Download Impaired glycogen synthesis causes metabolic

Microbiology (2012), 158, 3032–3043
DOI 10.1099/mic.0.062950-0
Impaired glycogen synthesis causes metabolic
overflow reactions and affects stress responses in
the cyanobacterium Synechocystis sp. PCC 6803
Marianne Gründel, Ramon Scheunemann, Wolfgang Lockau
and Yvonne Zilliges
Correspondence
Yvonne Zilliges
Humboldt-Universität zu Berlin, Institute of Biology/Biochemistry of Plants, Chausseestr. 117,
10115 Berlin, Germany
[email protected].
de
Received 17 August 2012
Revised
2 October 2012
Accepted 3 October 2012
The biosynthesis of glycogen or starch is one of the main strategies developed by living organisms
for the intracellular storage of carbon and energy. In phototrophic organisms, such polyglucans
accumulate due to carbon fixation during photosynthesis and are used to provide maintenance
energy for cell integrity, function and viability in dark periods. Moreover, it is assumed that
glycogen enables cyanobacteria to cope with transient starvation conditions, as observed in most
micro-organisms. Here, glycogen accumulates when an appropriate carbon source is available in
sufficient amounts but growth is inhibited by lack of other nutrients. In this study, the role of
glycogen in energy and carbon metabolism of phototrophic cyanobacteria was first analysed via a
comparative physiological and metabolic characterization of knockout mutants defective in
glycogen synthesis. We first proved the role of glycogen as a respiratory substrate in periods of
darkness, the role of glycogen as a reserve to survive starvation periods such as nitrogen
depletion and the role of glycogen synthesis as an ameliorator of carbon excess conditions in the
model organism Synechocystis sp. PCC 6803. We provide striking new insights into the complex
carbon and nitrogen metabolism of non-diazotrophic cyanobacteria: a phenotype of sensitivity to
photomixotrophic conditions and of reduced glucose uptake, a non-bleaching phenotype based
on an impaired acclimation response to nitrogen depletion and furthermore a phenotype of energy
spilling. This study shows that the analysis of deficiencies in glycogen metabolism is a valuable
tool for the identification of metabolic regulatory principles and signals.
INTRODUCTION
Glycogen is a highly branched polymer of glucose which
serves as a carbon and energy reserve in most organisms
(Preiss & Romeo, 1994). The synthesis of glycogen in
cyanobacteria and of starch in plants and algae starts with
the use of ADP-glucose as the glycosyl donor (Ball &
Morell, 2003). The synthesis of ADP-glucose from glucose1-phosphate and ATP, catalysed by ADP-glucose pyrophosphorylase (AGPase), also referred to as glucose-1-phosphate adenylyltransferase, defines both the first committed
step of glycogen synthesis and the major rate-controlling
step of glycogen synthesis. AGPase is finely tuned through
allosteric activation by 3-phosphoglycerate, the product of
the carboxylating reaction of ribulose-1,5-bisphosphate
carboxylase/oxygenase, and is inhibited by orthophosphate.
This regulation of AGPase in cyanobacteria most likely
evolved to couple storage polysaccharide synthesis to
photosynthetic CO2 fixation, and was conserved during
chloroplast evolution for starch biosynthesis in plants
Abbreviation: AGPase, ADP glucose pyrophosphorylase.
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(Ball & Morell, 2003). The glucose moiety of ADP-glucose
is transferred to the non-reducing end of a linear a-1,4
glucane primer, a reaction catalysed by glycogen synthases.
The a-1,6-glycosidic branches are introduced into the
growing polymer by the branching enzyme (Preiss &
Romeo, 1994).
In many micro-organisms, glycogen plays a major role in
the survival under adverse conditions. Accumulation of
glycogen is enhanced when a macronutrient, e.g. a nitrogen
or sulfur source, is limiting but a carbon source is present
in sufficient amounts. Various bacterial species accumulate
glycogen in the stationary phase, when growth stops
because of either depletion of a nutrient, such as nitrogen,
sulfur or phosphate, or an unfavourable pH (Preiss, 1984).
Glycogen enables the cells to cope with transient starvation
conditions that may result in a significant loss of viability
in glycogen-deficient strains (Preiss, 1984). Several studies
have associated bacterial glycogen metabolism with environmental survival, symbiotic performance, colonization and
virulence, but the exact roles of the polymer may differ from
strain to strain (Wilson et al., 2010).
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Cyanobacterial energy and carbon metabolism
In cyanobacteria, the ability for glycogen synthesis appears
to be conserved since the respective genes are present in all
cyanobacterial genomes annotated so far (Beck et al., 2012).
However, the number of isoenzymes involved in glycogen
biosynthesis and degradation varies between cyanobacterial
species. It is thought that cyanobacteria store a portion of
the photosynthetically generated carbohydrate as glycogen,
which is assumed to be the substrate of endogenous
metabolism during periods of darkness (Stanier & CohenBazire, 1977). It has been hypothesized that glycogen acts as
a dynamic reserve in cyanobacteria with the dual function as
a reserve and a buffer substance (Carr, 1988).
One way to understand the function of glycogen in
phototrophic cyanobacteria is to generate a biological
system that lacks genes that are involved in glycogen
synthesis. First knockout mutagenesis of AGPase and
glycogen synthase in Synechococcus elongatus PCC 7942
and of AGPase in Synechocystis sp. PCC 6803 revealed a
role for AGPase and glycogen in acclimation and tolerance
to salt and oxidative stress (Miao et al., 2003a; Suzuki et al.,
2010). However, important aspects of the role of glycogen
metabolism in central carbon metabolism and in acclimation processes to macronutrient changes have not yet
been investigated. In this study, we address (i) the role of
glycogen as a respiratory substrate in periods of darkness,
(ii) the role of glycogen as a reserve to survive starvation
periods such as nitrogen depletion and (iii) the role of
glycogen synthesis as an ameliorator of carbon excess via a
comparative physiological and metabolic analysis of
knockout mutants defective in AGPase or in both glycogen
synthases in the model organism Synechocystis sp. PCC
6803 (hereafter referred to as Synechocystis).
METHODS
Bacterial strains and growth conditions. The strains of
Synechocystis used in this study were propagated on BG11 agar plates
(Rippka et al., 1979). Liquid cultures of Synechocystis wild-type and
mutant strains were grown at 28 uC under continuous illumination
with white light in a range of 60–100 mmol photons m22 s21 (as
indicated in the respective experiments) and continuous aeration
(0.03 %, v/v, CO2) in BG11 medium containing 20 mM HEPES buffer
(pH 8.0) and 17.6 mM sodium nitrate as nitrogen source (+N), or in
BG110, which lacks sodium nitrate (2N). Nitrogen deprivation was
achieved by a repeated washing–centrifugation procedure and
resuspension in BG110 medium. Cultures were split and nitrate was
added again to one of them (control). Photomixotrophic growth was
achieved by the addition of glucose in a final concentration of 5 mM to
agar plates and liquid cultures, respectively. A light–dark cycle was
usually (or as indicated) performed with synchronized cells at
alternating light and dark periods of 16 (100 mmol photons m22 s21)
and 8 h, respectively. For strain maintenance and pre-culturing of the
mutants, antibiotics were used at the following concentrations: 14 mg
chloramphenicol ml21; 75 mg kanamycin ml21. Escherichia coli strain
DH5a was used for all plasmid constructions. E. coli cultures were
grown in LB medium, supplemented with antibiotics at standard
concentrations as appropriate.
