Growing the
1. Introduction
Ribonucleoside diphosphate reductase (RNR) of
The best-known defective RNR mutant in
The RNR101 protein is inactivated at 42°C
These results are consistent with RNR having a thermoresistance period due to protection by some subcellular structure. This enzyme has been proposed to be part of a complex for the biosynthesis of dNTP (Mathews, 1993) therefore the association with this complex might explain such protection. We have proposed that, as a component of the replication hyperstructure, the RNR101 protein would be protected from thermal inactivation and that this would suffice to allow chromosome replication for 50 min in restrictive conditions (Guzmán et al., 2002; 2003, Molina & Skarstad, 2004; Guarino et al., 2007a; 2007b; Riola et al., 2007).
Supporting this model, RNR has been colocalized with the replisome-associated proteins DnaB helicase and DNA polymerase τ subunit, and with the fork-associated protein SeqA (Fig. 2) (Sánchez-Romero et al., 2010).
Furthermore, a hyperstructure containing RNR101 impairs replication fork progression even at the permissive temperature (Guarino et al., 2007a). Arrest of replication forks is known to cause double-strand breaks, DSBs (Bierne & Michel, 1994; Kuzminov, 1995). We have shown that the number of DSBs in the
Relevant phenotype | % linear DNA a | |
|
4.58 | ±2.51 |
|
15.18 | ±2.83 |
|
6.74 | ±2.60 |
|
5.72 | ±1.41 |
|
24.79 | ±7.05 |
|
12.64 | ±4.70 |
|
5.94 | ±2.19 |
It is intriguing that rifampicin or cloramphenicol addition, as well as the presence of a
In studying replication in the
Consequently, we propose that a reduction in the number of forks replicating the chromosome results in an improvement in the quality of replication that allows the deficient replication hyperstructure of the
2. Reduction in the overlap of replication rounds improves fork progression at the restrictive temperature in a nrdA101 strain
We have previously shown that, due to an elongation of the replication period lasting more than twice the cell cycle at 30°C (
2.1. Experimental approach
In contrast with eukaryotic organisms, the time required to replicate a single chromosome (
As explained above, the overlap of the replication rounds depends on two parameters, the generation time, τ, and the
2.2. Increasing the generation time
We lowered
As expected from growing the bacteria in a carbon source different from glucose, a lengthening of the generation time and a lowering in the number of overlapped replication rounds,
2.3. Reducing the C period
2.3.1. By the presence of dnaA defective alleles
The presence of
When incubated for 4 h at 42°C in the presence of cephalexin, the DNA content per cell in
We have verified that the number of overlapped replication forks per chromosome at 30°C could be also lowered by introducing
These results suggest that a lowering in the number of replication forks running along the chromosome could improve the progression of replication in the
In addition, over-expression of the
Strains | τ | ΔG30°C | 42°C | 42°C/ΔG30°C | n | ori/ter | C |
|
78 | 0.55 | nt | nt | 1.37 | 2.58 | 107 |
|
79 | 1.03 | 0.52 | 0.50 | 2.36 | 5.13 | 186 |
|
105 | 0.59 | 0.83 | 1.40 | 1.46 | 2.75 | 153 |
|
112 | 0.35 | 0.58 | 1.65 | 0.91 | 1.87 | 102 |
|
78 | 0.55 | 1.40 | 2.54 | 1.37 | 2.58 | 107 |
|
80 | 0.45 | 0.48 | 1.06 | 1.15 | 2.21 | 92 |
|
82 | 1.00 | 0.45 | 0.45 | 2.30 | 4.92 | 188 |
|
81 | 0.73 | 0.70 | 0.95 | 1.76 | 3.38 | 142 |
|
85 | 0.55 | 0.95 | 1.80 | 1.26 | 2.39 | 107 |
hyperstructure would increase the supply of dNTP to the replication enzymes (Pato, 1979; Mathews, 1993).
