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Applied and Environmental Microbiology, August 1999, p. 3651-3659, Vol. 65, No. 8
Department of Biological Sciences, University
of Stirling, Stirling FK9 4LA, United Kingdom
Received 12 March 1999/Accepted 24 May 1999
Diel periodicity in the expression of key genes involved in carbon
and nitrogen assimilation in marine Synechococcus spp. was
investigated in a natural population growing in the surface waters of a
cyclonic eddy in the northeast Atlantic Ocean.
Synechococcus sp. cell concentrations within the upper
mixed layer showed a net increase of three- to fourfold during the
course of the experiment (13 to 22 July 1991), the population
undergoing approximately one synchronous division per day. Consistent
with the observed temporal pattern of phycoerythrin (CpeBA)
biosynthesis, comparatively little variation was found in
cpeBA mRNA abundance during either of the diel cycles
investigated. In marked contrast, the relative abundance of transcripts
originating from the genes encoding the large subunit of ribulose
bisphosphate carboxylase/oxygenase (rbcL) and glutamine
synthetase (glnA) showed considerable systematic temporal
variation and oscillated during the course of each diel cycle in a
reciprocal rhythm. Whereas activation of rbcL transcription was clearly not light dependent, expression of glnA
appeared sensitive to endogenous changes in the physiological demands
for nitrogen that arise as a natural consequence of temporal
periodicity in photosynthetic carbon assimilation. The data presented
support the hypothesis that a degree of temporal separation may exist between the most active periods of carbon and nitrogen assimilation in
natural populations of marine Synecoccoccus spp.
Marine Synechococcus spp.
are among the most abundant and cosmopolitan members of the
photosynthetic picoplankton: a taxonomically mixed assemblage of small
(<2.0-µm diameter) prokaryotic and eukaryotic microorganisms that
account for the major fraction of primary production in the world's
open oceans (40). Although genetically divergent (14,
51, 54, 66, 67), oceanic strains of Synechococcus spp.
belong to a physiologically coherent group (marine cluster A) of
cyanobacteria within the unicellular order
Chroococcales (59). All known isolates are
non-nitrogen-fixing, obligate photoautotrophs (59, 62) that
produce spectroscopically distinct biliproteins (including
phycoerythrin [PE]) as accessory components of their light-harvesting
apparatuses (1, 35, 36). Many are also capable of
flagellum-free swimming motility and, in one demonstrated case at
least, display positive chemotaxis toward a variety of organic and
inorganic nitrogenous compounds (64).
Following their first description in 1979 (22, 61), a
reasonable (if incomplete) understanding has developed of the
biological and physicochemical factors that regulate the growth and
productivity of marine Synechococcus spp. in the world's
oceans (18, 19, 23, 24, 29, 46, 47, 56, 60, 65, 66). While
there are undoubted exceptions to the paradigm, these organisms are generally at their most conspicuous (but not necessarily their most
productive) in oligotrophic surface waters, where the limited supply of
inorganic nutrients appears to exclude competition in the form of
larger and potentially faster-growing eukaryotic phytoplankton (19, 23, 62).
Estimates of Synechococcus sp. growth rates in open waters
are somewhat variable, but for active populations they tend to cluster
around a mean of about one division per day (7, 18, 29, 56).
Like many groups of marine phytoplankton, there is mounting evidence
that the daily progression through the Synechococcus sp.
cell cycle may be entrained in situ by the natural diel alternation in
irradiance. While a number of previous studies have clearly hinted at
the phenomenon (7, 8, 25, 62), the recent cytological
investigation conducted by Vaulot and coworkers (56) was the
first to confirm that DNA replication and cell division can become
tightly synchronized in Synechococcus spp. growing in open
waters. If such rhythmic behavior is a common feature of natural
populations, an obvious rationale can be developed to explain several
earlier reports (20, 25, 34) of diel oscillations in cell
rRNA content and macromolecular composition in these organisms. Like
the diel phasing of DNA replication and cytokinesis (56),
temporal patterns of this type are an entirely predictable feature of
synchronously dividing populations.
