was
a study of the other enzymes and proteins associated with the
polymerases. In our work on DNA polymerase-α, we were
routinely able to isolate the enzyme as part of a large multiprotein complex (Bryant et al., 1992), similar to the situation
with mammalian polymerase-α. ‘Dissection’ of the complex
mostly proceeded by biochemical assay of enzyme activities.
Thus it was established that primase was tightly linked to the
polymerase, suggesting that, as in other systems, polymeraseα-primase formed a sub-complex, probably of four subunits. Topoisomerase I was certainly present and possibly
topoisomerase II. Topoisomerase I is essential for removing
the supercoiling during movement of the replication forks
while topoisomerase II resolves the knots in DNA that occur
when two replication forks back into each other (Bryant and
Dunham, 1988). In collaboration with Dr Donato Chiatante,
both these enzymes, extracted from isolated whole nuclei,
were carefully characterized (Chiatante et al., 1991, 1993;
Chiatante and Bryant, 1994; Chiatante et al., 1997).
The polymerase-α complex also contains an unusual DNAbinding protein which, in a collaboration with Dr Jack Van’t
Hof, was shown to recognize ds–ss junctions in DNA (Burton
et al., 1997). This could certainly function as a primer recognition protein (Burton et al., 1997; Bryant et al., 2000b), similar to that described in mammalian cells. However, in view
of the close association between primase and polymerase-α,
it is difficult to understand the significance of such a role
(discussed in Bryant, 2008, 2010). Further, in a collaboration
with Dr Louise Anderson at the University of Illinois, it was
demonstrated that PRP is a ‘moonlighting’ population of the
cytoplasmic enzyme, PGK (Anderson et al., 2004); again this
is similar to the situation in mammalian cells. Finally, a DNA
helicase activity which was not an MCM protein was detected
by biochemical assay, while MCM3 was detected immunologically. The MCM complex is generally regarded as the replicative helicase, working in front of polymerase-primase to keep
the replication fork moving, so the role of a separate helicase
is as yet unknown.
With modern, fast protein sequencing techniques and with
a wider range of antibodies available, much more rapid progress could be made in analysing these protein complexes.
The main point here is that the synthesis phase of DNA replication requires a large number of proteins which associate
to form a loose complex, probably grouped as two sub-complexes. Further, the interaction of these two sub-complexes
with the template changes during the replication of each
tract of DNA. On the lagging strand, primase gives way to
polymerase-α (see above) and then the primase-polymerase-α
complex is displaced by polymerase-δ; on the leading strand
it is DNA polymerase-ε that displaces primase-polymerase-α.
What is now needed is to understand how these complexes
assemble together on the template with the proteins in the
right relationship to each other. Further, it would be helpful
to know how enzymes working downstream of the polymerases (in particular, the ligases needed for joining the nascent
DNA strands: Bray et al., 2008) interact with these polymerase complexes. The role of accessory proteins in polymerase
complexes remains unclear as do the factors involved in regulating the correct assembly of the complexes.
This leads us to consider the third strand of polymerase-related research: the pre-replicative events that happen before DNA polymerase-α-primase is loaded onto
the template. Much of our understanding of this came
initially from the study of cell cycle mutants in fission
yeast (Schizosaccharomyces pombe) and budding yeast
(Saccharomyces cerevisiae) but we are now building up a
clearer picture of what happens in plants. The key point is
that there are several steps, involving a number of different
proteins that must occur in sequence between the recognition
of the replication origins (see below) and the loading of the
initiating DNA polymerase. These steps culminate in the formation of the pre-replicative complex. I have reviewed this
feature of DNA replication relatively recently (Bryant, 2010;
2566 | Bryant
Bryant and Aves, 2011); but here the focus is on the main
steps (omitting in particular, regulation by phosphorylation
of some of the main ‘players’: see below and Bryant, 2010).
