Key words: apical dominance, bud dormancy, cell cycle arrest, cell quiescence, cell profileration, ..... the bud mass, it is not surprising that we would detect only ...
Plant Molecular Biology 29: 255-265, 1995. © 1995 Kluwer Academic Publishers. Printed in Belgium.
255
Cell cycle regulation during growth-dormancy cycles in pea axUlary buds Michelle L. D e v i t t and Joel P. Stafstrom * Plant Molecular Biology Center and Department of Biological Sciences, Northern Illinois University, DeKalb, IL 60115, USA (* author for correspondence) Received 16 January 1995; accepted in revised form 12 July 1995
Key words: apical dominance, bud dormancy, cell cycle arrest, cell quiescence, cell profileration, Pisum sativum
Abstract
Accumulation patterns of mRNAs corresponding to histones H2A and H4, ribosomal protein genes rpL27 and rpL34, MAP kinase, cdc2 kinase and cyclin B were analyzed during growth-dormancy cycles in pea (Pisum sativum cv. Alaska) axillary buds. The level of each of these mRNAs was low in dormant buds on intact plants, increased when buds were stimulated to grow by decapitating the terminal bud, decreased when buds ceased growing and became dormant, and then increased when buds began to grow again. Flow cytometry was used to determine nuclear D N A content during these developmental transitions. Dormant buds contain G 1 and G2 nuclei (about 3:1 ratio), but only low levels of S phase nuclei. It is hypothesized that cells in dormant buds are arrested at three points in the cell cycle, in midG1, at the Gt/S boundary and near the S/G2 boundary. Based on the accumulation of histone H2A and H4 mRNAs, which are markers for S phase, cells arrested at the G~/S boundary enter S within one hour of decapitation. The presence of a cell population arrested in mid-G~ is indicated by a second peak of histone m R N A accumulation 6 h after the first peak. Based on the accumulation of cyclin B mRNA, a marker for late G2 and mitosis, cells arrested at Gx/S begin to divide between 12 and 18 h after decapitation. A small increase in the level of cyclin B m R N A at 6 h after decapitation may represent mitosis of the cells that had been arrested near the S/G2 boundary. Accumulation of MAP kinase, cdc2 kinase, rpL27 and rpL34 mRNAs are correlated with cell proliferation but not with a particular phase of the cell cycle.
Introduction
Axillary buds of Alaska pea plants can be stimulated to undergo more than one complete
growth-dormancy cycle ( d o r m a n t ~ g r o w i n g ~ dormant~growing) over a period of six days [33, 36]. There are four dormant buds at the second node of intact pea plants, all of which begin
The nucleotide sequence data reported will appear in the EMBL, GenBank and DDBJ Nucleotide Sequence Databases under the accession numbers U10041 (His2A), U10042 (His4) and U10047 (rpL34).
256 to grow 8 h after decapitation. After two to three days, rapid growth by the largest bud inhibits further growth of the smaller buds; the small buds become fully dormant four to five days after decapitation. Small buds resume growing within one day of removing the large bud. Expression of ribosomal protein gene L27 (rpL27) has been used as a growth-specific marker to analyze growthdormancy cycles in pea buds [36]. In situ hybridization experiments showed that rpL27 m R N A levels increase in all regions of large and small buds within 1 h of decapitation. It appears that regulatory signals from the plant are transported very rapidly into the buds and that all bud cells are competent to respond to these signals. In small buds, rpL27 m R N A levels decline when they become dormant again and then increase after they are stimulated to resume growing. The rapidity and ease with which growth states can be interconverted make pea buds an ideal system in which to analyze cellular and molecular events associated with cell cycle control. The proliferating and quiescent cell cycle states, as well as particular phases of the cell cycle, are characterized by specific patterns of gene expression. For example, histone proteins and mRNAs accumulate to high levels during S phase in all eukaryotes [26, 30, 38]. Other proteins regulate passage through cell cycle 'checkpoints', that is, they assure that one phase is completed before the next can begin [27]. Maturation-promoting factor (MPF) is a complex of cyclin B and cdc2 kinase, a cyclin-dependent kinase (CDK); M P F regulates entry into mitosis in all eukaryotes [ 15, 28]. Passage through other cell cycle checkpoints is probably regulated by other cyclin/CDK complexes in association with several additional factors [ 14]. In plants, m R N A for cdc2 kinase is present in proliferating cells and in cells that are 'competent to divide' [12,24]; and cyclinB m R N A accumulates predominantly during late G2 and mitosis [ 13]. MAP kinase is a component of many intracellular signal transduction pathways, including pathways involved in cell cycle regulation [17]. In vertebrates, MAP kinase m R N A levels are similar in quiescent and proliferating cells [2, 11 ]. In contrast, accumulation of
plant MAP kinase m R N A occurs predominantly in proliferating cells [18, 34]. MAP kinase enzyme activity in animals, fungi and plants is regulated by phosphorylation of specific amino acids [5,17]. In this report we describe the cloning of histone H2A (His2A), histone H4 (His4) and ribosomal protein L34 (rpL34) cDNAs. Accumulation of mRNAs corresponding to these clones, as well as rpL27, MAP kinase [34], cdc2 kinase [6] and cyclin B [ 16] clones, were analyzed in developing pea buds. The D N A content of bud nuclei was determined using flow cytometry. Data from both types of experiments was used to analyze cell cycle kinetics during reversible growth-dormancy cycles. We demonstrate close correlations between the dormant state of bud development and cell quiescence, and between bud growth and cell proliferation. In addition, we show that cell cycle in dormant buds is arrested in mid-G1, and near the G1/S and S/G2 boundaries. Materials and methods
Plant material Peas (Pisum sativum L. cv. Alaska) were grown in a growth chamber maintained at ca. 22 ° C under a 16 h light/8 h dark photoperiod. Plants were 7 to 8 days old at the beginning of an experiment. The two largest buds at node 2, which are referred to as 'large' and 'small' buds, were analyzed. Large and small buds on intact 7-day old plants measured ca. 1.5 mm and 1.0 mm, respectively; two other buds at node 2 are much smaller [35, 36]. Buds were stimulated to grow by decapitating the main shoot directly above node 2. Large buds were collected at several time points during the first 24 h after decapitation and small buds were collected dally through six days after decapitation. By five days after decapitation, rapidly growing large buds inhibited the further growth of small buds, which became dormant again. Dormant small buds were stimulated to grow by removing the large bud. Small buds then were collected at several time points during the ensuing 24 h.