It has been shown that the
2.3.2. By increasing the number of copies of the datA sequence
The
2.3.3. By deleting the DARS sequence
The DnaA protein is a member of the AAA+ ATPase family and has an exceptionally high affinity for ATP/ADP (Sekimuzu et al., 1987; Kaguni, 2006). The level of cellular ATP-DnaA oscillates during the replication cycle, peaking around the time of initiation (Kurokawa et al., 1999).
Katayama's group has recently found two chromosomal intergenic regions termed DARS1 and DARS2 (
Our data show that decreasing the number of replication rounds (
Furthermore, a lower availability of wild type DnaA protein induced by the presence of extra copies of the
These observations, together with our data, are consistent with the idea that the progression of replication forks is not merely responsive to elongation factors (dNTP pools or proteins engaged in elongation) but also to the number of forks running along the chromosome. We suggest that the best explanation for the reduction of the
3. Stalled multifork chromosomes as the cause of aberrant DNA segregation and cell death in the nrdA101 mutant at the restrictive temperature
Growth of
Cell viability was studied in all the growth media and strains described above. Cells were grown at 30°C and when the cultures reached mid-logarithmic phase (about 0.1 OD550), an aliquot of each culture was incubated at 42°C and the number of viable cells were estimated by serial dilution and plating on rich medium at 30°C. Viability is expressed relative to the onset of treatment. Growing
Nucleoid segregation analysis was performed in aliquots of the cultures incubated at 42°C in the presence of cephalexin (50 µg/ml) for 4 hours plus, during the last 20 min, chloramphenicol addition(200µg/ml) to condense nucleoids. Micrographs of DAPI stained cells show a high number of cells containing an abnormal number of nucleoids randomly distributed along the filaments (Fig. 5) (Riola et al., 2007). An increased number of cells containing normal and well-segregated nucleoids were found in cells grown in arabinose or in glycerol (Fig. 5). The anomalous number and distribution of nucleoids found in the
The above results reveal a good correlation between the overlap of replication rounds and aberrant nucleoid segregation and cell lethality. This correlation is consistent with the hypothesis that these problems are associated with a highly forked chromosome structure. The detrimental effects of such chromosomes are reduced or eliminated by any environmental or genetic modification that reduces replication overlap. We therefore suggest that the observed morphological alterations of
overlaps, stalled forks have less opportunity to be repaired and restarted and this interferes with subsequent forks. This results in chromosomal abnormalities, disrupted chromosome and nucleoid segregation, loss of cell division, and, finally, cell death.
DNA topology has been found to play an important role in the segregation of duplicated chromosomes (Dasgupta et al., 2000; Holmes & Cozarelli, 2000). Consequently, a disturbed DNA topology due to a highly forked chromosome structure, could contribute to the altered nucleoid segregation observed in the
4. The number of replication rounds in the chromosome limits the replication rate of individual forks
In the
It is difficult to decide whether the reduction in the number of forks is the consequence of an increased replication rate (as
The second proposition is that the elongation rate increased as a consequence of the reduction of the number of forks or the replication overlap. This reduction in the number of the forks would be caused by the deficiency of any factor required for the initiation step since this would result in the delay of the initiation of replication.
In the above work, we have shown that a decrease in the growth rate of the
An unified explanation for all the results presented here is difficult to find. Clearly though, the underlying mechanism should explain the precise correlation between initiation and elongation that tunes DNA replication to any environmental circumstance. Whatever the nature of this mechanism, reduction in the number of forks per chromosome or decreased overlapping of consecutive replication rounds might increase the elongation rate by providing
a better overall chromosome structure, including discrete regional organization and supercoiling domains,
an increased availability of a limiting constituent required for replication and/or for segregation, and
an increased time for the repair and restart of a stalled fork so as to avoid collision with the next fork.
This homeostatic regulation between the numbers and velocities of forks would also explain how the replication rate compensates for widely varying replication origins and activities in eukaryotes (Conti et al., 2007).