Plasmids and mutagenesis. For insertion mutagenesis, the glgC
ORF (locus slr1176), the glgA1 ORF (locus sll0945) and the glgA2 ORF
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(locus sll1393) were amplified with #glgC.fw (59-GTGTGTTGTTGGCAATCGAG) and #glgC.rv (59-GCCCAGTCCTAGGGGTAAAA), with
#glgA1.fw (59-CGACGGTATGAAGCTTTTATTTG) and #glgA1.rv (59CCGGCGGAACGGTACCAAC) and with #glgA2.fw (59-GGCCAGGGGAATTCTCCTCCAG) and #glgA2.rv (59-GCGGATAATACTGAACGAAGCTTTG), ligated into the pJET1.2 vector (Fermentas) or pUC19
(New England Biolabs) and disrupted by deletion–insertion mutagenesis (for glgC) or insertion mutagenesis (for glgA1 and glgA2) of a
chloramphenicol resistance cassette from pACYC184 (New England
Biolabs) or a kanamycin resistance cassette from pUC4K (GE
Healthcare) in sense as well as in antisense orientation to the respective
gene in order to check for polar effects. The constructs were used
to transform Synechocystis as described by Ermakova et al. (1993).
Transformants were restreaked six times and analysed by Southern blot
hybridization to detect the level of segregation of genome copies.
Southern blot analysis. Genomic DNA of Synechocystis was
extracted as described by Franche & Damerval (1988). After digestion
of DNA with the restriction endonucleases NcoI, XbaI, SmaI and
EcoRI (Fermentas), electrophoresis and gel blotting to nylon
membranes (Amersham Hybond-N, GE Healthcare) were performed.
DNA probe fragments, obtained by PCR for glgC with #glgC150.fw
(59-TCCCGTCAGTAATTGCATCA) and #glgC330.rv (59-AGTGCCCTGAAACCAATCAG) and from restriction with NcoI and XbaI of
pUC19/glgA1 and pUC19/glgA2, respectively, were labelled with
[a-32P]dCTP with the DecaLabel DNA labelling kit (Fermentas) as
recommended by the manufacturer. DNA:DNA hybridizations
were performed according to standard protocols (Sambrook et al.,
1989).
Analysis of growth, pigmentation, photosynthetic capacity and
cell size. Growth characteristics of the cultures during exponential
growth (OD750 0.2–1.5) were analysed by daily measurement of
OD750 and of the chlorophyll content (Tandeau de Marsac &
Houmard, 1988). Whole cell absorbance spectra were recorded in the
wavelength range from 550 to 750 nm on a Shimadzu UV-2401PC
spectrophotometer equipped with an integrating sphere and were
corrected for residual cell scattering at 750 nm and normalized to the
chlorophyll a peak at 683 nm. Cultures used for the estimation of
growth rates did not contain antibiotics to reduce side effects. The
growth rate was calculated in the exponential growth phase as the
linear regression of the natural logarithm of chlorophyll content [mg
chlorophyll (ml cell suspension)21] verus time. Photosynthetic
capacity of cell suspensions containing 15 mg chlorophyll ml21 was
determined; the suspensions were washed and resuspended in BG11
with 25 mM NaHCO3. Oxygen evolution rates were measured with a
Clark-type (DW1) oxygen electrode (Hansatech) at 25 uC illuminated
with white light (intensity of 300 mmol photons m22 s21). Particle
size distribution was measured with the laser diffraction particle
analyser ANALYSETTE 22 (Fritsch). Cell concentration was determined
by counting in a Neubauer counting chamber.
Viability test. For spot assay experiments, cells were grown to
OD750 0.5–0.7 and adjusted to a chlorophyll content of 5 mg ml21 at
the beginning of the experiment. Aliquots (7 ml) of three different
dilutions (1 : 1, 1 : 10, 1 : 100) were dropped on BG11 agar plates; the
plates were incubated at 28 uC under continuous or alternating (12 h
light/12 h dark) illumination (60 mmol photons m22 s21) for 7 days.
Determination of glycogen content. For isolation of glycogen,
cell pellets of 1–3 ml culture were washed three times with sterile
water, suspended in 30 % (w/v) KOH and incubated at 95 uC for 2 h.
Precipitation of glycogen was achieved by the addition of ice-cold
ethanol to a final concentration of 70–75 % (v/v) and subsequent
incubation on ice for at least 2 h. After a centrifugation step (10 min,
4 uC, 10 000 g) the glycogen pellet was washed twice with 70 and 98 %
(v/v) ethanol, and dried in a speed-vac (10 min, 60 uC). The isolated
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M. Gründel and others
2
gC
Δg
l
Δg
l
W
(a)
T
gA
Determination of extracellular metabolites. For analysis of
extracellular metabolites the filtered (costar Spin-X 0.22 mm Nylon
PP Tube; Corning) supernatants of cell suspensions were directly
analysed by ion exclusion chromatography using a Shim-pack SCR102H column and Guard Column SCR-102H (Shimadzu) with 4 mM
p-toluenesulfonic acid aqueous solution as mobile phase, a flow rate of
0.8 ml min21 and a column temperature of 46 uC. Electroconductivity
of organic acids was analysed by the post-column pH-buffered addition
of 16 mM Bistris, 100 mM EDTA in 4 mM p-toluenesulfonic acid
aqueous solution. Identification of the respective peaks and quantification via calibration curves was achieved using commercial
standards (Sigma Aldrich). Additionally, optical enzymic tests were
performed for quantification of extracellular pyruvate (Lamprecht &
Heinz, 1984) and 2-oxoglutarate (Senior, 1975). Both methods gave
essentially identical results. Glutamate was quantified in a colorimetric
assay using glutamate dehydrogenase, NAD, iodonitrotetrazolium and
phenazine methosulfate (500 nm).
gA
(Hexokinase method) from Thermo Fisher Scientific.