257
Isolation of growing bud-specific cDNA clones Two c D N A libraries were made previously using A + R N A isolated from growing buds (i.e., 24 h after decapitation). First, a 2 - G E M 4 library was screened by differential hybridization to isolate clones that were up-regulated in growing buds [36]. Three growing bud clones were sequenced and found to contain incomplete coding sequences. Full-length c D N A clones were isolated from the second c D N A library, which had been prepared in the p S P O R T - I plasmid vector [34]. Complete sequences of these clones were determined by dideoxy sequencing of several subclones (Sequenase kit, US Biochemical).
RNA gel blotting Total bud R N A (10 #g/lane) was separated by denaturing formaldehyde gel electrophoresis and blotted onto nylon membranes [36]. Randomprimed 32p-labeled probes were prepared from gel-purified c D N A inserts using the DecaPrime kit (Ambion). Blots were prehybridized in 50~o formamide, 5 x Denhardt's reagent, 0 . 5 ~ SDS, 5 x SSPE and 100 #g/ml salmon sperm D N A for 2 h at 42 ° C, and hybridized overnight under the same conditions. Blots were washed in 0.2 x S SC and 0 . 1 ~ SDS at 55 °C and exposed to X-ray film. Blots were stripped and reprobed up to two times. The c D N A clones analyzed were: rpL34, His2A and His4 (this report); rpL27 [36]; M A P kinase [34]; and cyclin B and cdc2 kinase (clones pcycPsl and pcdkPsl, respectively, were obtained from T. Jacobs [6, 16]). A r R N A clone was used as a loading control (the p H A 2 was obtained from N. Polans).
Flow cytometry Nuclei were prepared according to a method provided by T. Jacobs (personal communication). Axillary buds (40 per time point) were fixed on ice for 30 min in 4~o formaldehyde in wash buffer ( 1 0 m M Tris-HC1 pH 7.4, 1 0 m M EDTA,
100 mM NaC1). The buds were rinsed four times (3 min each) in wash buffer. Buds were transferred to a microfuge tube containing nuclear isolation buffer (wash buffer plus 0.1 ~o Triton-X 100) and crushed with a glass pestle. Nuclei were filtered through a 20/~m nylon filter to remove debris and then pelleted at 300 x g (1800 rpm in an Eppendorf microfuge) for 5 min at 4 ° C. The final pellet was resuspended in 70}'0 EtOH and stored at 4 °C for up to 3 weeks. Nuclei were stained with phosphate-buffered DAPI (4,6diamidino-2-phenylindol). For each sample, the D N A content of 10000 nuclei was determined using a P A R T E C - P A S I I I flow cytometer. Individual nuclei were assigned to G1, S or G2 populations according to the algorithm of Dean and Jett [ 3 ]. These experiments were performed in the laboratory of Dr David Grdina (Argonne National Laboratory) with the assistance of Dr Jeffrey Murley.
Results
Isolation and sequence analysis of growth-associated cDNA clones A lambda growing bud c D N A library was differentially screened for clones with increased expression in growing buds. The D N A sequences of three growth-associated clones were similar to histone H2A, histone H4 and ribosomal protein L34 genes from other species. None of these c D N A s contained a complete coding sequence. A second growing bud c D N A library, which was prepared in a plasmid vector, was used to isolate clones containing complete coding sequences. The amino acid sequences of the His2A, His4 and rpL34 clones were compared to similar sequences from the G e n B a n k / E M B L databases (Fig. 1). The deduced amino acid sequence of our His2A clone is identical to another pea His2A clone at 141 of 149 residues; our clone encodes one fewer residue (Fig. 1A). The amino acid sequence of our pea His4 clone differs from a tomato clone at just one residue (Fig. 1B) and is identical to a maize clone (not shown). Sequences
258 A.
HiS2A
Ps2 Psl
MDASTKVKKGAGGRK GGGPRKKAVTRSVRA GLQFPVGRIGRFLKK GRYAQRVGTGAPVYL AAVLEYLAAEVLELA ...... T ............... ~ ............... ~ ....................................