Diel rhythms in the synthesis and accumulation of various mRNAs in
natural populations of Synechococcus spp. and other marine cyanobacteria have also been described (10, 26, 37, 38, 70),
but whether these might have their origin in cell cycle-related events
has not received consideration. Cell division in the marine Synechococcus sp. strain WH7803 is known to be under
circadian control (50), and there is convincing preliminary
evidence that expression of the rbcL gene (encoding the
large subunit of the Calvin cycle enzyme, ribulose bisphosphate
carboxylase/oxygenase [RubisCO]) may also be regulated in this way
(37). It is an open question, however, whether
rbcL transcription in Synechococcus spp. is
controlled directly by as-yet-unidentified clock genes or is regulated
in response to temporal oscillations in other metabolic processes that
may themselves be circadian in nature.
The RubisCO genes of marine Synechococcus spp. are
phylogenetically distinct from those of other cyanobacteria and may
have been acquired comparatively recently by lateral gene transfer (63). In the marine Synechococcus sp. strain
WH7803, both RubisCO subunits are cotranscribed with a homologue of the
Synechococcus sp. strain PCC7942 ccmK gene
(63), which encodes a structural component of the
carboxysome, the site of CO2 fixation in cyanobacteria. RubisCO expression is enhanced considerably by light and, as in other
cyanobacteria (11, 27), appears to be regulated primarily at
the level of transcription. In this regard, the temporal pattern of
RubisCO expression displayed by Synechococcus sp. strain
WH7803 in culture closely parallels that seen in natural populations of
photosynthetic picoplankton growing in subtropical oceanic surface
waters (37, 38).
By analogy to the adaptive strategies adopted by other cyanobacteria
growing at low concentrations of combined nitrogen, the principal route
of inorganic nitrogen assimilation in marine Synechococcus spp. is thought to occur via the glutamine synthetase (GS)-glutamate synthase pathway and to be dependent upon photosynthetically generated ATP and reductant (31, 53). While the existence of
alternative pathways has not been excluded experimentally, from what is
known of the kinetic properties of these systems in other cyanobacteria (32) it appears unlikely they could make other than a very
minor contribution to N assimilation in Synechococcus spp.
at the low ambient nitrogen concentrations typical of oceanic surface waters.
The pathways of carbon and nitrogen assimilation are metabolically
linked at the level of GS in cyanobacteria, and the enzyme is
extensively regulated in response to a variety of environmental signals. GS is rapidly inactivated in the dark and also by ammonium ions in the well-studied model system Synechocystis sp.
strain PCC6803 and several other cyanobacteria (45, 49). The
nature of the reversible modification system present in
Synechocystis sp. strain PCC6803 is distinct from that
described for the GS of enteric bacteria, however, in that
adenylylation is not involved. Nevertheless, some elements of the
signal transduction system that lead to the activation or inactivation
of GS do appear to be conserved between cyanobacteria and enteric
bacteria. The regulatory protein PII (encoded by glnB) is
present in both groups but in cyanobacteria is activated by
phosphorylation rather than uridinylylation (16). As with
GS, activation of PII is regulated by nitrogen availability and is
dependent upon photosynthetic electron transport (17, 45).
Transcription of glnA and glnB in
Synechococcus sp. strain PCC 7942 and
Synechocystis sp. strain PCC 6803 is regulated by the
positive transcription factor NtcA, a functional analogue of the
nitrogen regulator of enteric bacteria, NtrC (17, 57). NtcA
is widely distributed among cyanobacteria (including the marine strain
Synechococcus sp. strain WH7803 [28]) and
in the absence of ammonium binds to the promoter regions of a suite of N-regulated genes, including glnA and glnB.
Transcription of glnA increases two- to threefold in cells
of Synechococcus sp. strain PCC7942 switched from ammonium-
to nitrate-containing growth medium and by a somewhat greater margin
(5- to 10-fold) during nitrogen deprivation (13).
