Plant Cell Physiol. 46(6): 931–936 (2005)
doi:10.1093/pcp/pci100, available online at www.pcp.oupjournals.org
JSPP © 2005
Circadian G2 Arrest as Related to Circadian Gating of Cell Population
Growth in Euglena
Aoen Bolige, Shin-ya Hagiwara, Yulan Zhang 1 and Ken Goto 2
Laboratory of Biological Rhythms, Obihiro University of Agriculture and Veterinary Medicine, Obihiro, 080-8555 Japan
;
cycle machinery. One of these has only recently been found in
the algal flagellate Euglena gracilis grown photoautotrophically, and it determines when in the circadian cycle light is
effective for inducing the capability of G2-phase cells to
progress to cell division in darkness (Hagiwara et al. 2002).
That is, whereas darkness prevents G2 cells from progressing to
mitosis unless they have been irradiated sufficiently beforehand
to acquire the ability for, or commit to, cell division in the following darkness (Hagiwara et al. 2001), a circadian rhythm
regulates the timing of the photoinduction of the ability of cells
to proceed to mitosis in the absence of light, such that while 6 h
of illumination is not at all effective around subjective dawn, at
around subjective dusk it does induce this ability (Hagiwara et
al. 2002). This circadian light response may underlie the ‘photoinducible phase’ for photoperiodic induction of cell reproduction in this alga (Hagiwara et al. 2002), because the external
coincidence model of photoperiodism supposes that a long-day
response occurs only when light falls during the ‘photoinducible phase’ (cf. Davis 2002).
This study is concerned primarily with another type of circadian rhythm, which may be intimately related to the rhythm
mentioned above and has been known for over four decades to
regulate the timing of cell population growth in many organisms (Sweeney and Hastings 1958, Sweeney 1987, Edmunds
1988, Goto and Johnson 1995), including cyanobacteria (Mori
et al. 1996) and humans (Canaple et al. 2003). Although the
underlying clock may be common to both, the rhythms may
differ in their downstream (output) pathways, and will be designated as CRdusk for the former and CRG2 for the latter; the
former is responsible for the maximal photoinduction at dusk,
while the latter prevents G2 cells from progressing to mitosis,
as concluded in this study.
Perhaps, the most extensive studies on CRG2 have been
conducted in E. gracilis (Edmunds and Funch 1969, Goto et al.
1985, Edmunds 1988, Carré and Edmunds 1993) in which cell
population growth is timed to occur only during subjective
night, i.e. an endogenous state that is not only attained during
12 h dark intervals of 24 h light/dark (LD) cycles but also
recurs with a circadian period, 26 h in this alga, in an environment without an external time cue; the subjective night is from
subjective dusk or circadian time (CT) 12 to subjective dawn or
CT24 (=00).
Cell population growth is gated to occur in particular
circadian phases, which has been known for over four decades in various organisms including cyanobacteria and
human. However, little is known as to which cell cycle
phases from G1 to M are primarily regulated by the
circadian rhythm or when in a circadian cycle this primary
regulation takes place. We report here that in the flagellate
alga Euglena gracilis grown photoautotrophically, the
circadian rhythm primarily prevented developmentally
matured G2 cells from progressing to mitosis, such that cell
population growth occurred only during subjective night.
In addition, we found that the circadian rhythm also arrests
G1-to-S and S-to-G2 transitions at particular circadian
phases.
Keywords: Cell cycle — Circadian rhythms — Euglena gracilis — G2 arrest — Mitosis.
Abbreviations: CT, circadian time; DD, continuous darkness;
FCM, flow cytometry; GT, generation time; LD:1,1, alternation of 1 h
light plus 1 h darkness; LL, continuous light; MPF, maturationpromoting factor.
Introduction
The molecular mechanisms underlying cell cycle progression have been well studied in the past decade. Progression is
regulated by the autonomous cycling of the cell cycle engine,
which is composed of maturation- or mitosis-promoting factor
(MPF), and through various checkpoints (Hartwell and Weinert
1989, Murray and Kirshner 1989, Elledge 1996, Nurse 2000,
De Veylder et al. 2003, Murray 2004). Recently, Matsuo et al.