PS2 PSl
GNAARDNKKNRISPR HLLLAVRNDVELGKL LAGVTIAYGGVLPNI NPVLLPKRTESAA-S ........................ E ............................ K.~..A.
B. PS
MSGRGKGGKGLGKGG
.......................
Ps
HARRKTVTAMDVVYA
Le
............................
AKRHRKVLRDNIQGI
TKPAIRRLARRGGVK
RISGLIYEETRGVLK
IFLENVIRDAVTYTE
: .............................................
~ .....
LKRQGRTLYGFGG
75 75 103 103
rpL34
Ps
MVQRLTYRRRHSYAT
Nt
........ K .....................
Ps Nt
149 150
His4
Le
C.
APKSPSKAKKTPKKA T ..............
75 75
KSNQHRVVKTPGGKL
VYQTTKKRASGPKCP
VTGKRIQGIPHLRPT
EYKRSRLSRNRRTD-N
~--~ .........................................
RAYGGVLSGGAVRER IIRAFLVEEQKIVKK VLKIQKTKEKQAAKN ......... S .......................... A...L...~
75 75 120 120
Fig. 1. Comparison of deduced amino acid sequences of pea c D N A clones with selected homologous sequences. A. Alignment of two pea histone H2A clones: Ps2, this report; Psl [19]. B. Alignment of histone H4 clones from pea (Ps) and tomato (Le [1]). C. Alignment of ribosomal protein L34 clones from pea (Ps) and tobacco (Nt [9]). Dots indicate identity; dashes indicate gaps that were inserted to maximize alignment; underlined residues are considered to be similar.
ofhistone H4 proteins are known to be very highly conserved [45]. Tobacco is the only other plant from which a rpL34 c D N A has been isolated [9]. The pea and tobacco rpL34 clones are identical at 113 of 120 residues.
Patterns of mRNA accumulation during growthdormancy cycles As outlined in the Introduction, node 2 small buds can be stimulated to undergo more than one complete growth-dormancy cycle over a six day period. The expression pattern of rpL27 was analyzed previously [36] and is included here for comparison (Fig. 2). The relative abundance of rpL27 m R N A was very low in dormant buds on intact plants, it increased significantly in growing buds two days after decapitation, and then it declined when buds became dormant again between three and five days after decapitation. If the large bud was removed five days after decapitating the main shoot, small buds resumed growing and again contained high levels ofrpL27 m R N A (5 + 1 days). Growing 5 + 1-day buds probably contained more rpL27 m R N A than 'growing' 2-day buds because the latter already had begun to reenter dormancy (compare with growing 24 h buds
in Fig. 3). Accumulation patterns for m R N A s corresponding to the three clones described in this report (His2A, His4 and rpL34) and for m R N A s corresponding to the M A P kinase, cdc2 kinase and cyclin B clones were quite similar to the rpL27 pattern, but a few differences can be noted. For example, all of these m R N A s were relatively more abundant than rpL27 at three days after decapitation, that is, during the early phase of the growth-to-dormancy transition. Among the clones studied, dormant buds contained relatively higher levels of cdc2 kinase m R N A than of the other messages. Large axillary buds were used to examine the dormancy-to-growth transition during the release of buds from apical dominance. In dormant buds on intact plants, the steady-state level of all seven m R N A s analyzed was very low or undetectable (Fig. 3). The levels of cdc2 kinase, M A P kinase, rpL27 and rpL34 m R N A s increased within one hour of decapitation, reached a maximum level by three to six hours, and persisted at these high levels through 24 h. Cyclin B m R N A showed a small increase at six hours, was reduced to the dormancy level between 9 and 12 h and then reached its highest level at 18 and 24 h. Levels of the two histone m R N A s increased to relatively high levels within one hour of decapitation,
259
Fig. 2. RNA gel blot analysis of growth-dormancy cycles in
node 2 small buds. A 10 ~tg portion of total RNA was separated by denaturing formaldehyde gel electrophoresis, transferred to a nylon membrane and hybridized with a 32p-labeled probe. Numbers above each lane indicate days after decapitating the main stem; 5 + 1 days indicates removal of the large bud at 5 days and analysis of small buds 1 day later. Developmental stages are: D, dormant; G to D, growth-to-dormancy transition; G, growing. Messenger RNAs analyzed were: rpL27, rpL34, His2A, His4, MAP kinase, cdc2 kinase and cyclin B. A rRNA probe was used as a loading control. reached their highest levels at 6 h, declined at 9 h, and increased to and remained at high levels at 12 through 24 h.