Enhanced glnA transcription in the absence of ammonium leads
to parallel increases in GS synthesis and activity (12, 13), and a similar correlation among nitrogen regimen, glnB mRNA
abundance, and PII protein levels has been reported recently for
Synechocystis sp. strain PCC6803 (16, 52).
Expression of glnA at the transcriptional level is not light
dependent in Synechococcus sp. strain PCC 7942 (6), but like psbA (encoding the D2 protein of
photosystem II) and the rbcL gene of marine
Synechococcus spp. (37), there is some evidence
for circadian control (30).
Establishing how temporal and spatial variability in the environment
structures the molecular biological responses of natural populations of
marine microorganisms is a central theme in contemporary biological
oceanography. The present field-based study was designed to examine the
temporal (diel) periodicity in the abundance of three different
Synechococcus sp. mRNAs that encode proteins central to
photosynthetic light harvesting (PE; cpeBA), carbon fixation (RubisCO; rbcL), and nitrogen assimilation (glutamine
synthetase; glnA). The aims of the investigation were to
assess the extent to which the three genes might be differentially
expressed during the diel cycle and to relate the findings to the
temporal pattern of Synechococcus sp. population growth and
cell division in situ. The data reported reveal a quasireciprocal
rhythm in the relative abundance of rbcL and glnA
mRNAs and point to at least some degree of temporal separation between
the most active phases of C and N assimilation in natural populations
of Synechococcus spp.
Observations were made during a 10-day period in July 1991 at a
series of drifting stations located within a cyclonic eddy in the
northeast Atlantic Ocean (Fig. 1). The
study site was selected during passage of the research vessel (RRS
Charles Darwin; cruise CD61) to the sea area to the south of
Iceland following receipt of thermal infrared satellite images of the
region on 10 July 1991. During the previous month an extensive
mesoscale (~250,000-km2) bloom of coccolithophorid algae
had occurred in these subpolar waters (15, 21), but by the
time the research vessel arrived on station (12 July 1991) all surface
expression of the bloom had disappeared (21). Mapping of
surface hydrographic and biological properties within the eddy revealed
that the postbloom phytoplankton community comprised a mixed assemblage
of micro- and picophytoplankton dominated by diatoms and
microflagellates and by Synechococcus spp., respectively.
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Diel Rhythms in Ribulose-1,5-Bisphosphate Carboxylase/Oxygenase
and Glutamine Synthetase Gene Expression in a Natural Population of
Marine Picoplanktonic Cyanobacteria (Synechococcus
spp.)
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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FIG. 1.
Track of the ARGOS drifter deployed on 13 July 1991 (Julian day 194). The position of the drifter at 0000 hours on each
subsequent day is indicated by the day number. (Data courtesy of Bob
Barrett and Robin Pingree.)
A drogued (10 m) ARGOS drifter was deployed at the center of the eddy (61°05'N, 20°W) on 13 July 1991. Thereafter, the research vessel was operated in Lagrangian mode by maintaining close station with the drifter until the morning of 22 July 1991 (for further details of the study site and its hydrographic properties, see reference 21). Plankton samples were obtained from near-surface (~5 m) waters by using an outlet of the ship's nontoxic seawater supply (residence time from inlet to outlet, ~2 min) or from discrete depths by using 30-liter GO-Flo bottles (General Oceanics, Miami, Fla.) deployed from a Kevlar line. Surface nutrient concentrations were determined continuously while on station with a Technicon autoanalyzer and standard analytical procedures (5, 21).
Synechococcus sp. cell counts. The abundance of Synechococcus sp. cells was estimated by epifluorescence microscopy (19). Seawater samples of known volume (10 to 100 ml) were prescreened through a 250-µm-mesh plankton net, and phytoplankton cells were collected by filtration on 25-mm-diameter, 0.2-µm-pore-size polycarbonate membranes (Nuclepore Corp.) overlaying a Whatman GF/F support filter. When necessary (i.e., during times of excessive ship movement), filtered samples mounted on glass slides in nonfluorescent immersion oil were stored in darkness at 4°C, but in all cases they were counted onboard ship within 24 h of collection.