Mutants unable to synthesize glycogen were not impaired
in their photoautotrophic growth. On agar plates (Fig. 1a)
as well as in liquid cultures (Fig. 2a) a similar growth
behaviour was observed for the wild-type and all knockout
mutants. Growth rates of 0.32 mg chlorophyll (ml cell
suspension)21 per day for wild-type and 0.30 mg chlorophyll
(ml cell suspension)21 per day for the mutants were
measured under constant illumination with 40 mmol
photons m22 s21 and constant aeration with 0.03 % (v/v)
CO2 (Fig. 2a). Even though no significant growth differences
were detectable, obvious phenotypes of glycogen-deficient
mutants (DglgA1/DglgA2, DglgC) were observed under strict
photoautotrophic conditions: (i) elevated sensitivity to high
light intensities (not shown) and to low inocula (Fig. 1a) and
(ii) significant reduction in photosynthetic capacity. Oxygen
evolution rates of 212±8 (SD) [mmol O2 h21 (mg
gl
Quantification of extracellular glucose. The extracellular glucose
content was determined in 10 ml medium with the glucose reagent
Reduced photosynthetic capacity and high-lightsensitive phenotype
1 /Δ
glycogen was resupended in 100 mM sodium acetate (pH 4.5)
and enzymically hydrolysed to glucose by treatment with 2 mg
amyloglucosidase ml21 (Roche Diagnostics) at 60 uC for 2 h. Glycogen
content was determined with the glucose reagent (Hexokinase method)
from Thermo Fisher Scientific according to the manufacturer’s
instructions.
RESULTS
Genes responsible for glycogen synthesis in cyanobacteria
have been identified by their similarity to the respective
homologues for AGPase and glycogen synthase from E.
coli, plants and green algae. The genome of Synechocystis
encodes one AGPase homologue (glgC: slr1176) and two
glycogen synthase homologues (glgA: sll0945, sll1393)
(Nakao et al., 2010). ORF sll0945 encodes a bacterial-type
glycogen synthase, homologues of which are present in all
cyanobacteria, subsequently referred to as glgA1 (Cid et al.,
2002). The additional (starch-type) glycogen synthase
(sll1393) is referred as glgA2, according to the annotated
homologue in Synechococcus sp. PCC 7002.
Complete inhibition of glycogen synthesis was achieved by
mutagenesis of either AGPase (DglgC) or simultaneous
inactivation of both glycogen synthases (DglgA1/DglgA2),
respectively. The knockouts were verified by Southern blot
analyses (not shown). The inability of the mutants to
synthesize glycogen was checked after nitrogen depletion,
conditions that trigger massive glycogen accumulation in
wild-type cells as a short-term response of the general stress
acclimation process (see Fig. 4a below). By contrast, single
knockout mutants of glycogen synthases, DglgA1 and DglgA2,
respectively, were still able to accumulate similar amounts of
glycogen as the wild-type (not shown). Consequently, either
the double knockout mutant DglgA1/DglgA2 or the single
knockout mutant DglgC was used to analyse the impact of
incapacities in glycogen synthesis on the cellular stress
responses and on the carbon and nitrogen metabolism.
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1:10
1:100
(b)
1:1
Light–dark cycle
Knockout of glycogen synthesis
Continuous light
1:1
1:10
1:100
Fig. 1. Growth and viability under light–dark cycle conditions.
Growth of wild-type (WT) and mutants defective in glycogen
synthesis (DglgA1/DglgA2; DglgC) under continuous (a) and light
(12 h)–dark (12 h) cycle conditions (b). Strains in the exponential growth phase with a chlorophyll content of 5 mg ml”1 were
plated at different dilutions [1 : 1 (undiluted), 1 : 10, 1 : 100] on
BG11 agar, pH 8.0 and incubated under continuously (a) or
alternated (b) illumination of 60 mmol photons m”2 s”1 at 28 6C
for 7 days. The effect was verified by three independent spot
assays.
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Microbiology 158
Cyanobacterial energy and carbon metabolism
Impairment of viability under stress conditions
Photoautotrophic growth
(a)
Drastic phenotypes occurred when cells were stressed by
varying conditions such as alternating light–dark periods
(A), photomixotrophic growth in the presence of CO2 and
glucose (B), and nitrogen starvation (C).
Chlorophyll (mg ml–1)
30
25
20
(A) Restricted viability under alternating light–dark
periods. Synchronized cells of Synechocystis (wild-type)
accumulated glycogen in minor levels of 2.0–2.3 mg glucose
equivalents (mg chlorophyll)21 during the 16 h light period
(140 mmol photons m22 s21) and dissipated barely half
of it to levels of 1.2–1.3 mg glucose equivalents (mg
15
10
5
0
0
20
40
60
80
100
120
140
Time (h)
Chlorophyll (mg ml–1)
30
6
25
5
20
4
15
3
10
2
5
1
0
0
0
20
40
60
80
100
120
Glucose (mmol l–1)
Photomixotrophic growth
(b)
140
Time (h)
WT
ΔglgA1/ΔglgA2
WT
ΔglgA1/ΔglgA2
ΔglgC
ΔglgC
Fig. 2. Growth and viability under photomixotrophic conditions.
Growth of wild-type (WT) and mutants defective in glycogen
synthesis (DglgA1/DglgA2; DglgC) in batch cultures in the
absence (a) or presence (b) of glucose (Glc). Glucose was added
at 5 mM (b). All strains were incubated under continuous
illumination of 60 mmol photons m”2 s”1 at 28 6C for 6 days.
Growth curves (solid lines) were calculated by the diurnal
determined chlorophyll content [mg chlorophyll (ml cell suspension)”1] of respective cultures (first y-axis), showing mean±SD
from three independent experiments. Glucose consumption
(dotted lines) was calculated by the diurnal determined glucose
levels [mmol glucose (l supernatant)”1] of respective supernatants (second y-axis) show mean±SD from three independent
experiments.
chlorophyll)21] in the wild-type and of 174±7 (SD) [mmol
O2 h21 (mg chlorophyll)21] in DglgA1/DglgA2 and DglgC
mutants were determined under saturating illumination
with white light.
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chlorophyll)21 during the 8 h dark period. The growth
rate (wild-type) was slightly reduced to 0.22 mg chlorophyll
(ml cell suspension)21 per day compared with strict photoautotrophic conditions. In contrast, cells incapable of
glycogen synthesis (DglgA1/DglgA2, DglgC) were restricted
in their viability under this light–dark regime (Fig. 1b). The
loss in viability became most evident in spot assays where
a minimal inoculum of cells has been tested. Based on
identical inocula, the colour intensity of the colony spots
differed significantly between wild-type and mutants after
only a few days of growth. The wild-type cells formed deep
blue–green colonies even from highly diluted inocula (Fig.
1b, 1 : 100). Much larger inocula of mutant cells were
required to obtain colonies of similar cell density (Fig. 1b,
1 : 1).
(B) Photomixotrophy-sensitive phenotype. Synechocystis
is capable of growing photomixotrophically and uses both
CO2 and glucose as carbon source in parallel under these
conditions. In this case, it is assumed that the synthesized
glycogen is a product not only of photosynthesis but also of
glucose conversion (Smith, 1982). Under the conditions
tested, glycogen levels of 1.0–2.0 mg glucose equivalents
(mg chlorophyll)21 were determined in the presence of
5 mM glucose and 0.03 % (v/v) CO2, similar to the value
under strictly photoautotrophic conditions (see Fig. 4d).