Flow cytometry
The D N A content o f bud nuclei was determined by flow cytometry. D o r m a n t large buds contained nuclei with the G1 (2C) and Gz (4C) content o f D N A in a ratio o f about 3:1 (Fig. 4A). The computer algorithm used to assign peaks indicated that about 10~o o f the nuclei contained an inter-
Fig. 3. RNA gel blot analysis of the dormancy-to-growth transition in node 2 large buds. Methods are as for Fig. 2. Numbers above each lane indicate hours after decapitation. Developmental stages are: D, dormant; D to G, dormancyto-growth transition; G, growing. Messenger RNAs analyzed were: rpL27, rpL34, His2A, His4, MAP kinase, cdc2 kinase and cyclin B. A rRNA probe was used as a loading control.
mediate level o f D N A , which would be indicative o f S phase. Since only very low levels o f histone m R N A s could be detected in d o r m a n t buds (Figs. 2 and 3), a measured value o f about 10~o "S phase' nuclei probably should be interpreted as equivalent to zero actual S phase cells. One day after decapitation, S phase nuclei represented a significant proportion o f the total (Fig. 4B). Five days after decapitation, small buds had b e c o m e d o r m a n t again. The flow cytometry profile for these buds (Fig. 4C) was quite similar to that o f d o r m a n t large buds. One day after removing the large bud, small buds again contained a high proportion o f S phase nuclei (Fig. 4D). The percentage o f nuclei in each phase of the cell cycle was determined for small buds during more than one
260
B.
A,
D.
C.
[] G1
o>, E
[]S [] G2
3
U-
2
i
. ,,
,,
°
.,.,,,
.2~ 0
32
64
96
0
32
64
:
~ ..,,~
96
32
64
0
96
32
64
96
DNA Content Fig. 4. Flow cytometry analysis of nuclear DNA content of axillary buds. Nuclei were isolated from: A. Dormant large buds (from intact plants). B. Growing large buds (24 h after decapitation). C. Dormant small buds (5 days after decapitation). D. Growing small buds (large buds were removed at 5 days and small buds were collected 1 day later).
c o m p l e t e g r o w t h - d o r m a n c y cycle (Fig. 5). T h e p r o p o r t i o n o f S p h a s e nuclei w a s at the b a s a l level o f a b o u t 10 ~ in d o r m a n t b u d s (0 days), i n c r e a s e d to a b o u t 30~o in g r o w i n g b u d s (1 a n d 2 days), declined during the g r o w t h - t o - d o r m a n c y transition (3 to 5 days), a n d t h e n i n c r e a s e d after these b u d s w e r e stimulated to r e s u m e g r o w i n g (5 + 1 days). A t e a c h stage o f d e v e l o p m e n t , an increase in the p r o p o r t i o n o f S nuclei o c c u r s in parallel with a d e c r e a s e o f G1 nuclei, a n d vice versa. G 2
80
60
"5
...El ,EY" El,/ \
q
40
...... [] ........ G~
~ 40-
.... o ....
.......13
G2
20 0
t
'\
-I ........D ........ G1
i
...0 .... ~ ........
/
D'...., D
40-
't]
S
20o,O..O.-.-o ....
20- d
t
j~...D..,...o......o,,,,.q\ B.
60 "/
Z
---0---
D
-...-. - o[]- -........ GI S
"~
60-
"b".-......d/
A. [
~''°'"'~'.-.
"~ Z
80
80
~dzq~q. _
v
0
j
i
0
3
6 Hours
i
|
i
t
i
i
i
0
1
2
3
4
5
5+1
Days After D e c a p i t a t i o n
Fig. 5. Cell cycle kinetics in small buds during more than one
complete growth-dormancy cycle. Cell cycle phases were determined by flow cytometry analysis of bud nuclei. Main shoots were decapitated on day 0 and large buds were removed at 5 days, which promoted cell proliferation in small buds at 5 + 1 days.
...
9
,,,'"
12
",,,
15
...,o..,.o
1 8 21
24
After Decapitation
Fig. 6. Cell cycle kinetics in axillary buds
during the dormancy-to-growth transition. Cell cycle phases were determined by flow cytometry analysis of bud nuclei. A. Large buds; hours after decapitating the main shoot. B. Small buds; 5 days after decapitating the main shoot, dormant small buds were stimulated to grow by removing the large bud at time 0. Arrows indicate time points prior to increases in the proportions o f S phase nuclei.
261 nuclei constitute about 15 to 20~o of the total at each stage. Nuclear D N A content was determined for large and small buds during the first 24 h after being released from apical dominance, that is, during the dormancy-to-growth transition (Fig. 6). If the 10~o 'S phase' nuclei in dormant buds were divided equally among the G~ and G2 populations, the G I : G 2 ratio would be between 3:1 (large buds) and 4:1 (small buds). Cell cycle kinetics are slightly different in large and small buds. In large buds, the proportion of S phase nuclei increased after 6 and 15 h and represented 3 0 ~ of the total at 24 h (Fig. 6A, arrows). In small buds, a single increase occurred after 9 h; after 24 h S phase nuclei represented 38~o of the total (Fig. 6B, arrow). Based on patterns of histone m R N A accumulation, it is likely that cells actually enter S phase sooner than indicated by these experiments (see Discussion). At each stage buds contained about 15 to 20 ~o G2 nuclei.