5. Balance between the number of origins and elongation rates as a general regulatory mechanism in the control of eukaryotic cell cycle
In eukaryotic cells, the DNA replication program is organized according to multiple tandem replicons that span each chromosome. Each replicon is replicated bidirectionally by a pair of replication forks that increase their rates up to three fold towards the end of S phase. Furthermore, the rate of the replication fork progression varies up to ten-fold or more depending on the distance between origins in different conditions or cell types (Housman & Huberman, 1975; reviewed in Herrick, 2010). Two replication regimes with distinct kinetics govern duplication of the genome: in the first half of the S phase, when the gene-rich euchromatin is predominantly replicated, the density of the activated replication origins steadily increases to about twice the initial value; during the second half of the S phase, when the gene-poor heterochromatin tends to be replicated, the density of active replication origins increases substantially by about ten fold (Herrick & Bensimon, 2008). It has been proposed that this mechanism would guarantee the rapid and complete duplication of the genome. Nevertheless, in mammalian cells the relationship between origin activation, the size of replicons (50-300kb) and the existence of multiple potential origins remains to be elucidated (Herrick, 2010).
The efficient duplication of the eukaryotic genome depends on the orderly activation of the origins, estimated to be ten thousand, and on the proper progression of their forks. The coordinated activation of origins is insufficient on its own to account for timely completion of genome duplication when interorigin distances vary significantly and fork velocities are constant. Therefore the coordination and compensation between origin spacing and fork progression may be one of the mechanisms for the complete duplication of the genome in the limited amount of time of the S phase. By using a single-molecule approach based on molecular combing, the interorigin distances and replication fork velocities over extensive regions of the genome have been measured in both primary keratinocytes and cancer cells (Conti et al., 2007). This study provides evidence for the direct correlation between the interorigin distances and the replication rates, insofar as the further the origins are from one another, the faster the forks progress. These results are in agreement with the results of this and other studies of
Figure 3 in Conti et al., 2007 shows a significant linear correlation between these two parameters in eukaryotic cells, consistent with a biological mechanism that coordinates replication fork progression with interorigin distance. The mechanism that allows replication forks to adjust their speed is unknown. Nevertheless the possibilities for the nature of this mechanism are similar to the ones proposed above for
6. Concluding remarks
In this work we show that reducing the number of replication forks per chromosome in
variations in the availability of some limiting hyperstructure component which might lead to assembly of an inefficient hyperstructure when a high number of forks compete for this component, or
the structural constraints caused by a chromosome undergoing several rounds of replication running at the same time.
Results from other research groups, reviewed above, and comparison with DNA replication in eukaryotes provide further evidence that, in widely different systems, the initiation and the elongation of chromosome replication are not independent processes.
Acknowledgments
We are very grateful to Kirsten Skarstad and Tsutomu Katayama for bacterial strains and plasmids. We especially thank Encarna Ferrera for her technical help. This work was supported by grant BFU2007-63942 from the Ministerio de Ciencia e Innovación. IS, CM and MAS-R acknowledge the studentship from Junta de Extremadura.
References
- 1.
Anglana M. Apiou F. Bensimon A. Debatisse M. 2003 Dynamics of DNA replication in mammalian somatic cells: nucleotide pool modulates origin choice and interorigin spacing. 114 385 94 . - 2.
Atlung T. Hansen F. G. 2002 Effect of different concentrations of H-NS protein on chromosome replication and the cell cycle in Escherichia coli. J Bacteriol184 1843 1850 . - 3.
Blinkova A. Hermandson M. Walker J. R. 2003 Suppression of Temperature-Sensitive Chromosome Replication of an Escherichia coli dnaX(Ts) Mutant by Reduction of Initiation Efficiency. .185 3583 3595 . - 4.
Bierne H. Michel B. 1994 When replication forks stop. 13 17 23 . - 5.
Bussiere D. E. Bastia D. 1999 Termination of DNA replication of bacterial and plasmid chromosomes .31 1611 1618 . - 6.
Conti C. Sacca B. Herrick J. Lalou C. Pommier Y. Bensimon A. 2007 Replication fork velocities at adjacent replication origins are coordinately modified during DNA replication in human cells. .18 3059 3067 . - 7.