Origins that have been recognized by an origin-recognition
complex (see below) are activated by the binding of CDC6
and the MCM2–7 helicase complex. Sometimes this event is
referred to as licensing of the origin. CDC6 is then displaced
and degraded and is replaced by MCM10; MCM2–7 and
MCM10 then recruit CDC45 to the origin and this in turn,
loads primase-polymerase-α onto the template (but whether
just as the primase-polymerase tetramer or as a larger complex is not clear).
The fourth line of DNA polymerase-related research concerns regulation. Regulation can be approached in at least four
ways. The first is post-translational regulation of pre-existing
enzymes. The second, moving up a level, is the control of
transcription of the genes encoding the polymerases and of
other S-phase associated proteins. The third, one level higher
still, concerns the hormones and other factors that interact
with the transcriptional regulators, often acting via cyclin–
cdk complexes. This starts to bring us into contact with plantspecific regulatory systems and leads to the highest level of
all, regulation of all aspects of cell cycle activity in relation
to plant growth, development, and morphogenesis. Space
does not permit detailed discussion of the two higher levels
of regulation: the reader is referred to papers by Murray and
his colleagues (Meijer and Murray, 2000; Dewitte et al., 2003;
Menges and Murray, 2008; Nieuwland et al., 2009; Xie et al.,
2010) and by Sanchez et al. (2012).
In relation to transcriptional control, many proteins active
in the S-phase are regulated by a sequence of events involving RB (retinoblastoma) protein (in plants, RBR: retinoblastoma-related protein) and the E2F/DP family of transcription
factors (Attwooll et al., 2004). The E2F/DP transcription factors are kept inactive by RBR protein. The phosphorylation
of RBR (in plants by CYCD–CDKA; Menges and Murray,
2008) causes it to dissociate from E2F/DP which then upregulate, in late G1, the genes encoding S-phase proteins.
In animal cells, it is known that all three replicative DNA
polymerases and PCNA are amongst the proteins regulated
in this way. This has not been directly shown for the plant
DNA polymerases although patterns of polymerase activity in relation to DNA replication are consistent with this
system being operative. This has been clearly demonstrated
for plant PCNA (Uemukai et al, 2005) and CDC6 (de Jager
et al., 2001). It is also clear however, that not all of the proteins involved in DNA replication are regulated strictly in
relation to S-phase. In our own work, for example, while
the transcription of the gene encoding CDC6 (see above) in
tobacco BY2 cells occurs only in late G1 and S, transcription
of the gene encoding MCM3 occurs throughout the cell cycle
(Dambrauskas et al., 2003).
At the post-translational level of regulation, DNA
polymerase-α is a phospho-protein which, in maize (corn),
becomes phosphorylated by a CDK–cyclin complex during the S-phase (Roig and Vasquez-Ramos, 2003). It is difficult to ascertain the role of this phosphorylation from the
data on maize but there is evidence from mammals that
Cdk2–cyclinA-mediated phosphorylation of both the catalytic (180 kDa) and the 68 kDa subunits increases polymerase
activity. Curiously, the enzyme also appears to be stimulated
by an interaction between hyper-phosphorylated RB protein
and the catalytic subunit, an interaction that can only occur if
the 68 kDa subunit is phosphorylated (Takemura et al., 2006).
The possible interaction between DNA polymerase-α and
hyperphosphorylated RB protein introduces another aspect
of post-translational control, namely that enzyme activities may be regulated by interaction with other proteins.
This may occur within specific groupings of polypeptides,
as with the heterotetrameric DNA polymerase-α-primase.
Interactions can also occur between less tightly linked proteins. The significant stimulation of DNA polymerase-α
activity in vitro by primer-recognition protein has already
been mentioned (even if we do not understand the in vivo
role of this interaction). Another example concerns PCNA,
the DNA polymerase-δ processivity factor. In Arabidopsis
it has been shown that two SET-domain proteins, ATXR5
and ATXR6 interact with PCNA and that this interaction
relates to PCNA’s role in DNA replication (Raynaud et al.,
2006). ATXR6, in particular, accumulates in the nucleus
during the S-phase; this involves a transcriptional up-regulation under the control of the E2F transcription factor(s) as
described above.