Discussion
Dormant axillary buds of pea can be stimulated to grow, to become dormant again and then to resume growing, all within six days [36]. In the present study, we analyzed the cell cycle during these growth-dormancy cycles. Flow cytometry was used to determine the proportions of cells in each phase of the cell cycle at each stage of development. A limitation of this technique is that it cannot indicate where within a cell cycle phase a cell is located. For example, a cell with a 2C or G~ D N A content may have just completed mitosis, it may be ready to enter S phase or it may be quiescent. Flow cytometry is a much more powerful and informative technique when the expression of molecular markers are examined in parallel [10]. In this report, seven mRNAs were analyzed whose patterns of accumulation are correlated with bud growth and cell proliferation. c D N A clones corresponding to histone H2A, histone H4 and ribosomal protein L34 genes were isolated and characterized during these studies (Fig. 1). The ensuing discussion is focused on two
general questions. First, to what extent are two cellular states (proliferation and quiescence) related to two organ-level states (growth and dormancy)? And second, where in the cell cycle are dormant bud cells arrested and what molecular changes occur when they are stimulated to proliferate?
Cell proliferation and quiescence during growthdormancy cycles We showed previously that accumulation of rpL27 m R N A is tightly linked to bud growth [36], and here we show that the pattern of rpL34 m R N A accumulation is quite similar (Fig. 2). In general, cytoplasmic ribosomal protein genes are regulated coordinately [23], although variable patterns of expression have been noted (e.g., [9]). Patterns of accumulation for His2A, His4, MAP kinase, cdc2 kinase and cyclin B mRNAs also are quite similar to the rpL27 pattern (Fig. 2). Histone mRNAs accumulate predominantly during S phase in all eukaryotes that have been studied [26, 30, 38]. Typically, S phase or replicationdependent accumulation is about 20-fold higher than accumulation during other phases of the cell cycle. Replication-independent accumulation of histone mRNAs may be driven by the same promoters that are responsible for replicationdependent transcription [20, 39]. Based on the absence of other proliferation-specific markers in dormant pea buds, the very low levels of histone mRNAs in these buds probably are due to replication-independent accumulation (Fig. 2). In plants, cyclin B m R N A accumulation is correlated with late G2 and M phases [13]. Since cells rarely arrest during S or M phases [40, 43], histone and cyclin B mRNAs also are excellent markers for cell proliferation. Accumulation of MAP kinase and cdc2 kinase mRNAs are closely associated with proliferating cells but not with particular phases of the cell cycle (see below). Differentiated plant cells contain little cdc2 kinase m R N A whereas quiescent cells that are 'competent to divide' contain higher levels
262 [ 12, 24]. Quiescent cells in dormant pea buds also contain some cdc2 m R N A but considerably less than proliferating cells (Figs. 2 and 3). Based on their nuclear D N A content, the proportion of S phase (proliferating) cells is much lower in dormant buds than in growing buds (Figs. 4, 5 and 6). Together these results indicate that bud dormancy in pea is closely correlated with cellular quiescence and bud growth is correlated with cell proliferation. The proliferation-to-quiescence transition, which occurs in small buds when they re-enter the dormant state (Figs. 2 and 5), has not been studied in any plant system. The quiescent state might be induced by reduced activity of positive cell cycle regulators such as cyclin/CDK complexes or MAP kinase, by increased activity of as yet unidentified tumor suppressor-like proteins, or both. The speed and ease with which the quiescent and proliferating states can be interconverted will allow such questions to be addressed using the pea bud system. Recently, a cyclin promoter:GUS fusion was shown to be expressed in dormant axillary buds of transgenic Arabidopsis plants (Fig. 1H in [8]), a finding that might appear to contradict our resuits (Figs. 2 and 3). Based on the relative levels of G U S staining in Arabidopsis root apices and dormant buds, the rate of cell division in the buds would appear to be much lower. In addition, G U S staining in these buds appears to be confined to the shoot apical meristem and perhaps one or two of the youngest leaf primordia; the level of staining in older primordia is considerably lower. Based on histological studies of dormant pea buds, the mitotic index in the apical meristem and the two youngest leaf primordia is relatively high; in contrast, virtually no mitotic figures were found elsewhere in the bud (J. P. Stafstrom, in preparation). Thus, cell division in the apices of dormant buds of both species may be quite similar. Since the areas of dormant pea buds containing dividing cells represents less than 1 ~o of the bud mass, it is not surprising that we would detect only low levels of cyclin m R N A in these buds.