Cooper S. Helmstetter C. E. 1968 Chromosome replication and the division cycle of Escherichia coli B/r. 31 519 540 . - 8.
Dasgupta S. Maisnier-Patin S. Nordstrom K. 2000 New genes with old modus operandi. The connection between supercoiling and partitioning of DNA in Escherichia coli .1 323 327 . - 9.
Dix D. E. Helmstetter C. E. 1973 Coupling between chromosome completion and cell division in Escherichia coli .115 786 95 . - 10.
Fuchs J. A. Karlstrom H. O. Warner H. R. Reichard P. 1972 Defective gene product in dnaF mutant of Escherichia coli. 238 69 71 . - 11.
Fujimitsu K. Senriuchi T. Katayama T. 2009 Specific genomic sequences of E. coli promote replicational initiation by directly reactivating ADP-DnaA .23 1221 33 . - 12.
Gines-Candelaria E. Blinkova A. Walker J. R. 1995 Mutations in Escherichia coli dnaA which suppress a dnaX(Ts) polymerization mutation and are dominant when located in the chromosomal allele and recessive on plasmids.177 705 715 . - 13.
Gon S. Camara J. E. Klungsoyr H. K. Crooke E. Skarstad K. Beckwith A. J. 2006 A novel regulatory mechanism couples deoxyribonucleotide synthesis and DNA replication in Escherichia coli. 25 1137 1147 . - 14.
Grompe M. Versalovic J. Koeuth T. Lupski J. R. 1991 Mutations in the Escherichia coli dnaG gene suggest coupling between DNA replication and chromosome partitioning. 173 1268 1278 . - 15.
Guarino E. Jiménez-Sánchez A. Guzmán E. C. 2007 Defective ribonucleoside diphosphate reductase impairs replication fork progression in Escherichia coli. 189 3496 501 . - 16.
Guarino E. Salguero I. Jiménez-Sánchez A. Guzmán E. C. 2007 Double-strand break generation under deoxyribonucleotide starvation in Escherichia coli. 189 5782 6 . - 17.
Guzmán E. C. Caballero J. L. Jiménez-Sánchez A. 2002 Ribonucleoside diphosphate reductase is a component of the replication hyperstructure in Escherichia coli. 43 487 95 . - 18.
Guzmán E. C. Guarino E. Riola J. Jiménez-Sánchez A. 2003 Ribonucleoside diphosphate reductase is a functional and structural component of the replication hyperstructure in1 29 43 . - 19.
Hanke P. D. Fuchs J. A. 1983 Regulation of ribonucleoside diphosphate reductase mRNA synthesis in Escherichia coli .154 1040 1045 . - 20.
Herrick J. 2010 The dynamic replicon: adapting to a changing cellular environment .32 153 64 . - 21.
Herrick J. Sclavi B. 2007 Ribonucleotide reductase and the regulation of DNA replication: an old story and an ancient heritage. 63 22 34 . - 22.
Herrick J. Bensimon A. 2008 Global regulation of genome duplication in eukaryotes: an overview from the epifluorescence microscope .117 243 260 . - 23.
Hill T. M. Sharma B. Valjavec-Gratian M. Smith J. 1997 sfi-independent filamentation in Escherichia coli is lexA dependent and requires DNA damage for induction. 179 1931 9 . - 24.
Holmes V. F. Cozzarelli N. R. 2000 Closing the ring: links between SMC proteins and chromosome partitioning, condensation, and supercoiling .97 1322 1324 . - 25.
Housman D. Huberman J. A. 1975 Changes in the rate of DNA replication fork movement during S phase in mammalian cells.94 173 181 . - 26.
Jaffe A. D’Ari R. Norris V. 1986 SOS-independent coupling between DNA replication and cell division in E. coli. .165 66 71 . - 27.
Jannière L. Canceill D. Suski C. Kanga S. Dalmais B. Lestini R. Monnier A. F. Chapuis J. Bolotin A. Titok M. Le Chatelier E. Ehrlich S. D. 2007 Genetic evidence for a link between glycolysis and DNA replication. 2(5):e447. - 28.