Simultaneous assimilation of CO2 and glucose led to an
enhanced exponential growth rate of 0.38 mg chlorophyll
(ml cell suspension)21 per day in wild-type (Fig. 2b),
whereas the growth rates of both mutants (DglgA1/ DglgA2,
DglgC) were reduced to 0.20 mg chlorophyll (ml cell
suspension)21 per day under identical conditions (Fig.
2b). Moreover, the glucose uptake rates differed between
wild-type and glycogen-deficient mutants. Based on the
addition of 5 mM glucose to each culture, glucose levels
were diminished to half within 50 h in wild-type cultures,
whereas in cultures of mutants impaired in glycogen
synthesis this level was not reached before 100 h (Fig. 2b).
On agar plates containing glucose, cells with impaired
glycogen synthesis were severely restricted in their viability
and were unable to resume growth even after several days
(not shown).
(C) Loss of viability under nitrogen starvation. Nitrogen
depletion strongly promoted glycogen accumulation in
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M. Gründel and others
wild-type cells. Here, glycogen levels of 67.5±3.7 mg
glucose equivalents (mg chlorophyll)21 were reached
under light intensities of 100 mmol photons m22 s21
after 48 h of nitrogen deprivation (Fig. 4a below). Longterm nitrogen depletion led inevitably to a loss of viability
for mutants that lacked the capacity for glycogen synthesis
(DglgA1/DglgA2, DglgC). In spot assays, cells of cultures
starved for nitrogen for up to 7 days were transferred to
nitrogen-containing agar plates (Fig. 3). The wild-type cells
formed blue–green colonies even at high dilutions (Fig. 3b:
1 : 100) and even after a prolonged period of nitrogen
starvation (Fig. 3b, 7 days). In contrast, viability of mutant
cells (DglgA1/DglgA2, DglgC) was so strongly affected within
a few days of nitrogen depletion that regrowing was
impeded (Fig. 3b).
0.14–0.17 mM were measured in the culture supernatants
(Fig. 4b, c, white bars), similar to recently published results
(Carrieri et al., 2012). Glucose addition also led to an
accumulation of pyruvate to levels of 1.0–1.4 mM and of 2oxoglutarate to levels of 0.11–0.13 mM in the medium of
these mutant cultures within 48 h (Fig. 4e, f, white bars).
These energetically relevant metabolites were not detected
under either strictly photoautotrophic conditions in the
supernatants of glycogen-deficient mutants (Fig. 4, grey
bars) or the tested stress conditions in the supernatants of
wild-type cells (Fig. 4) or of the single knockout mutants
DglgA1 and DglgA2 (not shown). Both lysis and general
leakage effects could be excluded: first, no phycobiliproteins
were detectable in the supernatants by spectrophotometric
analysis. Second, the highly abundant intracellular amino
acid glutamate was not detected in any of the supernatants
(not shown).
‘Overflow metabolism’: release of pyruvate and 2oxoglutarate
Upon nitrogen starvation, wild-type cells did not accumulate glycogen when kept in darkness, and not even
traces of energetically relevant metabolites were detectable
in the supernatants of glycogen-deficient mutants under
this condition. However, glycogen accumulation in wildtype cells as well as metabolite release from cells incapable
of glycogen synthesis was detectable about 5 h after
switching on the light (not shown). The release of pyruvate
and 2-oxoglutarate by the glycogen-deficient mutants is
thus a light-dependent process, like glycogen accumulation
in wild-type cells.
gA
Δg 1 /Δ
lg glg
C
T
Δg
l
W
W
T
Δg
l
gA
Δg 1 /Δ
lg glg
C
A
A
2
(a)
2
To prove the role of glycogen synthesis as an ameliorator
of carbon excess, metabolite profiles of culture supernatants of the wild-type and of the knockout mutants
incapable of glycogen synthesis (DglgA1/DglgA2, DglgC)
were determined. Under conditions of nitrogen deprivation
or photomixotrophic growth, the mutant cells massively
lost energetically relevant metabolites such as pyruvate
and 2-oxoglutarate. After 48 h of nitrogen starvation,
pyruvate levels of 0.4–0.6 mM and 2-oxoglutarate levels of
-N
Length of nitrogen depletion (days)
0
1
2
4
7
(b)
1:1
+N
1:10
gA
Δg 2
lg
C
gl
W
T
gA
Δg 2
lg
C
1 /Δ
gA
Δg
l
Δg
l
gA
1 /Δ
gl
W
T
gA
Δg 2
lg
C
W
T
gl
1 /Δ
Δg
lg
A
gA
Δg 2
lg
C
gl
1 /Δ
2
C
Δg
lg
gA
gl
W
T
gA
Δg
l
Δg
l
gA
1 /Δ
W
T
1:100
Fig. 3. Viability test under nitrogen-depleted conditions. (a) The chlorotic process in cells of wild-type (WT) and of mutants
defective in glycogen synthesis (DglgA1/DglgA2; DglgC) in response to nitrogen starvation. (b) Viability test of nitrogen-starved
cells of wild-type (WT) and mutants defective in glycogen synthesis (DglgA1/ DglgA2; DglgC) under nitrogen-replenished
conditions. Differentially prolonged starved cells (0, 1, 2, 4, 7 days) were plated at different dilutions [1 : 1 (undiluted), 1 : 10,
1 : 100] on BG11 agar, pH 8.0 and incubated under continuous illumination of 60 mmol photons m”2 s”1 at 28 6C for 7 days.
The effect was verified by four independent spot assays.