There are doubtless many more details concerning the
subtleties of post-translational control that remain to be discovered. However, it is worth emphasizing at this point that
plants certainly do regulate the rate of DNA synthesis in
vivo. This may be achieved by varying the number of replication origins that are activated, as discussed below. However,
in some situations, for example, the induction of flowering
in Sinapis alba, the increased rate of DNA replication and
hence shortening of the S-phase cannot be attributed solely
to the use of more replication origins. It must also involve
an increase in the actual rate of synthesis (Jacqmard and
Houssa, 1988). Given what we know about the interaction
of enzymes and other proteins with the template it seems
unlikely that more molecules of a particular enzyme could
be recruited. This suggests, in turn, that the activity/activities
of one or more enzymes (whether topoisomerase, working in
front of the replication fork, MCM-helicase separating the
strands, primase and the polymerases actually synthesizing
the daughter strands) is/are stimulated. As already shown,
there are several mechanisms of achieving such a stimulation.
We simply do not know which mechanisms operate, nor on
which enzymes.
DNA replication origins
DNA replication origins are perhaps the most frustrating
structures in DNA on which to work. They are readily visualized by fibre autoradiography. By appropriate modifications
to the technique, it can be shown that replication proceeds
outwards in two directions from each origin and, further, that
the speed (‘fork rate’) at which this happens can be measured.
In any one plant tissue, origins are relatively evenly spaced,
leading to questions about the type of chromatin feature that
Cell cycling with the SEB: a personal view | 2567
would lead to this spacing. Is it a regularly occurring DNA
sequence or a feature of higher-order chromatin structure,
or both, or neither? Further, in any one DNA fibre, some
tracts of DNA were not active in replication while others
were active. This gave rise to the concept of replicon families, or time-groups, each with their own ‘window’ during the
S-phase in which they are active in replication. Jack Van’t
Hof (Van’t Hof et al., 1978) went on to analyse this in detail
for Arabidopsis thaliana, showing that its small genome was
organized as just two replicon families. This is also indicated
by the results of a more recent molecular analysis of just
one Arabidopsis chromosome (Lee et al., 2010). This leads
to further questions, in particular, what is it that makes one
replicon different from another? Do their replication origins
differ or is it a matter of chromatin organization and hence
accessibility?
Thus, fibre autoradiography gives us a lot of information but raises many questions. It would be helpful to isolate and characterize the origins and then to investigate their
interaction with the DNA replication machinery. But this is
where the frustration lies. Fibre autoradiography gives no
clue about sequences or about chromatin organization. The
investigator does not know what to look for. Further, each
origin is active only transiently, unlike gene promoters; it is
therefore very difficult to catch a replication origin in the act.
This then describes the state of affairs at the time I started to
be interested in replication origins; my interest was focused
on how the replicative enzymes interacted with the origins
and the role these interactions might have in the regulation
of replication.
As this work continued, two models began to dominate,
as I have described elsewhere (Bryant, 2010). One was replication in mammalian cells of the virus SV40. This virus utilizes host replicative machinery except for a virus-encoded
helicase that recognizes a specific sequence in the viral
genome. The second was the replication of plasmids containing ars1 elements from budding yeast (Saccharomyces
cerevisiae). The ars element is an 11 bp, AT-rich motif
which confers on any plasmid the ability to be replicated
in a yeast cell. It was quickly established that the motif did
indeed represent an essential sequence of the yeast replication origin. This was further confirmed when a complex of
six proteins, the origin recognition complex, was isolated
from yeast and shown to bind in a sequence-dependent
manner to the yeast replication origin (Bell and Stillman,
1992). Everything was pointing to sequence as a key feature
of replication origins.