Determination of PE concentrations.
Synechococcus sp.
PE concentrations were determined as previously described with
Synechococcus sp. strain WH7803 cells as the reference
standard (69). Seawater samples (1 to 2 liters) were
fractionated through 47-mm-diameter 2.0- and 0.6-µm-pore-size polycarbonate filters (Poretics Corp.) under gentle vacuum (<10 cm of
Hg), and Synechococcus sp. cells retained on the
0.6-µm-pore-size filter were washed into 1.5 ml of Whatman
GF/F-filtered seawater (collected from a depth of 100 m). Owing to
equipment failure, it was not possible to carry out PE analyses at sea.
The concentrated samples were centrifuged at 13,000 × g for 1 min, the seawater supernatant was aspirated, and the
pelleted phytoplankton cells were taken up in 1 ml of 50% (vol/vol)
aqueous glycerol and stored in darkness at 
20°C. PE concentrations
were estimated on return to the laboratory with a Perkin-Elmer LS5
spectrofluorimeter and the instrument settings and correction
procedures previously reported (69).
RNA extraction and Northern analysis. Seawater samples (10 to 20 liters) were fractionated through 90-mm-diameter, 2.0- and 0.6-µm-pore-size polycarbonate filters (Poretics Corp.) at negative vacuum pressures of 10 and 60 cm of Hg for the 2.0- and 0.6-µm-pore-size filters, respectively. Routinely, this fractionation procedure results in the retention of greater than 90% of the original Synechococcus sp. biomass on the 0.6-µm-pore-size filter whereas eukaryotic cells are either trapped by the 2.0-µm-pore-size filter or disrupted by the high vacuum pressure employed during filtration through the 0.6-µm-pore-size filter. The efficiency of the procedure was monitored by epifluorescence microscopic examination of the cell material retained on the 0.6-µm-pore-size filter. In all cases, the only intact chlorophyll-containing cells observed also exhibited PE fluorescence; i.e., only Synechococcus sp. cells were present in the 0.6- to 2.0-µm size-fractionated samples under the conditions applied.
Sample volume was determined by the amount of seawater that could be filter fractioned within 15 min of sample collection, and all nocturnal samples were processed in a darkened laboratory. Synechococcus sp. cells retained on the 0.6-µm-pore-size filter were washed with 5 ml of TEN buffer (10 mM Tris-HCl [pH 8.0], 1 mM EDTA, 250 mM NaCl), resuspended in 1 ml of ice-cold extraction buffer (100 mM LiCl, 50 mM Tris-HCl [pH 7.5], 1 mM EGTA, 1% [wt/vol] sodium dodecyl sulfate [SDS]), and snap frozen prior to storage at20°C until further processing ashore.
Total RNA was isolated by a hot-phenol extraction procedure
(70). Synechococcus sp. cells were homogenized in
extraction buffer at 65°C for 1 to 2 min and deproteinized with an
equal volume of hot (65°C) phenol-chloroform-isoamyl alcohol
(25:24:1) for 5 min. The aqueous phase was recovered by centrifugation
(13,000 × g for 5 min) and reextracted with
chloroform-isoamyl alcohol (24:1) at ambient temperature. Nucleic acids
were precipitated from the aqueous phase with 2.5 volumes of 100%
ethanol at 20°C for 24 h and recovered by centrifugation
(13,000 × g for 20 min). The pelleted material was
washed in 75% (vol/vol) ethanol and taken up in 100 µl of DNase
buffer (100 mM sodium acetate, 10 mM MgCl2). DNA was
hydrolyzed at 37°C for 30 min in the presence of 10 U of DNase (RNase
free; Boehringer Mannheim). Following inactivation and removal of DNase
by phenol-chloroform extraction, RNA was pelleted by ethanol
precipitation and taken up in 50 µl of diethylpyrocarbonate-treated
deionized water. RNA concentrations were determined by absorbance at
260 nm, and the integrity of the samples was confirmed by
electrophoresis through formaldehyde agarose gels stained with ethidium
bromide (3).