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Microbiology 158
Cyanobacterial energy and carbon metabolism
Glycogen
Pyruvic acid
(b)
1.4
60
1.2
50
40
30
20
10
0
0
24
48
0
24
48
0
24
0.8
0.6
0.4
0.2
48
0
ΔglgC
ΔglgA1/ΔglgA2
24
48
0
24
48
0
24
1.2
50
40
30
20
10
0
24
48
WT
0
24
48
ΔglgA1/ΔglgA2
0
24
48
0.4
0.3
0.2
0,1
0.0
24
48
0
24
48
0
24
48
ΔglgC
ΔglgA1/ΔglgA2
WT
(f)
1.0
0.8
0.6
0.4
0.2
0.0
0
0.5
0
2-Oxoglutaric acid (mmol l–1)
60
Pyruvic acid (mmol l–1)
1.4
0.6
48
(e)
70
0.7
ΔglgC
ΔglgA1/ΔglgA2
WT
(d)
Glycogen [µg (µg chlorophyll)–1]
1.0
0.0
WT
+Glc
(c)
2-Oxoglutaric acid (mmol l–1)
70
Pyruvic acid (mmol l–1)
Glycogen [µg (µg chlorophyll)–1]
(a)
-N
2-Oxoglutaric acid
0
ΔglgC
24
WT
48
0
24
48
ΔglgA1/ΔglgA2
0
24
48
ΔglgC
0.7
0.6
0.5
0.4
0.3
0.2
0,1
0.0
0
24
WT
48
0
24
48
ΔglgA1/ΔglgA2
0
24
48
ΔglgC
Fig. 4. Metabolite profile analysis under conditions of carbon excess. Comparative analysis of intracellular glycogen levels (a, d)
and extracellular pyruvate (b, e) and 2-oxoglutarate (c, f) levels in wild-type (WT) and mutants defective in glycogen synthesis
(DglgA1/ DglgA2; DglgC) as short-term response (0, 24, 48 h) to carbon excess such as nitrogen starvation (a–c) and glucose
addition (d–f) (for respective growth curves see Fig. 2). Strains at exponential growth phase were split: one served as a control
(grey bars) and the other one (white bars) was treated as indicated. Data for nitrogen depletion (a–c) show mean±SD origin
from four different experiments and data from glucose feeding (d–f) show mean±SD from two different experiments. The
glycogen level is calculated from glucose equivalents (mg) related to chlorophyll content (mg). Pyruvate and 2-oxoglutarate levels
[mmol (l supernatant)”1] were determined via ion exclusion chromatography using standard calibration curves. Data were
completely coinciding by optic enzymic tests.
Impaired acclimation response upon nitrogen
depletion: the non-bleaching phenotype
Unicellular, non-diazotrophic cyanobacteria such as
Synechocystis undergo substantial changes in response to
nutrient starvation. Two characteristics are the completion
of cell division in the absence of further cell growth and the
degradation of the light-harvesting antennae, the phycobilisomes, by a proteolytic process termed chlorosis (Schwarz
& Forchhammer, 2005). Chlorosis becomes evident by a
bleaching process of the cells that results in a colour change
from blue–green to yellow–green (Fig. 3a, wild-type).
Knockout mutants that lack the ability for glycogen
synthesis were not able to degrade phycobilisomes. These
cells maintained their greenish-blue colouring despite
nitrogen depletion (Fig. 3a, DglgA1/DglgA2, DglgC). In whole
cell absorbance spectra, the phycocyanin peak (628 nm) was
unchanged in the DglgA1/DglgA2 and DglgC mutants (Fig. 5),
whereas in the wild-type (Fig. 5) and in the single knockout
mutants DglgA1 and DglgA2 (not shown) a distinct decline in
the phycocyanin peak was detectable within a few hours of
nitrogen depletion. Moreover, mutant cells incapable of
glycogen synthesis (DglgA1/DglgA2, DglgC) were not able to
divide after nitrogen deprivation. Total cell concentration as
well as the chlorophyll content per cell remained nearly
http://mic.sgmjournals.org
constant under these conditions. In contrast, wild-type cells
duplicated once whilst their chlorophyll content dropped
from about 8.0±0.661028 to 4.4±0.761028 mg per cell.
DISCUSSION
Knockout of glycogen synthesis in cyanobacteria
The ability to synthesize glycogen seems to be conserved in
cyanobacteria. The genes glgC (encoding the AGPase), glgA
(encoding the glycogen synthases) and glgB (encoding the
branching enzyme), which are involved in the synthesis of
the polymer, have been found in each of the so-far annotated
cyanobacterial genomes (Beck et al., 2012; Nakao et al.,
2010). In unicellular, non-diazotrophic cyanobacteria such
as Synechocystis sp. PCC 6803 (this study) and S. elongatus
PCC 7942 (Suzuki et al., 2010), glycogen synthesis was
abrogated by the independent knockout of either AGPase or
glycogen synthase(s). Neither glycogen (this study, Fig. 4)
nor the respective enzymic activities (Suzuki et al., 2010)
were detectable in cellular extracts of the mutants. Thus,
both enzyme activities are essential for glycogen synthesis,
and AGPase is the only source of ADP-glucose for glycogen
and probable glycosylglycerol biosynthesis in Synechocystis
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3037
M. Gründel and others
WT
ΔglgA1/ΔglgA2
ΔglgC
550
600
650
700
750
Wavelength (nm)
Fig. 5. Progression of phycobilisome degradation under conditions of nitrogen depletion. Progression of phycobilisome degradation (phycocyanin peak at 628 nm) was monitored by the analysis
of whole cell absorbance spectra of wild-type (WT), DglgA1/
DglgA2 and DglgC mutants after 0 h (solid line) and 24 h (dashed
line) of nitrogen depletion. After nitrogen step-down strains were
continuously illuminated with 100 mmol photons m”2 s”1. Spectra
were normalized to the chlorophyll peak (683 nm). The effect was
verified in 10 independent experiments.
(Hagemann & Erdmann, 1994). Manipulation of the
branching enzyme simply affects the structure of synthesized
glycogen (Yoo et al., 2002). Our knockout data (reported
here) show that the synthesis of ADP-glucose defines the
first committed step of glycogen synthesis in Synechocystis
similar to that observed in plants and heterotrophic bacteria
(Ball et al., 2011).
Starch-type enzymes in cyanobacteria and the
early loss of glycogen synthesis during
endosymbiosis
The ability for storage of carbon as glycogen, which is well
conserved in free-living cyanobacteria, was lost in the very
first steps of endosymbiosis (Nowack et al., 2008). All
recent Archaeplastida synthesize starch, a glucose polymer
that is structurally and physically different from glycogen
(Ball et al., 2011). Due to an increasing detection of starchlike polymers in present-day cyanobacteria and due to the
phylogenetic origin of the granule-bound starch synthase
from bacteria, it is assumed that ancient cyanobacteria
were capable of starch and/or glycogen synthesis. In line
with this, a few enzymes of the cyanobacterial polyglucan
(glycogen) metabolism clearly have a common origin with
enzymes involved in starch metabolism (Patron & Keeling,
2005). Synechocystis sp. PCC 6803 possesses two isoenzymes of polyglucan synthase: ORF sll0945 (glgA1) clusters
together with eubacterial glycosyltransferases and ORF
sll1393 (glgA2) clusters together with plant and algal soluble
(not granule bound) starch synthases (Cid et al., 2002;
Patron & Keeling, 2005). Both types of synthases are
3038
involved in the polyglucan synthesis in Synechocystis, as
demonstrated by the fact that both genes had to be
inactivated for complete inhibition (see Results). Moreover,
single knockout mutagenesis revealed that both synthases can
replace the activity of the other, and synthesize similar
amounts of polyglucans under the tested physiological
conditions (see Results). Potentially distinctive roles of these
isoenzymes in polyglucan (glycogen or starch-like) biosynthesis were not detectable with the applied methods and thus
remain undefined.