(2003) demonstrated that in the mouse, the activity of MPF follows a circadian rhythm with a phasing such that it can be the
direct cause of the circadian timing of mitosis, although the
activity of WEE1 kinase, which inhibits MPF activity, remains
high when mitosis occurs or MPF is active. However, most cell
cycle biologists have overlooked the fact that the cell cycle
machinery is not the sole way of regulating the progression of
the cell cycle.
There are at least two kinds of circadian control over cell
cycle progression, both of which run independently of the cell
1
2
Present address: Institute of Crop Breeding & Cultivation, Chinese Academy of Agricultural Sciences, Beijing, China.
Corresponding author: E-mail, [email protected]; Fax, +81-155-49-5612.
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Circadian gating of G2-to-M transition in Euglena
most importantly, we can assess whether the cells are developmentally ready for cell division by placing the culture in LD:
1,1 into DD (Hagiwara et al. 2001, Hagiwara et al. 2002).
Results
Fig. 1 Cell population growth in LD:1,1. Cultures dividing arrhythmically in LL were transferred to LD:1,1 at either 4 (filled circles) or
6 klux (open circles). This transfer was sufficient to evoke circadian
rhythms in gated cell growth. Hatched and open bars represent subjective nights and days, respectively, in Fig. 1–3.
An important question remains unsolved: which cell cycle
phase from G1 to M is the primary site of circadian regulation,
and at which circadian phases does this regulation take place?
The work by Carré and Edmunds (1993) is intriguing because
they found a considerable number of cells in G2 + M phase in
an achlorophyllous mutant ZC strain of E. gracilis grown heterotrophically, even in the subjective day when cell population
growth does not take place. This led us to ask whether those G2
+ M-phase cells are developmentally ready for cell division: if
so, they may serve as unequivocal evidence for a circadian
rhythm that prevents mature G2 + M-phase cells from completing cell division during the subjective day. The work of
Matsuo et al. (2003) also suggests the circadian inhibition of
the G2-to-M transition, because, although surgical removal of
a portion of the liver induces widespread proliferation among
residual hepatocytes and DNA synthesis occurs maximally at a
fixed time after surgery, the resulting G2 cells will enter mitosis only at a particular phase of the circadian cycle.
This study sought to answer these questions by using
photoautotrophic E. gracilis under 2 h LD cycles of 1 h light
plus 1 h darkness (LD:1,1), where both CRdusk and CRG2 interact. We chose this protocol instead of using the heterotrophic ZC
mutant in DD for the following reasons. First, the most extensive studies on CRG2 have been conducted under these conditions (Goto et al. 1985, Edmunds 1988). Secondly, cell cycle
speeds can be manipulated easily by changing light intensities
and we are interested in showing the independence of the timing of cell cycle transitions from cell cycle speeds. Finally, and
Circadian control of cell population growth
In E. gracilis, circadian gating of cell population growth
can be manifested without prior entrainment to 24 h LD cycles.
Therefore, when asynchronously dividing cultures in LL were
placed into LD:1,1, which does not serve as a time cue
(Edmunds 1988), the cell cycle became regulated by a circadian rhythm, and cell population growth did not occur during
the subjective day lasting ∼13 h of each ∼26 h cycle (Fig. 1);
the gated cell population growth began at 24, 50, 76 and 102 h
in LD:1,1. Since the period length was similar at 4 or 6 klux,
while the average generation time (GT) differed by >2-fold (55
or 26 h, respectively), we concluded that circadian rhythm in E.
gracilis is distinct from the cell cycle machinery, as was demonstrated in Chlamydomonas reinhardtii (Goto and Johnson
1995).
Note that as the onset of cell population growth is defined
as CT12 (subjective dusk) and the circadian period was ∼26 h
in this alga, the circadian rhythm was reset to CT14 at the
transfer from LL to LD:1,1.
Circadian rhythms in cell cycle transitions
To understand why cell population growth does not occur
during the subjective day, the DNA contents of 11 cultures
grown in LD:1,1 were determined by flow cytometry (FCM).
The kinetic behavior of the cell cycle phases, from 15 circadian cycles for the 11 cultures, was averaged to a single circadian cycle for statistical analysis. Average data for five
cycles with a factorial increase in cell number or step size from
1.23 (GT of 87.1 h) to 1.46 (GT of 47.6 h) are shown in Fig.