Northern dot blots were performed as described previously
(70) following heat denaturation of samples (1 to 2 µg of
RNA) in 200 µl of 5× SSC (1× SSC is 0.15 M NaCl, 15 mM Na citrate, pH 7.5)-10 mM EDTA at 65°C for 15 min. Following transfer under gentle vacuum (<4 cm of Hg) to positively charged nylon membranes (Boehringer Mannheim), the blots were air dried and the RNA was immobilized by UV irradiation at 305 nm.
Dot blots were hybridized at high stringency with double-stranded DNA
probes derived from the oceanic cyanobacterium Synechococcus sp. strain WH8103 (68, 71). The probes employed were (i) a 699-bp BamHI-BglII fragment of pRBGL1.4, which
includes the 5' region and the first 630 nucleotides of
rbcL; (ii) an EcoRI-SalI fragment of
pPE1.3, which includes 198 bp of upstream sequence, the coding region
of cpeB, and the 5' end of cpeA of the class 1 PE
operon; and (iii) the EcoRI insert of pGlnA1.1, which
includes most of the coding region of glnA plus 350 bp of
upstream sequence. Probe DNA was isolated in low-melting-point agarose
following restriction digestion of plasmid DNA, and the recovered
fragments were purified with glass milk. DNA was labelled with
digoxigenin-dUTP (Boehringer Mannheim) by random priming according to
the manufacturer's recommendations.
Membranes were prehybridized for 4 h at 55°C in a solution
containing 50% (vol/vol) formamide, 5× SSC, 0.1% (wt/vol) SDS, 1%
(wt/vol) blocking reagent (Boehringer Mannheim), and 5% (wt/vol) dextran sulfate. Probe DNA (25-ng/ml final concentration) and sheared
salmon sperm DNA (100-µg/ml final concentration) were denatured at
100°C for 10 min and added to fresh solution prior to overnight
hybridization at 55°C.
Northern dot blots were stringency washed twice in 2× SSC-0.1%
(wt/vol) SDS at ambient temperature for 5 min and twice again at 65°C
in 0.2× SSC-0.1% (wt/vol) SDS for 30 min. Hybrids were detected by
immunochemistry with alkaline phosphatase-conjugated anti-digoxigenin
and the chemiluminescent substrate AMPPD (Tropix Inc.) in conjunction
with Amersham Hybond enhanced chemiluminescence film (70).
The resulting luminographs were quantified by densitometry with a GDS
8000 gel documentation system (UVP Ltd.) and the Gel Works 1D Advanced
software package.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Filter fractionation of Synechococcus spp. An essential requirement of several of the analyses carried out during the present study was the development of a rapid isolation procedure to concentrate Synechococcus sp. cells from bulk phytoplankton samples. The differential-fractionation technique employed was a minor refinement of an earlier protocol (69) that had been field tested during previous cruises in the northeast Atlantic Ocean (RRS Discovery cruise D189 and RRS Charles Darwin cruise CD47) and also by others for phytoplankton communities sampled off the North Carolina coast (25). In confirmation of the absence of eukaryotic algae from the fractionated samples prepared for the Northern analyses, only 16S and 23S rRNA bands were detected on denaturing RNA gels (Fig. 2) plus an additional band of intermediate molecular mass which is a diagnostic degradation product of the 23S rRNA subunit found in cyanobacterial RNA samples extracted in buffers containing low concentrations of Mg2+ ions (25, 48).
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Temporal variability in Synechococcus sp. abundance and
PE content.
Within the eddy, Synechococcus sp. cell
concentrations were elevated (~2-fold) in comparison to those of
surrounding waters and showed a net increase of 3- to 4-fold between
the first and last days on station (Table
1). During this period, the sea surface temperature increased by ~1°C, leading to the development of some weak secondary structure in the upper 10 m of the water column toward the end of the study (Table 1 and Figure
3, top). Nitrate concentrations within
the upper mixed layer were within the range of 0.6 to 2.1 µM, whereas
ammonium concentrations increased with depth from the surface (<0.1
µM) to reach 0.15 to 0.2 µM near the base of the mixed layer
(5).