Cell cycle arrest in dormant axillary buds
Sucrose-starved root tips of pea and several other species, as well as cells within dormant seeds, contain cells arrested in G 1 and G2 [40]. The 'principal control point model' of Van't Hof and Kovacs [43] hypothesized that passage through a control point (or checkpoint) requires the presence of specific regulatory proteins. It is now known that activated M P F and other cyclin/CDK complexes play key roles in this regulation (reviewed in [4, 14, 15, 28, 29]). Cells in dormant pea buds also are arrested in G1 and G2 (Figs. 4A and 4C). We used the kinetics of histone and cyclin m R N A accumulation (markers for S phase and late G2/M, respectively) to determine where within these phases cells are arrested. While histone mRNAs accumulate predominantly in S phase, a few slight exceptions to this general rule should be noted. For example, Tanimoto and coworkers [38] demonstrated that histone m R N A accumulation in pea root apices begins before the onset of S phase and ceases before D N A synthesis has been completed. S phase in these cells takes about 6 h [43]. Based on this fact and data of Tanimoto et al. [38], we calculate that histone m R N A accumulation in pea root cells begins about 1 h before the S phase begins and ceases 1/2 h before the end of S phase. Histone m R N A accumulation and S phase can be uncoupled using the D N A polymerase inhibitor aphidicolin [31 ], but this observation has no direct bearing on endogenous control of the cell cycle as occurs in developing pea buds. In pea buds, histone m R N A accumulation begins within one hour of decapitation, whereas the first indication of an increase in D N A content as determined by flow cytometry does not occur until 7 h (compare Figs. 3 and 6A). The 6 h delay seen in the flow cytometry data may be due to the fact that D N A content of cells must be considerably greater than 2C before the computer scores them as S phase cells (notice the overlaps in the computer-assigned peaks in Fig. 4). Thus, we consider histone m R N A accumulation to be a better method for identifying S phase cells and for analyzing cell cycle kinetics. Histone mRNAs begin to accumulate in axil-
263 lary buds within one hour of decapitation, reach a maximal level at six hours, decline at nine hours, and then increase to and remain at a high level between 12 and 24 h (Fig. 3). This pattern of may be due to two (or more) populations of cells arrested at different positions in G1. Cells that began to accumulate histone mRNAs within 1 h of decapitation (Fig. 3) must have been arrested near the G1/S boundary. The decline in histone mRNAs at 9 h might occur as these cells completed S and entered G2. A D N A synthesis phase lasting 6 to 9 h is consistent with previous studies on cell cycle kinetics in pea shoot apices [21, 22]. A second population of dormant bud cells appears to be arrested in mid-Gp Entry of these cells into S would account for a second increase in histone gene expression beginning at 12 h after decapitation (Fig. 3). The large increase in cyclin m R N A accumulation seen at 18 and 24 h may represent mitosis of the cells that had been arrested in G1. A small, transient increase in the level of cyclin m R N A 6 h after decapitation may represent mitosis of cells that had been arrested in G2 (Fig. 3). Cytological studies confirm that the earliest increase in the number of mitotic figures occurs 6 h after decapitation (J. P-. Stafstrom, in preparation). The G2 population may be arrested near the G2/M boundary, but experience a lag of about 6 h before dividing. Alternatively, these cells may be arrested nearer to the S/G2 boundary and, upon being stimulated to proliferate, traverse G2 before dividing. The latter interpretation is consistent with a G2 duration of about 5.5 h (see below). The duration of each phase of the cell cycle can be determined if (1)the relative duration of each phase is known and (2)the absolute duration of one or more phases is known. With regard to the first requirement, an unsynchronized cell population in growing buds comprises about 50~o G1 cells, 30~o S cells and 20~o G2 cells (Fig. 5, 2 days after decapitation). The approximate duration of each phase was calculated as follows. Based on accumulation of cyclin mRNA, cells that had been arrested at G~/S begin to divide between 12 and 18 h after decapitation (Fig. 3). S + G2, the time until the onset of mitosis, repre-
sents 50~o of the cell cycle, so the duration of a complete cell cycle would be 24 to 36 h. Since cyclin expression had reached its maximal level by 18 h, it is likely that cells began to enter mitosis at an earlier time (e.g., 15 h). Therefore, we adopt the working hypothesis that cell generation time in growing pea buds is about 30 h. Mitosis is presumed to last 1 h [22]. If we assume a cell generation time of 30 h (and subtract 1/2 h from both G1 and G2 to account cells in mitosis), the approximate duration of each cell cycle phase is: M, 1 h; G1, 14.5 h; S, 9 h; and G2, 5.5 h. Using different methods, the average cell generation time for the pea shoot apex was found to be 26 to 30 h and the durations of all phases were similar to our estimates [21, 22]. The G1/S and G2/M boundaries are considered to be the primary cell cycle checkpoints [29]. However, cell cycle arrest at several other checkpoints is well documented. Our results suggest that cells in dormant buds are arrested in G2, but perhaps earlier than G2/M, and at two or more points in G1 (see above). In sucrose-starved pea root apices, cells that will give rise to vascular parenchyma are arrested in late S or early G2 [41, 44] and other cell populations are arrested at G1/S and at an earlier stage of G1 [42, 43]. The latter population might be equivalent to the mid-G1 population described here. Sucrosestarved root apices of other species have GI:G2 ratios ranging from 3:1 to 1:3, presumably because different combinations of checkpoints are utilized [40]. Each cell cycle checkpoint is probably controlled, at least in part, by a cyclin/CDK complex [ 14]. Humans contain at least 11 cyclins and a similar number of CDKs, which could easily account for the several arrest points that have been documented [ 14, 25]. Arabidopsis and maize each contain at least three families ofcyclin genes, which are differentially expressed according to cell type, cell cycle phase, or both [7, 32, 37]. Thus, plants contain the regulatory machinery to account for several cell cycle checkpoints. Ongoing experiments in our laboratory should help to clarify aspects of cell cycle arrest in dormant buds and cell proliferation in growing buds. Further studies of node 2 small buds are of par-
264 ticular interest: cells may be arrested at only one point in G 1 (compare Figs. 6A and 6B); and 'small' buds that have re-entered the dormant state five days after decapitation are actually about three times longer and ten times more massive than 'large' buds on intact plants [34], which will make it easier to collect large quantities of buds at each developmental stage. Results from preliminary experiments indicate that cells arrested in G1 and G2 are distributed equivalently in all regions of the bud and in all cell types (J. L-. Stafstrom, in preparation). The spatial distribution of these cell populations is being studied further by in situ hybridization analysis of histone and cyclin mRNAs.