2A and B, and data for all 15 cycles, with a step size from 1.23
to 1.75 (GT of 32.2 h), are shown in Fig. 2C. The flux rates of
all three transitions, i.e. the cell numbers per 2 h that underwent
G1-to-S, S-to-(G2 + M) and (G2 + M)-to-G1 transitions, displayed circadian rhythms (Fig. 2B).
Circadian gating of G2-to-M transitions
Cells in the G2 + M phase were present throughout the
26 h cycle (Fig. 2A, C). However, these cells did not complete
cell division during the subjective day (CT00 to CT12, Fig. 1,
2A), suggesting that either the G2-to-M or the M-to-G1 transition did not occur. In order to determine which transition did
not occur, we observed temporal changes in mitotic cell numbers under LD:1,1 at 9 klux (Fig. 3). Mitotic cells appeared at
46 and 72 h (i.e. CT08; Fig. 3A), while population growth
began 4 h later, at 50 and 76 h (i.e. CT12; Fig. 3C), indicating
that mitosis lasted ∼4 h. There were few, if any, mitotic cells
from CT00 to CT08. Therefore, G2 + M-phase cells (Fig. 1,
2A; 20–30% as estimated by FCM) were in fact G2-phase cells.
Circadian gating of G2-to-M transition in Euglena
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Fig. 3 Mitosis under LD:1,1 (9 klux). Vertical bars represent the
maximum (A) or onset of (C) cell population growth. (A) Cells in
interphase, prophase or mitosis (i.e. prophase to telophase). (B) Cells
in metaphase, anaphase or telophase. (C) Cell population growth.
Fig. 2 Cell cycle transitions in LD:1,1. For simplicity, the number of
cells in the first sample was assigned the value 100. Averaged values
and SEMs from five circadian cycles (step size between 1.23 and 1.46)
are shown in (A) and (B), while values from all 15 cycles in 11 independent cultures (step size between 1.23 and 1.75) are shown in (C).
(A), (C), number of cells; (B), flux rates per h.
Consequently, G2 cells did not progress to mitosis between
CT00 and CT08.
Furthermore, since cell division ended at CT00, the 4 h
duration of mitosis in LD:1,1 indicates that the G2-to-M transition began to stop at approximately CT20. Therefore, the G2to-M transition did not occur between CT20 and CT08, which
resulted in the circadian ‘gating’ of both mitosis between CT08
and CT20 and cell population growth (or cytokinesis) during
the subjective night (CT12 to CT00).
A most important question is why the G2-to-M transition
did not occur between CT20 and CT08. Were the G2 cells
present during this period not ready for cell division developmentally, in terms of cell cycle phase? Or, if they were ready,
did a circadian rhythm prevent the developmentally mature G2
cells from progressing to mitosis? To answer these questions,
the culture under LD:1,1 was transferred to DD, because darkness prevents G2-phase cells of the photoautotrophic Euglena
from progressing to cell division if they are not ready for cell
division in darkness, whereas sufficient light irradiation affords
them the capability of, or the commitment to, cell division in
darkness (Hagiwara et al. 2001, Hagiwara et al. 2002).
As shown in the bottom curve in Fig. 4, the DD transfer at
CT01 did not completely suppress cell population growth during the following subjective night; moreover, it did not change
the timing of cell division, indicating unequivocally that there
were G2 cells, 14% of the total, that were committed to cell
division by the transfer at CT01 and that they did not progress
to mitosis until CT08. Therefore, cell population growth did
not occur even in the presence of the committed G2 cells.
CT20 is the final phase of the entry into mitosis (Fig. 3)
that, once started, can be completed in DD (Hagiwara et al.
2001, Hagiwara et al. 2002). Not surprisingly, DD transfer at
CT21, ∼1 h after the final phase, had little effect on the cell
population growth in the current night, as can be seen in the
small changes in the factorial increase of cell numbers with the
DD transfer. The small effect in the current night also shows
that if there were committed G2 cells, they did not progress to
mitosis between CT21 and CT24 (=CT00) or CT08. The pres-
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Circadian gating of G2-to-M transition in Euglena
ence of committed G2 cells at CT21 is evidenced by the additional cell population growth, although much smaller than in
the previous example, during the next night.
The earlier transfer to DD (CT12 or CT16) did not give
rise to cell population growth the next night (upper two curves
of Fig. 4), indicating that all of the cells that had committed by
CT12 or CT16 progressed to mitosis by CT20 in the current
night. As expected, the factorial increase in the cell numbers of
the darkened cultures was smaller than in the previous cycles
under LD:1,1.