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Diel variability in Synechococcus mRNA abundance. Changes in the relative abundance of three Synechococcus sp. mRNAs were measured over the course of two diel cycles (17 to 18 and 19 to 20 July 1991) toward the end of the period on station. Comparatively little temporal variability in cpeBA mRNA levels was observed; transcript levels oscillated somewhat (dynamic range, ~2), but no consistent evidence of a distinct temporal pattern was evident during either diel cycle (Fig. 4a and b). By contrast, marked temporal variability in rbcL and glnA expression occurred and resulted in about an order of magnitude difference in mRNA levels between the respective daily maxima and minima in the abundance of both transcripts (Fig. 4c to f).
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Diel patterns of Synechococcus sp. cell division and PE synthesis. The entrainment of Synechococcus sp. cell division to a particular phase of the diel cycle (Fig. 3b) has been documented previously for both coastal and open waters (7, 8, 42, 56, 62). In agreement with the observations reported here, these earlier studies indicate that oceanic populations most frequently undergo division during the late evening or night. It is intriguing, therefore, that cell division in rapidly growing coastal communities is initiated somewhat earlier in the diel cycle (8, 62). While such contrasting behavior may point to fundamental differences in cell cycle control between coastal and oceanic Synechococcus spp., the few seasonal records available provide some evidence that the peak in the frequency of dividing cells is shifted to a later phase in the diel cycle in coastal waters during the autumn and winter months (8, 62). Perhaps under these circumstances, in situ growth rate (one or fewer divisions per day) and environmental constraints on the diel phasing of the cell cycle more closely mirror those typical of Synechococcus sp. populations growing in open waters under presumably less favorable physicochemical conditions. Such a contention is certainly supported by the observation that phosphate additions to surface seawater samples from the Mediterranean Sea advanced subsequent progression through the Synechococcus sp. cell cycle by several hours (56).
Two distinct temporal patterns of DNA synthesis have been described for Synechococcus spp. growing in laboratory culture under alternating light-dark cycles (2, 4), but neither pattern is consistent with the development of true cell cycle synchrony. In both coastal and oceanic isolates a significant proportion of cells enter the light cycle in G2 having already completed progression through S during the latter half of the previous photoperiod. In the natural populations studied by Vaulot and colleagues, by contrast, all Synechococcus sp. cells entered the photoperiod in G1 having progressed through G2/M during the previous evening and night (56). Similar significant differences between the cell cycle behavior of laboratory cultures and that of natural populations have also been reported for the related photosynthetic prokaryote Prochlorococcus marinus. Progression through the cell cycle is highly synchronized to the diel periodicity in irradiance in natural populations of P. marinus but, as with Synechococcus spp., this behavior is not reproduced under standard laboratory conditions (55, 56). The apparent discrepancy between laboratory models of Synechococcus sp. cell division and field observations remains unexplained but might be reconciled by examining the quantum requirement for passage through the light-dependent cell cycle checkpoints in G1 and G2 originally proposed by Chisolm and coworkers (2, 4). If the quantum requirement for progression through G1 into S phase is significantly higher than that required for progression through G2/M, then under natural illumination (in which, as Vaulot et al. [56] point out, incident irradiance shows a sinusoidal temporal distribution) cells may become stalled in G1 during the last few hours of fading daylight, whereas the dark block in G2 might still be passed. Quite clearly, such a scenario could lead to the rapid establishment of cell cycle synchrony under natural illumination because, irrespective of when division took place during the latter half the previous diel cycle, all cells should enter the next diurnal period in G1. It is not an absolute requirement of this model to propose the existence of two distinct light control circuits to regulate the G1 and G2 checkpoints in order to promote cell cycle synchrony under natural illumination. Newly born cells in G1 have approximately half the pigment complement and, presumably, half the light absorption capacity of G2 cells that are just about to undergo division (Fig. 3, bottom). In addition to the diel periodicity in DNA content that results from cell cycle synchronization (56), the entrainment of Synechococcus sp. cell division to the natural photoperiod predicts a number of other cell