Essential role of glycogen synthesis in dark–light
cycles
Probably the most important changes which cyanobacterial
metabolism has to accommodate to are the transitions
between light and dark periods. Cells incapable of glycogen
synthesis are obviously more sensitive to regular dark–light
transitions than wild-type cells (Fig. 1). The impaired
viability (reduced survival rate) of such cells supports the
idea of glycogen’s dual role as reserve and buffer substance
(Carr, 1988).
Dark–light transitions involve changes in cytoplasmic pH
and redox state, as well as changes in the intracellular
concentration of specific metabolites and metal ions (Smith,
1982). These factors mainly regulate the switch between
assimilatory (photosynthetic) and catabolic pathways in the
cyanobacterial cell (Smith, 1982). Transitions from dark to
light stress the cells, as reflected in an increased transcript
accumulation of stress genes (Gill et al., 2002). We assume
that both glycogen and glycogen synthesis have crucial roles
in this acclimation process, both acting as buffers and
cellular tools for compensation of such stressful energetic
transitions, mainly to ameliorate and to avoid futile cycles
during the process of changing photosynthetic activity and
of metabolic switching, similar to what is assumed for starch
synthesis in plants (Sun et al., 1999).
In addition to this regulatory function, the glycogen pool
might be the main endogenous reserve in Synechocystis that
provides maintenance energy for cell integrity, function
and viability in dark periods. Based on this assumption, the
inactivation of glycogen synthesis should result in a nonviable genotype, as predicted by flux-balance analysis of the
Synechocystis metabolic network (Knoop et al., 2010). Here,
glycogen metabolism appears to be vital to produce all
precursors required for biomass formation and to regenerate all metabolites and cofactors, especially in dark
periods. However, the complete segregation genotype of
both AGP and glycogen synthase mutants of Synechocystis
suggests the presence of other available (minor) endogenous respiratory substrates. For instance, starch-less mutants
of Nicotiana sylvestris accumulate sufficient soluble neutral
sugars during the day which obviously provide respiratory
substrates in the dark (Huber & Hanson, 1992). Thus, other
available substrates such as sugars, lipids or chlorophyll may
partly compensate for the lack of glycogen. However, the
restricted viability of the Synechocystis mutants (Fig. 1) and
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Microbiology 158
Cyanobacterial energy and carbon metabolism
the reduced respiratory oxygen consumption by S. elongatus
PCC 7942 (Suzuki et al., 2010) indicate that this polymer is
the major respiratory substrate in cyanobacteria. We
nevertheless assume that glycogen and glycogen synthesis
is primarily needed as a buffer substance and an ameliorator
in acclimation processes whenever metabolic switches in
amphibolic pathways occur. The polymer likely has a similar
role during transitions of cyanobacteria to periods of
nitrogen starvation or to photomixotrophic growth. Both
conditions conceivably mimic the physiological states of
cells under light–dark transitions (Lee et al., 2007) and will
be discussed below in more detail.
Essential role of glycogen synthesis for the
energy and carbon metabolism
Phenotypes that result from the inability to synthesize
glycogen are very prominent: (i) reduced oxygen evolution
rates (see Results), (ii) glucose-sensitivity (Fig. 2b), (iii) a
non-bleaching phenotype (Fig. 5) and (iv) overflow of
pyruvate and 2-oxoglutarate (Fig. 4). All detected characteristics occur explicitly in the light when glycogen synthesis is
needed and significant carbon fluxes are sensed by the cell.
In cyanobacteria, photosynthesis and carbon/nitrogen
assimilation are linked via the ferredoxin/thioredoxin
system. Prominent targets of the thioredoxin system are
enzymes of carbon and nitrogen assimilation, such as
ribulose-1,5-bisphosphate carboxylase/oxygenase and glutamine oxoglutarate aminotransferase, as well as all enzymes
involved in glycogen synthesis, such as phoshoglucomutase,
AGPase, glycogen synthase and branching enzyme (Lindahl
& Florencio, 2003) (see Fig. 6 for an overview). This
regulatory principle explains the fact that glycogen synthesis
and thus the phenotypes of glycogen deficiency occur only in
the light and stop immediately in the dark.
The significant reduction of photosynthetic capacity (see
Results) in the glycogen-deficient mutants is a common
feature of all currently published polyglucan-free mutants
in photosynthetic organisms (Eichelmann & Laisk, 1994;
Miao et al., 2003b; Sun et al., 1999; Suzuki et al., 2010). The
real cause has not yet been identified. We assume a role of
glycogen in photosynthetic reactions as has been suggested
for starch (Eichelmann & Laisk, 1994; Sun et al., 1999).
Glycogen synthesis has been proposed to act as an electron
acceptor and an ameliorator during periods of changing
photosynthetic activities. It has also been suggested that
glycogen metabolism is involved in anaplerotic reactions of
the Calvin–Benson–Bassham cycle during acclimation to
changing CO2 concentrations (Eisenhut et al., 2008).
Therefore, glycogen metabolism might be involved both
in maintaining the redox homeostasis and in the
acclimation to changes in photosynthetic activity.
Glycogen’s essential role in stress responses
(1) Metabolic switch into the ‘dormant state’:
accumulation of reserves. Various micro-organisms
http://mic.sgmjournals.org
including species of the cyanobacterial orders Nostocales
and Stigonematales are capable of forming specialized resting
cells in response to macronutrient starvation. In contrast,
Oscillatoriales and Chroococcales such as Synechocystis lack the
capacity to form such spore-like cells. Nevertheless, their cells
also switch stringently from a growth-related mode (active
state) to a maintenance/survival mode without growth and
multiplication (dormant state) (Kaprelyants et al., 1993).
During the transition of Synechocystis from an active to a
dormant state, several metabolic processes are substantially
changed (Krasikov et al., 2012; Aguirre von Wobeser et al.,
2011). Here, a massive accumulation of carbon reserve
polymers like glycogen (Fig. 4a) and polyhydroxybutyrate
(PHB), degradation of phycobiliproteins (Fig. 5) and an
activation of catabolic pathways are observed (Schwarz &
Forchhammer, 2005). The simultaneous synthesis of both
carbon reserve polymers by Synechocystis and some other
cyanobacteria is an exception to the principle that only one
kind of carbon reserve material is formed by a given species.
In this context, it has to be emphasized that a mutant of
Synechocystis defective in PHB synthesis (DphaAB) employs
in response to nitrogen starvation a similar metabolic
change, including phycobilisome degradation like the wildtype, and was only marginally affected in its viability
(Taroncher-Oldenburg & Stephanopoulos, 2000) (see
overview in Fig. 6). By contrast, mutants defective in
glycogen synthesis seem to be unable to switch: they are
incapable of cell division and of the chlorotic response (Fig.
5). Moreover, these cells lost their viability after a few days.
These data indicate that in Synechocystis glycogen synthesis
plays a more decisive role in the metabolic changes during
the acclimation response and in viability maintenance during
starvation periods than the other carbon reserve PHB. The
long survival time of Synechocystis correlates with the small
average chain length of cyanobacterial glycogen and sustains
glycogen’s global role as a durable energy reserve that is
essential for bacterial survival and persistence to stress
conditions (Preiss, 1984; Wang & Wise, 2011).