To some extent, this view was backed up by sequence
analysis of the so-far only unequivocally identified replication origin from plant nuclear DNA (Van’t Hof and Lamm,
1992). This origin lies within a 1500 bp region in the nontranscribed spacer of the repeated genes encoding rRNA
(Van’t Hof et al., 1987). It is AT-rich and contains four
matches to the yeast ars core sequence (Hernández et al.,
1
Originally named for conferring autonomous replication in
Saccharomyces, the terminology has now mutated to autonomously
replicating sequence.
1988); further, plant genomes in general contained numerous matches to this motif (Sibson et al., 1988), although
whether they functioned as origins in planta was never
demonstrated.
However, as reviewed in Bryant (2010) the consensus
began to move away from the idea that most replication
origins were, or contained, specific sequences. Even in fission yeast, the ORC binds to AT-rich tracts rather than to
highly specific AT-rich motifs. The dominant view is that,
although some unusual origins of replication do contain
specific essential AT-rich sequences, the majority of animal
replication origins, detected and analysed by modern techniques that allow genome-wide trawling and rapid sequence
analysis, do not (Bryant, 2010). Nevertheless, AT-richness
is a general feature and many have tracts of curved or bent
DNA. However, an unexpected feature is emerging as more
and more animal replication origins are analysed. Most,
but not all of the animal replication origins analysed to
date contain, in addition to AT-rich DNA, CpG islands
and short G repeats known as OGREs2 (Cayrou et al.,
2012; Maizels and Gray, 2013). The disposition of the G
repeats is such that the DNA is likely to form a transient
quadruplex (Maizels and Gray, 2013). Since the actual initiation site is situated a specific distance from the OGRE (e.g.
280 bp in mouse: Cayrou et al., 2012) it is suggested that,
in these OGRE-containing origins, the OGRE is essential
for initiation while the AT-rich region of each origin is the
site of initial strand separation. It is, however, still not clear
where the ORC binds within the overall structure of an origin. The newer genome-wide analyses and DNA combing
techniques have not yet been widely applied to plants but
the limited information available to us hints that replication origins in Arabidopsis also contain tracts rich in G and
C residues (Costas et al., 2011; Sanchez et al., 2012; Bass
et al., 2014).
The other origin-related topic that interested both
myself and Dennis Francis was the variation in the spacing and therefore the number of origins that are used
in any particular situation; see for example, Kidd et al.
(1992). Spacing can change in developmental transitions
such as the induction of flowering (Jacqmard and Houssa,
1988; Durdan et al., 1998); it can be modified by treatment with hormones and other growth regulators (Houssa
et al., 1994; Jacqmard et al., 1995; Mazzuca et al., 2000).
It can also be changed by ‘replicative stress’ (Woodward
et al., 2006). An example of the latter is the effect of
cross-linking the DNA prior to the onset of the S-phase,
as carried out by our research group in an attempt to
trap replication origins (Francis et al., 1985). What actually happened was that new origins, located between the
cross-links, were utilized as replication forks stalled at the
cross-links. Although this work got us no closer to isolating replication origins, it did lead us to suggest that there
replication origins of different ‘strengths’, a suggestion
amply confirmed by more recent work on the events that
lead to an origin being ‘fired’.
2
Origin G-rich repeated elements.
2568 | Bryant
These events have been mentioned in passing (above) and
have also been reviewed in more detail elsewhere (Bryant,
2010; Bryant and Aves, 2011). I want to focus here on the
first two or three steps. Origins that have been recognized and
bound by the ORC are ‘licensed’ by the binding of CDC6
and the MCM2–7 helicase complex (see Schultz et al., 2009,
for further discussion). In animal cells, MCMs 2–7 are in a
large excess over the amount required to license the origins
that are active in any one S-phase, (Woodward et al., 2006)
leading to the suggestion that many origins that are licensed
may not actually be used, or in the words of Mel de Pamphilis
(1993) ‘Many are called but few are chosen’. We may surmise,
on the grounds of cellular economy, that it is the next step
that is rate-limiting, namely the phosphorylation of ORC
and of CDC6 by CDC7/DBF4 and CDC28/CLN1 but it
may actually be any one of the steps prior to the loading of
polymerase-α-primase. Whichever of these steps it is, there
are still questions: what actually determines which licensed
origins become active and how is this process affected by
developmental transitions, hormones, and growth regulators?