Jiménez-Sánchez A. Guzmán E. C. 1988 Direct procedure for the determination of the number of replication forks and the reinitiation fraction in bacteria. 4 431 433 . - 29.
Kawakami H. Iwura T. Tanaka M. Sekimizu T. 2001 Arrest of cell division and nucleoid partition by genetic alterations in the sliding clamp of the replicase and in DnaA. 266 167 79 . - 30.
Kaguni J. M. 2006 DnaA: Controlling the Initiation of Bacterial DNA Replication and More. 60 351 71 . - 31.
Kim J. Wheeler L. J. Shen R. Mathews C. K. 2005 Protein-DNA interactions in the T4 dNTP synthetase complex dependent on gene 32 single-stranded DNA-binding protein .55 1502 1514 . - 32.
Kuzminov A. 1995 Collapse and repair of replication forks in Escherichia coli. 16 373 384 . - 33.
Kitagawa R. Mitsuki H. Okazaki T. Ogawa T. 1996 A novel DnaA protein-binding site at 94.7 min on the Escherichia coli chromosome. 19 1137 47 . - 34.
Kitagawa R. Ozaki T. Moriya S. Ogawa T. 1998 Negative control of replication initiation by a novel chromosomal locus exhibiting exceptional affinity for Escherichia coli DnaA protein 12 3032 43 . - 35.
Kurokawa K. Nishida S. Emoto A. Sekimizu K. Katayama T. 1999 Replication cycle-coordinated change of the adenine nucleotide-bound forms of DnaA protein in Escherichia coli .18 6642 6652 . - 36.
Løbner-Olesen A. Slominska-Wojewodzka M. Hansen F. G. Marinus M. G. 2008 DnaC Inactivation in Expression of Nucleotide Biosynthesis Genes. 3(8): e2984. - 37.
Malínsky J. Koberna K. Stanĕk D. Masata M. Votruba I. Raska I. 2001 The supply of exogenous deoxyribonucleotides accelerates the speed of the replication fork in early S-phase.114 747 50 . - 38.
Maisnier-Patin S. Nordstrom K. Dasgupta y. S. 2001 Replication arrests during a single round of replication of the Escherichia coli chromosome in the absence of DnaC activity .42 1371 1382 . - 39.
Mathews C. K. 1993 Enzyme organization in DNA precursor biosynthesis. 44 167 203 . - 40.
Messer W. Weigel C. 2003 DnaA as a transcription regulator. 370 338 49 . - 41.
Michel B. Grompone G. Flores M. J. Bidnenko V. 2004 Multiple pathways process stalled replication forks .101 12783 12788 . - 42.
Michelsen O. Teixeira de Mattos. M. J. Jensen P. R. Hansen F. G. 2003 Precise determinations of C and D periods by flow cytometry in Escherichia coli K-12 and B/r .149 1001 10 . - 43.
Molina F. Skarstad K. 2004 Replication fork and SeqA focus distributions in Escherichia coli suggest a replication hyperstructure dependent on nucleotide metabolism .52 1597 612 . - 44.
Morigen Boye. E. Skarstad K. Løbner-Olesen A. 2001 Regulation of chromosomal replication by DnaA protein availability in Escherichia coli: effects of the datA region. 1521 73 80 . - 45.
Morigen-Olesen Løbner.A Skarstad K. 2003 Titration of the Escherichia coli DnaA protein to excess datA sites causes destabilization of replication forks, delayed replication initiation and delayed cell division .50 349 62 . - 46.
Morigen Odsbu. I. Skarstad K. 2009 Growth rate dependent numbers of SeqA structures organize the multiple replication forks in rapidly growing Escherichia coli. 14 643 657 . - 47.
Naruyama H. Shimada M. Niida H. Zineldeen D. H. Hashimoto Y. Kohri K. Nakanishi M. 2008 Essential role of Chk1 in S phase progression through regulation of RNR2 expression . .374 79 83 . - 48.