(2) Chlorotic response. The observed non-bleaching
phenotype (lack of degradation of phycobilisomes) of
mutants incapable of glycogen synthesis (Fig. 5) is usually
assigned to the absence, modification or inhibition of
proteins that are involved in the nitrogen signalling cascade
(Schwarz & Forchhammer, 2005). Just a few metabolic
knockouts are known to cause a non-bleaching phenotype
(Lahmi et al., 2006; Morrison et al., 2005). Here, manipulations of pyruvate metabolism via knockout of alanine
dehydrogenase or of phosphotransacetylase in a Dhik33
(DdspA) background result in a deceleration but not in a total
inability of phycobilisome degradation, as is observed in cells
impaired in glycogen synthesis (see overview Fig. 6). Even
7 days of nitrogen starvation did not change the greenish
colour of glycogen-deficient mutant cells (Fig. 3a). Changes
in metabolite levels by manipulation of the pyruvate
metabolism, such as knockouts of alanine dehydrogenase
or glycogen synthesis, obviously influence the progression of
phycobilisome degradation (Fig. 4a, overview in Fig. 6). We
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3039
M. Gründel and others
AGP*
ATP
PPi
ADP-Glucose
Glucose-1-P
GS*
PGM*
Pi
ADP
Glycogen
GBE*
Glucose
G6PDH*
ATP
GLK
+
Glucose-6-P
+
Ribulose-5-P
Glycolysis
H2O
ADP
Xylulose-5-P
Pi
H2O
P
Acetyl-P i
ATP
CO2
NADPH+H+
H2O
Fructose-6-P
Pi
Ribose-5-P
Pentose
Seduheptulose-7-P
Fructose-1,6-bis-P
phosphate
Glyceraldehyde-3-P
+
Fructose-6-P
Dihydroxyacetone-P
H2O
Acetyl-P
+
NADP
Pi
NADPH+H+
GAP2
NAD
GAP1
Pi
NADH+H+
ADP
PhaE
PhaC
ATP
NADP+
H2O
H 2O
Pi
P-Enolpyruvate
AMP
P
H2iO
ATP
ADP
HS-CoA
ATP
Alanine
Glycolate
+
NAD
Pyruvate
+
2 Fdax
HS-CoA
CO P
2 Fd2red i
NAD
HS-CoA
CO2
+
NADH+H
+
NAD
NADH+H+
+
NH4
–
HCO3
2-Oxoglutarate
H2O
HS-CoA
Glyoxylate
Acetyl-P
ADP
ATP
HS-CoA
AMP
PPi
Acetate
+
HS-CoA
Citrate ATP
Oxaloacetate
+
+
NADH+H
+
NAD
NH4
H2O2
H2O
O2
+
NADH+H
PTA/ DspA
Acetyl-CoA
2x
Glutamate
Aspartate
NADH+H
NADPH+H+ NADP+
+
NAD
Acetaldehyde
Ethanol
NADPH+H+
+ CO2
NADP
cis-Aconitate
Malate
Pi
ADP
ATP
TCA cycle
H2O
ADP
Pi
Fumarate
+
+
NADPH+H
H2O
+
NADP 2-Oxoglutarate
Succinate
NADP
CO2
+
NADPH+H
+
NADH+H
2 Fdred
GOGAT*
ATP
HS-CoA
SuccinylADP
+
CoA
Cyanophycin
H2O
H2O
Glutamine
Isocitrate
FADH2
FAD+
Reductive
2-Phosphoglycolate
Glycerate-2-P
AcetoacetylCoA
PhaA
Ribulose-1,5-bis-P
O2
NADPH+H+
ALD
ADP
H2O
CO2
RubisCO*
2x
Glycerate-3-P
3-Hydroxybutanoyl-CoA
Pi
ATP
1,3-bis-P-glycerate
HS-CoA
Glyceraldehyde-3-P
pathway
Erythrose-4-P
PHB
Oxidative
NADP+
H2O
NADP NADPH+H
Sedoheptulose
1,7-bisphosphate
Glyconeogenesis
6-P-Gluconate
6-P-Gluconolactone
ADP
Succinate
semialdehyde
ATP
NH4+
NH4+
Glutamate
CO2
Glutamate
2-Oxoglutarate
ADP
Pi
NAD+
2 Fdax
GS
+
NADPH+H
+
NH4
+
NADP
CO2
H2O
4-Amino
butanoate
Glutamate
ATP
Arginine
assume that additional metabolic signals within the pyruvate
metabolism contribute to the acclimation response in
cyanobacteria. To what extent molecular processes such as
transcription, RNA processing, translation, post-translational
modifications and/or protein–protein interactions of the
3040
nitrogen signalling cascade are affected by these manipulations is as yet unknown.
(3) Energy spilling reactions. Cells incapable of glycogen
synthesis get into an acute dilemma since energy and
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Cyanobacterial energy and carbon metabolism
Fig. 6. Illustration of central carbon and reserve metabolism of Synechocystis sp. PCC 6803 according to Knoop et al. (2010).
Prominent enzymes of the reductive (RubisCO, ribulose-1,5-bisphosphate carboxylase/oxygenase) and oxidative pentose
phosphate pathways (G6PDH, glucose-6-phosphate dehydrogenase), of glycolytic (GLK, glucokinase; GAP1, glycerin-3phosphate dehydrogenase, form 1) and gluconeogenetic route (GAP2, glycerin-3-phosphate dehydrogenase, form 2), of
pyruvate metabolism (ALD, alanine dehydrogenase; PTA, phosphotransacetylase; DspA, sensory histidine kinase Hik33), of
glycogen synthesis (PGM, phosphoglucomutase; AGPase, ADP-glucose pyrophosphorylase; GS, glycogen synthases; GBE,
glycogen branching enzyme), of polyhydroxybutyrate (PHB) synthesis (PhaA, PHA-specific beta-ketothiolase; PhaCE, PHB
synthase) and of the nitrogen uptake system (GS, glutamine synthetase; GOGAT, glutamine-oxoglutarate aminotransferase)
were marked. Metabolic knockouts (shown by a black cross) that result in a glucose-sensitive phenotype or non-bleaching
phenotype upon nitrogen starvation are shown with red or green semi-circles, respectively. Prominent targets of the thioredoxin
system are marked with asterisks.
carbon excess cannot flow into glycogen synthesis. To
release the excess, cells lose significant amounts of pyruvate
and 2-oxoglutarate (Fig. 4). The amount of carbon released
into the medium by the glycogen-deficient mutants as
pyruvate and 2-oxoglutarate was calculated roughly to
correspond to 30–60 % of the carbon that would have been
used for glycogen synthesis by the wild-type under similar
conditions (Fig. 4). Massive release of carbon compounds
under conditions of unbalanced growth is well known as
‘overflow’ metabolism in heterotrophic bacteria. The
‘overflow’ metabolism is a form of ‘energy spilling’ (Russell,
2007). Characteristic of the overflow metabolism is the
excretion of partially oxidized metabolites such as 2oxoglutarate, pyruvate, succinate and gluconate, capsular
material and protein into culture medium (Russell & Cook,
1995). The mechanism of membrane passage of these
compounds is unclear for Synechocystis. Certainly, it does
not reflect a general leakage from the cells since a concomitant
release of glutamate, which is known to be a highly abundant
free amino acid in Synechocystis (Eisenhut et al., 2008), was
not observed in the culture medium (see Results). Our data
indicate that glycogen synthesis is an essential cellular tool for
integration of energy and carbon homeostasis to avoid energy
spilling reactions in Synechocystis.