Collectively, these results unequivocally reveal that the
absence of the G2-to-M transition between CT20 and CT08
does not result from an absence of committed G2 cells, and that
the timing of when G2 cells commit indirectly determines when
they progress to mitosis; G2 cells that committed to cell division between CT12 and CT20 progressed to mitosis by CT20
that night, but those that did so after CT20 did not progress to
mitosis until CT08 the following subjective day. Therefore, we
conclude that a circadian rhythm prevented the committed G2
cells from progressing to mitosis between CT20 and CT08.
In addition, note that since our cultures were not dividing
synchronously, i.e. all the cells did not divide in one night; a
simple inspection of Fig. 2B does not reveal information concerning the duration of the G1, S or G2 + M phases. Indeed, all
of the transitions, G1-to-S, S-to-G2 and G2 + M-to-G1, occurred
maximally at the same circadian phase, i.e. around subjective
midnight. This coordination is ascribable to the combined
effect of the circadian regulation of the timing of cell cycle
transitions and the fact that the average GT is longer than the
26 h circadian period.
Circadian gating of the S-to-G2 transition
Fig. 2A and C also reveals that the number of cells in G2 +
M phase did not change significantly between CT00 and CT12.
Since no cells exited from G2 + M phase during this period, the
constant number of G2 + M-phase cells can be attributed to the
fact that no cells entered the G2 + M phase. Therefore, there
were also no S-to-G2 transitions between CT00 and CT12, as
can also be seen from the flux rates of the S-to-G2 transition in
Fig. 2B, whereas there was a significant number of S-phase
cells during this period (Fig. 2A, C).
Note that while DNA synthesis occurred between CT00
and CT12, it was not completed. Ongoing synthesis was evident not only from the increase in the numbers of S-phase cells
during late subjective day (Fig. 2A), but also from the change
in the shape of the S compartment of the DNA histograms
throughout the subjective day (data not shown). Therefore, the
circadian gating of the S-to-G2 transition may result either from
slowing the progression of S phase or by arresting S-phase
cells late in S phase.
Circadian gating of the G1-to-S transition
The number of S-phase cells was constant between 62 and
66 h (CT23 to CT03) in LD:1,1 (Fig. 2A and C), during which
time the cells did not exit from S phase. Therefore, there were
also no entries into S phase (i.e. no transition from G1 to S).
The number of G1 cells was unchanged during this period (Fig.
2A and C), when G1 cells were not produced. The absence of
this transition is also shown by the low flux rates of G1-to-S
transition at 63 and 65 h (Fig. 2B).
At the 66th hour, after the interval of no G1-to-S transition from CT23 to CT03, the number of G1 cells began to
decrease, while the number of S cells began to increase, indicating that the G1-to-S transition began around CT03. In Fig.
2B, this is reflected in the positive flux rate of the G1-to-S transition at 67 h in LD:1,1.
Discussion
In this study, DNA FCM analysis, together with microscopic observations, revealed that in E. gracilis grown photoautotrophically under LD:1,1, G2-to-M transitions took place
only during a particular circadian phase, between CT08 and
CT20 (Fig. 2, 3), and that mitosis, once it had occurred at any
time between CT08 and CT20, was completed for ∼4 h (Fig.
3). Between CT20 and CT08, G2-to-M transitions did not
occur, even in the presence of committed G2 cells (Fig. 4).
Therefore, we conclude that a circadian rhythm (CRG2) prevents G2 cells from progressing to mitosis and thereby regulates the timing of the G2-to-M transition, such that cell
population growth is allowed to take place only during subjective night, between CT12 and CT24 (=00) (Fig. 1). Therefore,
we suggest that, biochemically, CRG2 somehow prevents the
activation of MPF between CT20 and CT08.
Unexpectedly, we also found that a circadian rhythm regulates the timing of the G1-to-S and S-to-G2 transitions that did
not take place between CT23 and CT01 or CT00 and CT12,
respectively (Fig. 2). With reasoning similar to that described
below for the circadian arrest of the G2-to-M transition, this
might be ascribable to circadian arrest or slow down. We
should examine whether there are G1 and S cells that are
developmentally ready for cell cycle transitions at the circadian
phases in which no G1-to-S and S-to-G2 transitions occur.