8.
9.
10.
11.
12.
Acknowledgements We thank Dr Thomas Jacobs for providing the cdc2 kinase and cyclin c D N A clones and for advice on flow cytometry, Drs Jeffrey Murley and David Grdina for help in performing the flow cytometry experiments, and Dr Neil Polans for the pHA2 clone. This work was supported by the Plant Molecular Biology Center, Northern Illinois University.
13.
14. 15. 16.
References 1. Brandstadter J, Rossbach C, Theres K: The pattern of histone H4 expression in the tomato shoot apex changes during development. Planta 192:69-74 (1994). 2. Chen R-H, Sarnecki C, Blenis J: Nuclear localization and regulation of erk- and rsk-encoded protein kinases. Mol Cell Biol 12:915-927 (1992). 3. Dean PN, Jett JH: Mathematical analysis of D N A distributions derived from flow microfluorometry. J Cell Biol 60:523-527 (1974). 4. Doerner PW: Cell cycle regulation in plants. Plant Physiol 106:823-827 (1994). 5. Duerr B, Gawienowski M, Ropp T, Jacobs T: MsERKI: a mitogen-activated protein kinase from a flowering plant. Plant Cell 5:87-96 (1993). 6. Feiler HS, Jacobs TW: Cloning of the pea cdc2 homologue by efficient immunological screening of PCR products. Plant Mol Biol 17:321-333 (1991). 7. Ferriera PC, HemerlyAS, de Almeida EnglerJ, BergouniouxC, Burssens S, Van Montagu M, EnglerG,
17.
18.
19.
20.
21. 22.
Inz6 D: Three discrete classes of Arabidopsis cyclins are expressed during different intervals of the cell cycle. Proc Natl Acad Sci USA 91:11313-11317 (1994). Ferriera PC, Hemerly AS, de Almeida EnglerJ, Van Montagu M, Engler G, Inz6 D: Developmental expression of the Arabidopsis cyclin gene cyclAt. Plant Cell 6: 1763-1774 (1994). Gao J, Kim S-R, Chung Y-Y, Lee JM, An G: Developmental and environmental regulation of two ribosomal protein genes in tobacco. Plant Mol Biol 25:761-770 (1994). Georgieva El, Lrpez-Rodas G, Hittmair A, Feichtinger H, Brosch G, Loidl P: Maize embryo germination. I. Cell cycle analysis. Planta 192:118-124 (1994). Gotoh Y, Moriyama K, Matsuda S, Okumura E, Kishimoto T, Kawasaki H, Suzuki K, Yahara I, Sakai H, Nishida E: Xenopus M phase MAP kinase: isolation of its cDNA and activation by MPF. EMBO J 10:2661-2668 (1991). Hemerly AS, Ferreira P, de Almeida Engler J, Van Montagu M, Engler G, Inz6 D: cdc2a expression in Arabidopsis is linked with competence for cell division. Plant Cell 5:1711-1723 (1993). Hirt H, Mink M, Pfosser M, BOgre L, Gyorgyey J, Jonak C, Gartner A, Dudits D, Heberle-Bors E: Alfalfa cyclins: differential expression during the cell cycle and in plant organs. Plant Cell 4:1531-1538 (1992). Hunter T, Pines J: Cyclins and cancer II: cyclin D and CDK inhibitors come of age. Cell 79:573-582 (1994). Jacobs TW: Control of the cell cycle. Devel Biol 153: 1-15 (1992). Jacobs TW, Prewett ME, Duerr BK, Feller HS, Dunphy J, Chen H, Poole J: Protein kinases in cell division control. In: Plant Protein Phosphorylation, Plant Protein Kinases and Phosphatases and Plant G-Proteins, 12th Plant Biochemistry, Molecular Biology and Physiology Symposium, pp. 67-68. University of Missouri, Columbia, MO (1993). Jonak C, Heberle-Bors E, Hirt H: MAP kinases: universal multi-purpose signaling tools. Plant Mol Bio124: 407416 (1994). Jonak C, Pay A, Brgre L, Hirt H, Heberle-Bors E: The plant homologue of MAP kinase is expressed in a cell cycle-dependent and organ-specific manner. Plant J 3: 611-617 (1993). Koning AJ, Tanimoto EY, Kiehne K, Rost T, Comal L: Cell specific expression of plant histone H2A genes. Plant Cell 3:657-665 (1991). Lepetit M, Ehling M, Chaubet N, Gigot C: A plant histone gene promoter can direct both replication-dependent and -independent gene expression in transgenic plants. Mol Gen Genet 231:276-285 (1992). Lyndon RF: Rates of cell division in the shoot apical meristem of Pisum Ann Bot 34:1-17 (1970). Lyndon RF: The cell cycle in the shoot apex. In: Balls M, Billett FS (eds) The Cell Cycle in Development and Dif-
265
23. 24.