cycle-related effects on the daily pattern of macromolecular synthesis. The most obvious of these is the exponential increase in stable RNA (rRNA and tRNA) and protein content that must accompany progression through the cell cycle. Bulk measurements of biosynthetic processes performed on natural populations are unlikely to resolve such temporal changes in cell composition, however, unless they are corrected for (or are independent of) loss terms such as grazing and advection as well as the influence of other members of the planktonic community. Nevertheless, where specific estimates of Synechococcus sp. cell composition have been attempted with natural populations the expected temporal pattern of macromolecular synthesis has emerged (20, 25). As a further case in point, the cell content of the biliprotein PE was measured directly during the present study and found to increase in a quasilogarithmic fashion throughout the daylight hours (Fig. 3, bottom). Assuming that the Synechococcus sp. population within the eddy was in balanced growth (a not-unreasonable assumption given the comparative lack of variability in the environment), this suggests a more or less constant rate of synthesis (relative to the general protein pool) during all phases of the cell cycle. Perhaps as a consequence of the comparatively short duration of the nocturnal period, the decline in cell PE content after dusk could be largely explained by the effect of dilution through cell division rather than by a nighttime decrease in the net rate of synthesis. Periodicity in energy supply need not in fact lead to very large variations in the rate of protein synthesis, at least not in cyanobacteria grown in light-dark cycles with comparatively long light periods (41). At lower latitudes, however, the nocturnal rate of PE synthesis in marine Synechococcus spp. does appear to be somewhat lower even after allowance for dilution effects following cell division (68). There is also some evidence that the rate of Synechococcus sp. ribosome synthesis declines during the rather longer hours of darkness experienced at more southerly locations during the summer months (25).Temporal variation in gene expression. Consistent with the diel pattern of PE synthesis, the relative abundance of Synechococcus sp. cpeBA mRNA showed comparatively little temporal variability when normalized, as in the present case, to total RNA. Since cell ribosome content must also be a temporal variable in synchronously dividing populations, the comparative lack of variability in the normalized abundance of cpeBA mRNA suggests that either the PE operon was expressed constitutively or the cpeBA message is unusually stable. In fact, the half-life of cpeBA mRNA in the marine cyanobacterium Synechococcus sp. strain WH7803 is of the same order (10 to 15 min) as those determined for several other genes (9, 28, 68), and there is no a priori reason to believe that the natural population should differ markedly in this regard.
The pronounced diel variability and periodicity in Synechococcus sp. rbcL and glnA mRNAs, by contrast, is much more consistent with the operation of specific controls at the transcriptional and/or posttranscriptional levels. Although some of the temporal variability in mRNA abundance may reflect global changes in transcription rates as the population progressed through the cell cycle, such effects are perhaps as likely to influence the overall abundance of transcripts from the PE operon as they are to influence the diel pattern of RubisCO or GS expression. It is significant, therefore, that the temporal periodicities in Synechococcus sp. rbcL and glnA mRNAs remain markedly conservative features of the natural population when each is normalized to cpeBA mRNA (Fig. 5). Irrespective of whether the PE operon was truly constitutively expressed or not, it is clearly apparent that rbcL and glnA were differentially regulated during the diel cycle, both with respect to each other and with respect to cpeBA.
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ACKNOWLEDGMENTS |
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This work was supported by research grants from the Natural Environment Research Council (NERC), the Joint Environment Programme of National Power and Powergen, and the University of Stirling.
I acknowledge access to ship time during the NERC Biogeochemical Ocean Flux Study and the encouragement and support of R. P. Harris, M. Whitfield, and others at Plymouth Marine Laboratory and the Marine Biological Association.
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FOOTNOTES |
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* Mailing address: Department of Biological Sciences, University of Stirling, Stirling FK9 4LA, United Kingdom. Phone: 44 1786 467784. Fax: 44 1786 464994. E-mail: michael.wyman{at}stir.ac.uk.
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REFERENCES |
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
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