Circadian arrest of G2-to-M transitions
The most important question is why the committed G2
cells did not progress to mitosis between CT20 and CT08.
Were they still not developmentally mature or ready for cell
division? Or, did a circadian rhythm prevent them from progressing to mitosis?
Why did committed G2 cells not progress to mitosis? As
shown in Fig. 4, all of the G2 cells that committed between
CT12 and CT20 progressed to mitosis by CT20 and completed
cell division by CT24 (=00) in the current night, whereas those
that committed after CT20 did not actually progress to mitosis
until CT08 the next day. Obviously, committed G2 cells could
not progress to mitosis between CT20 and CT08.
The circadian rhythm (CRG2) somehow prevents committed G2 cells from progressing to mitosis between CT20 and
Circadian gating of G2-to-M transition in Euglena
Fig. 4 Cell population growth after the transfer to DD from LD:1,1.
Cultures in LL at 6 klux were transferred to LD:1,1 and then transferred to DD at either CT12, CT16, CT21 or CT01; aligned from top
to bottom. Filled symbols represent growth curves in DD where the
first points marked the time of the transfer to DD. Open and hatched
rectangles represent subjective days and nights, respectively. Numbers
attached to the left of the growth periods each depict a factorial
increase of cell number in a circadian cycle.
CT08; how? One might suppose that the committed G2 cells
are still not developmentally ready for cell division, and that
they require a dark process for maturation. This hypothesis also
supposes that a circadian rhythm suppresses that developmental process between CT20 and CT08. Alternatively, one could
suppose that the committed G2 cells are developmentally ready
at the time of the commitment and that the circadian rhythm
prevents the mature G2 cells from progressing to mitosis. Since
the circadian timing of cell population growth was unaffected
by any change in cell cycle speed (Fig. 1), we suggest that the
latter is much more probable.
However, one might propose counter-hypotheses that do
not suppose an involvement of circadian G2 arrest. Therefore,
we will present evidence against such counter-hypotheses.
Is a ‘commitment timer’ involved in circadian gating of
cell population growth? There are a multitude of ‘commitment’ phenomena in E. gracilis grown photoautotrophically
935
(Hagiwara et al. 2001): the cells in G1 phase are able to commit to progressing to G2 phase, but not to cell division; S cells
to S-to-G2 transition or cell division; and G2 cells to cell division. The cell cycle phase to which they commit depends on
three endogenous factors: the later in each of the cell cycle
phases, G1, S or G2 (Hagiwara et al. 2001); the stronger the
redox signal originating in non-cyclic photosynthetic electron
transport that senses environmental light intensities (Hagiwara
et al. 2002); and the nearer to subjective dusk in a circadian
cycle (Hagiwara et al. 2002), the more likely cells are to commit to dark progression to the later phase.
If a circadian rhythm regulates the timing of commitment
to cell division, and if the post-commitment duration between
the time of commitment and cell division is maintained constant, as was believed to be ∼6 h in the case of C. reinhardtii in
which only the commitment by G1 cells to cell division is
known (Spudich and Sager 1980, Donnan and John 1983,
Krupinska and Humbeck 1994), it might be possible that the
combined effect of the circadian timing of commitment and of
developmental control is responsible for the circadian gating of
cell population growth, as we speculated previously (Goto and
Johnson 1995).
However, in photoautotrophs, commitment to cell cycle
transitions is required only for those transitions in darkness, but
not those in light (Hagiwara et al. 2001). Therefore, the
hypothesis, if valid, cannot be a universal mechanism because
both heterotrophs in DD (Edmunds 1974, Carré and Edmunds
1993) and photoautotrophs in LL (Sweeney and Hastings 1958,
Edmunds and Funch 1969, Goto and Johnson 1995, Mori et al.
1996) display circadian gating of cell population growth,
although it might be applicable, at least theoretically, to the
photoautotrophs either entrained to 24 h LD or free-running, as
in the present study under LD:1,1.
Is the hypothesis correct in the case of E. gracilis grown
photoautotrophically under LD:1,1? It is wrong as a whole, but
it is correct in that the photoinduction of the commitment in E.
gracilis is regulated by CRdusk (Hagiwara et al. 2002), such that
a 6 h light pulse near subjective dusk most profoundly induces
the commitment of G1, S and G2 cells to cell cycle transition,
whereas a light pulse near subjective dawn is ineffective. Nevertheless, the hypothesis is wrong, in that it supposes a constant duration post-commitment, from commitment to the
beginning of mitosis or cell division, because we have shown
that it varies depending on the circadian phase when commitment was achieved (Fig. 4).