There is still a lot to discover before we reach a fuller understanding of the regulation of DNA replication.
DNA endoreduplication
An analysis of cell cycle arrest in relation to the onset of differentiation in plant organs quickly reveals that there are cells
arrested in G1 with 2C amounts of DNA and cells arrested
in G2 with 4C amounts of DNA. This could be interpreted
as a reflection of the interactions between the control of the
cell cycle and the control of differentiation; some dividing
cells had simply progressed further through the cell cycle
when division activity was inhibited or lost. However, it is
not as simple as this. Firstly, as noted by Evans and Van’t
Hof (1974), the patterns of G1 or G2 arrest vary between
species which suggests subtle differences in timing/patterning
that may be species-specific. Secondly, in many types of differentiated cells, further rounds of DNA replication occur in
the absence of mitosis: the DNA is endoreduplicated. Thus,
in Arabidopsis leaves, there are many cells with 8C amounts
of DNA (Galbraith et al., 1991).
Storage tissues of seeds, such as cereal endosperm and the
cotyledons of leguminous plants such as pea and bean, contain many cells with endoreduplicated DNA, often up to 32C
and 64C. Similarly, cells in the endocarp of fleshy fruit, such
as tomato, may reach 256C or even 512C: Chevalier et al.,
2011, 2014). However, the most spectacular examples occur in
highly specialized cells such as those of the embryo suspensor
in dicots. In Phaseolus coccineus, for example, some cells in the
suspensor may attain DNA amounts of 16 384C, involving 13
rounds of DNA replication in the absence of mitosis.
In a conventional cell cycle there are multiple checkpoints
and other regulatory mechanisms to ensure that the licensing of the replication origins mentioned above can only occur
in cells that have passed through mitosis. In cells undergoing
endoreduplication, these controls are bypassed or over-ridden. Space does not permit a detailed discussion of this aspect
of endoreduplication except to say that there are mechanisms
operating at organ, tissue, and cell level that allow breakage
of the normal rules (Larkins et al., 2001; Ishida et al., 2010;
Chevalier et al., 2011; Sterken et al., 2012; Roeder, 2012; Jun
et al., 2013). My focus here is on biochemical aspects followed by a brief mention of the possible function(s) of DNA
endoreduplication.
The formation of the pre-replicative complex at the replication origins has been mentioned in earlier sections of
this review. In all respects it appears that this happens in the
same way in endoreduplication as in replication (Sabelli et al.,
2013). CDC6 and to a lesser extent CDT1 (see above and
Bryant, 2010) are especially important players at this stage of
the process (Castellano et al., 2001, 2004; Bryant, 2010). In a
‘normal’ cell cycle, phosphorylation of CDC6 followed by its
degradation is part of the transition from a licensed origin to
an activated origin. In cells undergoing repeated rounds of
DNA replication, this degradation does not occur. If CDC6
is over-expressed, DNA endoreduplication is induced in cells
which would otherwise be undergoing normal cell cycles.
Further, in tobacco BY2 cells cultured in the absence of
auxin, cell division does not occur but after four to five days,
endoreduplication is initiated (but see below). Associated
with this is a renewal of CDC6 expression (Bryant, 2010).
Although the pre-replicative events associated with endoreduplication parallel those in normal replication, there are questions about which DNA polymerase(s) is/are involved in the
synthesis phase. For example, we (Quélo et al., 2002) showed,
somewhat unexpectedly, that in cultured tobacco cells, endoreduplication still occurred if the replicative polymerases were
inhibited by aphidicolin; it was only prevented if an inhibitor of DNA polymerase-β was also added. Inhibition of only
polymerase-β did not prevent endoreduplication, suggesting
that the process may be mediated by the replicative polymerases and/or a repair-associated enzyme, DNA polymerase-β.