(4) The mystery of photomixotrophic sensitivity: role of
glycogen synthesis for stressful amphibolic reactions in
response to energetic and carbon transitions. Simul-
taneous assimilation of CO2 and glucose (mixotrophy)
results in a unique situation inside the cell. Directly opposed
pathways are used in parallel: glycolysis and gluconeogensis
as well as reductive (Calvin cycle) and oxidative pentose
phosphate pathways by sharing both enzymes and reaction
intermediates (Kaplan et al., 2008) (see overview in Fig. 6).
Such conditions increase the growth rate of Synechocystis
(Fig. 2b) and other cyanobacteria. In contrast, cells incapable
of glycogen synthesis have a significantly reduced growth
rate under these conditions (Fig. 2b). On agar plates
containing glucose, the cells lost their viability and were
unable to resume growth within a few days. This phenotype is referred to as ‘photomixotrophic sensitivity’. The
respective wild-type, however, retained the property of
growth in the presence of glucose under the otherwise
identical conditions. The photomixotrophic sensitivity
phenotype is also known from other metabolic knockouts
http://mic.sgmjournals.org
such as glycolytic unidirectional glycerol-3-phosphate
dehydrogenase 1 (gap1) (Koksharova et al., 1998; Osanai
et al., 2007) (see overview in Fig. 6). It is assumed that
glucose uptake inflicts lethality and leads to a disturbed
carbon flux and balance in such mutants (Kaplan et al.,
2008). The exact mechanism is as yet unclear. Our data on
overflow metabolism (Fig. 4) indicate that blocks in
glycogen synthesis lead to a severely disturbed carbon flux
and balance. We assume that glycogen synthesis enables the
cell to cope with intrinsic bottlenecks or deficits in the form
of anabolic and anaplerotic reaction sequences and thus
allows essential metabolic switches in response to energetic
and carbon transitions. To what extent metabolic routes
other than photosynthetic reactions are impaired by
knockout of glycogen synthesis is as yet unclear. It is
known that the darkness-dependent glucose sensitivity is
based primarily on inhibition of the oxidative pentose
phosphate pathway by knockout of glucose-6-phosphate
dehydrogenase (G6PDH) (Jansén et al., 2010). The oxidative pentose phosphate pathway is involved in not only
glycogen mobilization through night periods but also the
metabolization of organic substrates into glycogen in the
light (Smith, 1982). Therefore, we assume a close interaction
between oxidative pentose phosphate pathway and glycogen
metabolism. The G6PDH is controlled by fluctuating ATP
and NADPH concentrations as well as light-induced pH
changes in the extra-thylakoidal environment (Grossman &
McGowan, 1975). Glucose addition usually changes the
cytosolic pH and gene expression (Ryu et al., 2004). We
assume that a block of oxidative pentose phosphate pathway
and/or a pH shift cause an inhibition of G6PDH and thus
of glucose metabolization in cells defective in glycogen
synthesis. The putative impact of an acidification of growth
medium on viability, caused by the overflow of pyruvate and
2-oxoglutarate, was excluded by optimal HEPES buffering at
pH 8.0 and controlling. Dilution effects in batch cultures
(Fig. 2b) might attenuate putative acidification in contrast to
more concentrating effects (slow diffusion) on solidified
media.
The glucose uptake is also repressed in glycogen-deficient
mutants (Fig. 2b). This crucial effect does not correlate
with the level of growth restriction (Fig. 2b). We assume
that a currently unknown negative feedback loop from
central carbon metabolism acts on the glucose uptake
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M. Gründel and others
system similar to that observed for trehalose metabolism in
Saccharomyces cerevisiae (Teusink et al., 1998).
genes of the cyanobacterium Synechocystis PCC 6803. Photosynth Res
37, 139–146.
Conclusion
Franche, C. & Damerval, T. (1988). Tests on nif Probes and DNA
Hybridizations. In Cyanobacteria, pp. 803–808. Edited by L. Packer &
A. N. Glazer. San Diego, California: Academic Press, Inc.
In conclusion, cells incapable of glycogen synthesis are an
ideal tool for identification of metabolic regulatory principles and signals. These mutants will most likely contribute
enormously to the understanding of complex carbon and
nitrogen metabolism and of acclimation responses in
cyanobacteria.
Gill, R. T., Katsoulakis, E., Schmitt, W., Taroncher-Oldenburg, G.,
Misra, J. & Stephanopoulos, G. (2002). Genome-wide dynamic
transcriptional profiling of the light-to-dark transition in Synechocystis
sp. strain PCC 6803. J Bacteriol 184, 3671–3681.
Grossman, A. & McGowan, R. E. (1975). Regulation of glucose 6-
phosphate dehydrogenase in blue-green algae. Plant Physiol 55, 658–
662.
Hagemann, M. & Erdmann, N. (1994). Activation and pathway of
glucosylglycerol synthesis in the cyanobacterium Synechocystis sp.
PCC 6803. Microbiology 140, 1427–1431.
ACKNOWLEDGEMENTS
We thank our past colleagues Annegret Wilde and Kerstin Baier for
their support in knockout mutagenesis and our FORSYS cooperation
partners Ralf Steuer and Henning Knoop for metabolic network
analysis. Excellent technical assistance was provided by Gisa Baumert
and Lena Stansfeld. This work was financially supported by the
German Ministry of Science within the framework of FORSYSPartner (BMBF-31101939).
Huber, S. C. & Hanson, K. R. (1992). Carbon partitiong and growth of
a starchless mutant of Nicotiana sylvestris. Plant Physiol 99, 1449–
1454.
Jansén, T., Kurian, D., Raksajit, W., York, S., Summers, M. L. &
Maenpaa, P. (2010). Characterization of trophic changes and a
functional oxidative pentose phosphate pathway in Synechocystis sp.
PCC 6803. Acta Physiol Plant 32, 511–518.
Kaplan, A., Hagemann, M., Bauwe, H., Kahlon, S. & Ogawa, T.
(2008). Carbon acquisition by cyanobacteria: mechanisms, compar-
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