Nordlund P. Reichard P. 2006 Ribonucleotide reductases. 75 681 706 . - 49.
Odsbu I. Morigen Skarstad K. 2009 A reduction in ribonucleotide reductase activity slows down the chromosome replication fork but does not change its localization. 4 (10),e7617 EOF . - 50.
Olliver A. Saggioro C. Herrick J. Sclavi B. 2010 DnaA-ATP acts as a molecular switch to control levels of ribonucleotide reductase expression in Escherichia coli .76 1555 71 . - 51.
Pato M. L. 1979 Alterations of deoxyribonucleoside triphosphate pools in Escherichia coli: effects on deoxyribonucleic acid replication and evidence for compartmentation. 140 518 24 . - 52.
Pritchard R. H. Zaritsky A. 1970 Effect of Thymine Concentration on the replication velocity of DNA in a thymineless mutant of Escherichia coli. 226 126 130 . - 53.
Riola J. Guarino E. Guzmán E. C. Jiménez-Sánchez A. 2007 Differences in the degree of inhibition of NDP reductase by chemical inactivation and by the thermosensitive mutation nrdA101 in Escherichia coli suggest an effect on chromosome segregation .12 70 81 . - 54.
Salguero I. López Acedo. E. Guzmán E. C. 2011 Overlap of replication rounds disturbs the progression of replicating forks in a ribonucleotide reductase mutant of Escherichia coli .157 1955 1967 . - 55.
Sánchez Romero. M. A. Molina F. Jiménez-Sánchez A. 2010 Correlation between ribonucleoside-diphosphate reductase and three replication proteins in Escherichia coli. doi: 10.11 EOF 86/1471-2199-11-11. - 56.
Sekimizu K. Bramhill D. Kornberg A. 1987 ATP activates DnaA protein in initiating replication of plasmids bearing the origin of the E. coli chromosome. 50 259 65 . - 57.
Schaper S. Messer W. 1995 Interaction of the initiator protein DnaA of Escherichia coli with its DNA target. .270 17622 6 . - 58.
Skarstad K. Steen H. B. Boye E. 1985 Escherichia coli DNA distributions measured by flow cytometry and compared with theoretical computer simulations. 163 661 8 . - 59.
Skovgaard O. Lobner-Olesen A. 2005 Reduced initiation frequency from oriC restores viability of a temperature-sensitive Escherichia coli replisome mutant .151 963 73 . - 60.
Stepankiw N. Kaidow A. Boye E. Bates D. 2009 The right half of the Escherichia coli replication origin is not essential for viability, but facilitates multi-forked replication .74 467 479 . - 61.
Sueoka N. Yoshikawa H. 1965 The chromosome of Bacillus subtilis. I. Theory of marker frequency analysis .52 747 757 . - 62.
Torheim N. K. Boye E. Løbner-Olesen A. Stokke T. Skarstad K. 2000 The SeqA protein destabilizes mutant DnaA204 protein. Mol Microbiol.37 629 38 . - 63.
Uhlin U. Eklund H. 1994 Structure of ribonucleotide reductase protein R1. 13 533 539 . - 64.
Von Freiesleben. U. Rasmussen K. V. Atlung T. Hansen F. G. 2000 Rifampicin-resistant initiation of chromosome replication from oriC in ihf mutants .37 1087 1093 . - 65.
Werner R. 1971 Nature of DNA precursors. 233 99 103 . - 66.
Wheeler L. J. Rajagopal I. Mathews C. K. 2005 Stimulation of mutagenesis by proportional deoxyribonucleoside triphosphate accumulation in Escherichia coli. 4 1450 1456 . - 67.
Zaritsky A. Pritchard R. H. 1971 Replication time of the chromosome in thymineless mutants of Escherichia coli. 60 65 74 . - 68.
Zaritsky A. Vischer N. Rabinovitch A. 2007 Changes of initiation mass and cell dimensions by the ‘eclipse’.63 15 21 .