That conclusion may run counter to our understanding of
DNA biochemistry, but even more puzzling are the data from
Roy et al. (1988, 2007a, b). DNA endoreduplication occurs in
the cotyledons of developing mung bean seedlings (as in other
legumes) but the major polymerase in cotyledon cells is DNA
polymerase-β. The authors suggest that it is polymerase-β,
rather than the replicative polymerases, that mediates DNA
endoreduplication. The suggestion is supported by the finding that this particular form of polymerase-β can interact with
PCNA which significantly increases its processivity.
Several functions have been ascribed to DNA endoreduplication as reviewed by (Chevalier et al. 2011, 2014). The two most
widely suggested are that it drives cell expansion in situations
of rapid growth (cell volume increases in response to nuclear
volume: the karyoplasmic ratio theory) and/or that it allows
high levels of gene expression via a gene dosage effect, for
example, in order to lay down large amounts of storage products in seeds. In respect of the former, there are many examples of large cells with endoreduplicated DNA, including those
in fleshy fruit (mentioned above) but there are fruit with large
cells that do not contain endoreduplicated DNA. Chevalier
et al. (2011) review evidence showing that, in fruit, the occurrence or not of endoreduplication is correlated with the rate of
expansion: rapid expansion requires DNA endoreduplication.
Cell cycling with the SEB: a personal view | 2569
However, data from tobacco BY2 cells suggests that in some
systems, cell expansion may precede endoreduplication. As
mentioned above, sub-culture of cells in the absence of auxin
leads to cell expansion rather than division. Endoreduplication
certainly occurs but not in all cells, and only after there has
been significant cell expansion (Quelo et al., 1992).
In respect of the gene dosage hypothesis, the relationship
in legume cotyledons between DNA endoreduplication, cell
expansion, and storage product deposition are certainly consistent with the view. However, our study of oil-seed rape
(Brassica napus) cotyledons (Silcock et al., 1990) showed
that endoreduplication does not occur in the large lipid- and
protein-filled cells. Thus, plans to study the effects of disrupting endoreduplication were very effectively prevented, a clear
example of the frustrations that sometimes occur in research,
even when clear data have been obtained!
Acknowledgements
Bray CM, Sunderland PA, Waterworth WM, West CE. 2008.
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and evolutionary aspects. Annals of Botany 107, 1119–1126.
Bryant JA. 2008. Copying the template—with a little help from my
friends? In: Bryant JA, Francis D, eds. The eukaryotic cell cycle.
Abingdon: Taylor and Francis, 71–80.
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Bryant JA, Dunham VL. 1988. The biochemistry of DNA replication.
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I am very grateful to Walter Dewitte and Jim Murray and
to the SEB, for organizing the Plant Cell Cycle sessions at
the Society’s 2013 Annual Main Meeting in Valencia. I am
also grateful to all those with whom I have had the privilege
of collaborating over the years, including Louise Anderson,
Steve Aves, Donato Chiatante, Val Dunham, Dennis Francis,
Steve Hughes, Hilary Rogers, Tom Rost, Jack Van’t Hof and
Jean-Pierre Verbelen. I thank all those post-graduate students
and post-docs who have worked in my lab. The work of some
of these lab members is mentioned in this paper but all have
contributed a lot to ‘The Bryant lab’. Finally, I must mention with gratitude the work of Paul Fitchett who, over many
years, was the best research technician that one could wish
for: thorough, patient, very hard-working and with very high
ethical standards. Without him, the lab would have been very
much less productive.
Bryant JA, Francis D. 1985. The cell division cycle in plants. SEB
Seminar Series, 26. Cambridge: Cambridge University Press.
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