Is circadian gating of S-to-G2 transitions responsible?
Alternatively, one might argue that the circadian gating of G2to-M transitions between CT08 and CT20 results from the
hypothetical constant length, ∼20 h, of the G2 phase, because
G2 cells were produced only between CT12 and CT24 (Fig.
2B). This hypothesis is the same as the above hypothesis in its
basic concept. According to the difference in the circadian
rhythm that is assumed to be responsible for the circadian gating of the G2-to-M transition, there is a difference in the proc-
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Circadian gating of G2-to-M transition in Euglena
ess time interval, which is thought to be kept constant by a
developmental mechanism.
Whereas the hypothesis in the previous section assumes a
constant duration of the post-commitment phase of G2 cells,
this hypothesis makes the assumption for the entire G2 phase.
Since the latter should be longer than the former by the duration of the pre-commitment phase of G2 cells, i.e. the time
needed for commitment by G2 cells, the latter is most likely to
be more variable than the former. Therefore, this hypothesis is
more unreasonable than the previous hypothesis.
Finally, note that neither hypothesis can be reconciled
with the fact that the timing of the G2-to-M transition was
unchanged by differences in cell cycle speed, because a developmental mechanism is not usually responsible for maintaining a developmental process of relatively long duration. Rather,
such a mechanism either maintains a relatively short, and perhaps minimal, duration of cell cycle progression or it lengthens the minimum duration of a cell cycle phase. The former is
exemplified by the constant lengths of S (4.8 ± 0.6 h) and G2 +
M (3.3 ± 0.4 h) phase under LL regardless of cell cycle speed
(Hagiwara et al. 2001) and the latter by the lengthening of the
minimum length of ∼7 h of G1 phase with decreasing light
intensities under LL, which slows cell cycle progression (Hagiwara et al. 2001).
Materials and Methods
The algal flagellate E. gracilis Klebs (Z) was cultured photoautotrophically and axenically as in Hagiwara et al. (Hagiwara et al.
2001, Hagiwara et al. 2002). The cultures were magnetically stirred
and aerated at 25°C, and illuminated under LL from an array of daylight-white fluorescent lamps (Toshiba, Mellow White) at an intensity
ranging from 4 to 9.5 klux (56–133 µmol m–2 s–1). When they were in
log-linear growth mode, they were transferred to LD:1,1. A 7 ml
aliquot of the culture was automatically collected and fixed using
0.5 ml of 20% neutralized formalin containing 5% KCl every 2 h, and
cell numbers were counted with a Coulter Counter.
Kinetic behavior of cell cycle progression was analyzed by DNA
FCM together with microscopic observation of mitotic cells as in
Hagiwara et al. (Hagiwara et al. 2001, Hagiwara et al. 2002). Briefly,
the cells, sampled at 2 h intervals, were stained with propidium iodide
and then subjected to DNA FCM or microscopic observation. The
percentages of cells in G1, S and G2 + M phase were calculated applying a fitting equation of a diploid population supplied by ModFit LT of
Verity SoftwareHouse, Inc. (Topsham, ME, USA) to the FCM data.
The percentages of cells in mitosis were counted by microscopic
observation.The average GT of circadian cultures was estimated by the
formula, GT = (a circadian period, 26 h)×ln (2)/ln (a factorial increase
of cell number during a circadian cycle). The calculation of GT for
log-linear cultures was described in Hagiwara et al. (2001).
Acknowledgment
We are grateful to Dr. Carl H. Johnson of Vanderbilt University
(Nashville, TN, USA) for critical comments and grammatical correction of the first draft of this manuscript. We also thank Dr. Leland N.
Edmunds, Jr. of SUNY (Stony Brook, NY. USA) for his critical comments on the first draft. Y.Z. is the recipient of the JSPS Invitation Fellowship of Japan Society for the Promotion of Science in 1997 (No. S97155), and of the Invitation Fellowship of OASERD (Obihiro Asia
and the Pacific Seminar on Education for Rural Development) in
1998–2001.
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(Received September 8, 2004; Accepted March 29, 2005)