25.
26.
27. 28. 29. 30. 31.
32.
33.
34.
ferentiation, pp. 167-183. Cambridge University Press, Cambridge (1973). Mager WH: Control of ribosomal protein gene expression. Biochim Biophys Acta 949:1-15 (1988). Martinez MC, Jorgenson J-E, Lawton MA, Lamb CJ, Doerner PW: Spatial pattern of cdc2 expression in relation to meristem activity and cell proliferation during plant development. Proc Natl Acad Sci USA 89:7360-7364 (1992). Meyerson M, Enders GH, Wu C-L, Su L-K, Gorka C, Nelson C, HarlowE, TsaiL-H: A family of human cdc2-related protein kinases. EMBO J 11:2909-2917 (1992). Mikami K, Iwabuchi M: Regulation of cell cycle-dependent gene expression. In: VermaDPS (ed) Control of Plant Gene Expression, pp. 51-68. CRC Press, Boca Raton, FL (1992). Murray AW: Creative blocks: cell-cycle checkpoints and feedback controls. Nature 359:599-604 (1992). Murray AW, Hunt T: The Cell Cycle: An Introduction. W. H. Freeman, New York (1993). Murray JAH: Plant cell division: the beginning of START. Plant Mol Biol 2 6 : 1 - 3 (1994). Osley MA: Regulation of histone synthesis in the cell cycle. Annu Rev Biochem 60:827-861 (1991). Reichheld J-P, Sonobe S, C16ment B, Chaubet N, Gigot C: Cell cycle-regulated histone gene expression in synchronized plant cells. Plant J 7:245-252 (1995). Renaudin J-P, Colasanti J, Rime H, Yuan Z, Sundaresan V: Cloning of four cyclins from maize indicates that higher plants have three structurally distinct groups of mitotic cyclins. Proc Natl Acad Sci USA 91:7375-7379 (1994). Stafstrom JP: Axillary bud development in pea: apical dominance, growth cycles, hormonal regulation and plant architecture. In: Amasino RM (ed) Cellular Communication in Plants, pp. 75-86. Plenum Press, New York (1993). StafstromJP, Altschuler M, Anderson DH: Molecular cloning and expression of a MAP kinase homologue from pea. Plant Mol Biol 22:83--90 (1993).
35. Stafstrom JP, Sussex IM: Patterns of protein synthesis in dormant and growing vegetative buds of pea. Planta 176: 497-505 (1988). 36. Stafstrom JP, Sussex IM: Expression of a ribosomal protein gene in axillary buds of pea. Plant Physiol 100: 14941502 (1992). 37. Staiger C, Doonan J: Cell division in plants. Curr Opin Cell Biol 5:226-231 (1993). 38. TanimotoEY, RostTL, ComaiL: DNA replicationdependent histone H2A mRNA expression pea root tips. Plant Physiol 103:1291-1297 (1993). 39. Terada R, Nakayama T, lwabuchi M, Shimamoto K: A wheat histone H3 promoter confers cell division-dependent and -independent expression of the gusA gene in transgenic rice plants. Plant J 3:241-252 (1993). 40. Van't H o f J : The regulation of cell division in higher plants. In: Basic Mechanisms in Plant Morphogenesis, Brookhaven Symposia vol. 25, pp. 152-165 (1973). 41. Van't H o f J , Bjerknes CA, Lamm SS: Meristematic precursors of vascular parenchyma differentiate from G2 phase after replicating D N A discontinuously. Am J Bot 73:87-95 (1986). 42. Van't HofJ, Kovacs CJ: Mitotic delay in two biochemically different G 1 cell populations in cultured roots of pea (Pisum sativum). Radiation Res 44:700-714 (1970). 43. Van't HofJ, Kovacs CJ: Mitotic regulation in the meristem of cultured roots: the principal control point hypothesis. In: Miller MW, Kuehnert CC (eds) The Dynamics of Meristem Cell Populations, pp. 15-32. Plenum Press, New York (1972). 44. Van't HofJ, Lamm SS, Bjerknes CA: Detection of replication initiation by a replicon family in DNA of synchronized pea (Pisum sativum) root cells using benzoylated naphthoylated DEAE-cellulose chromatography. Plant Mol Biol 9:77-86 (1987). 45. Wells D, McBride C: A comprehensive compilation and alignment of histones and histone genes. Nucl Acids Res 17:r311-r346 (1989).
