Department of Plant Biology and Center for
the Study of the Early Events in Photosynthesis, Arizona State
University, Tempe, Arizona 85287-1601
Respiration in cyanobacterial thylakoid membranes is interwoven
with photosynthetic processes. We have constructed a range of mutants
that are impaired in several combinations of respiratory and
photosynthetic electron transport complexes and have examined the
relative effects on the redox state of the plastoquinone (PQ) pool by
using a quinone electrode. Succinate dehydrogenase has a major effect
on the PQ redox poise, as mutants lacking this enzyme showed a much
more oxidized PQ pool. Mutants lacking type I and II NAD(P)H
dehydrogenases also had more oxidized PQ pools. However, in the mutant
lacking type I NADPH dehydrogenase, succinate was essentially absent
and effective respiratory electron donation to the PQ pool could be
established after addition of 1 mM succinate. Therefore, lack of the
type I NADPH dehydrogenase had an indirect effect on the PQ pool redox
state. The electron donation capacity of succinate dehydrogenase was
found to be an order of magnitude larger than that of type I and II
NAD(P)H dehydrogenases. The reason for the oxidized PQ pool upon
inactivation of type II NADH dehydrogenase may be related to the facts
that the NAD pool in the cell is much smaller than that of NADP and
that the NAD pool is fully reduced in the mutant without type II NADH
dehydrogenase, thus causing regulatory inhibition. The results indicate
that succinate dehydrogenase is the main respiratory electron transfer pathway into the PQ pool and that type I and II NAD(P)H dehydrogenases regulate the reduction level of NADP and NAD, which, in turn, affects
respiratory electron flow through succinate dehydrogenase.
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INTRODUCTION |
Synechocystis sp. strain
PCC 6803, a unicellular cyanobacterium, contains a respiratory electron
transport chain on both the cytoplasmic and thylakoid membranes
(11, 23). The cytoplasmic membrane forms the inner
boundary of the periplasmic space and is known to contain proteins
typically associated with respiratory electron transport, such as
NAD(P)H dehydrogenase, cytochrome b6f, and terminal oxidases
(presumably predominantly Cyd, a putative quinol oxidase) (7, 11,
23). Two types of NAD(P)H dehydrogenase have been found in
cyanobacteria. One is a NADPH-preferring type I dehydrogenase (NDH-1)
that is encoded by ndh genes, consists of about a dozen
subunits, and contributes to a proton gradient across the membrane
(2). The second type of dehydrogenase is a NADH-oxidizing
type II dehydrogenase (NDH-2) consisting of a single subunit and
presumably not contributing to a proton gradient across the membrane.
Three genes for NDH-2 (ndbA, ndbB, and ndbC) are
found in Synechocystis sp. strain PCC 6803.
The thylakoid membrane contains both a photosynthetic electron
transport chain that includes photosystem I (PSI) and PSII and a
respiratory electron transport chain containing, among others, NDH-1,
succinate dehydrogenase (SDH), and a cytochrome
aa3-type terminal oxidase (CtaI) (5, 19,
24). Genes for a third terminal oxidase (CtaII) are present in
the genome, but no functional evidence for CtaII has been obtained thus
far. The respiratory and photosynthetic electron transport chains in
the thylakoids have electron carriers in common, including the
cytochrome b6f complex, the
plastoquinone (PQ) pool, and soluble redox-active proteins (21,
27).
Electron transport into and out of the PQ pool in cyanobacterial
thylakoids is complex in that many different pathways exist and the
relative rate at which electrons are transported via each pathway
depends on the capacity of the pathway and the availability of oxidized
substances to accept electrons (or reduced compounds to donate them),
etc. Three intersecting pathways traditionally have been viewed as
dominant in Synechocystis sp. strain PCC 6803 thylakoids.
The three pathways are linear photosynthetic electron transport,
respiratory transport from NADPH and succinate to cytochrome oxidase,
and cyclic electron transport around PSI (electrons at the acceptor
side of PSI returning to the PQ pool). However, electrons can easily
cross from one pathway to another at the level of the PQ pool,
cytochrome b6f, and/or soluble
carriers. For example, in the absence of PSI, PSII-generated electrons
are fed into cytochrome oxidase (25), and in darkness,
respiratory electrons are used to reduce the acceptor side of PSII if
terminal oxidases are blocked (8).
Even though a wealth of data has accumulated over the years regarding
individual pathways, very little is known regarding the relative
importance of the various electron transfer pathways in the organism in
vivo. However, this information is critical to an understanding of the
metabolism and its regulation in the organism. As cyanobacteria are
thought to be related to the ancestor of chloroplasts, understanding of
photosynthetic and respiratory electron transport interaction and
regulation is likely to also impact our understanding of such
processes in photosynthetic eukaryotes. Regulatory processes with
respect to energy requirement (ATP production) (17,
22-24), metabolic cofactor reduction-oxidation (such as NADP+ reduction at PSI, allowing CO2 fixation
via the Calvin cycle) (18), and redox sensing-poising of
the PQ pool for gene regulation (8, 16) are well
established phenomenologically, but the primary signals and mechanisms
for such regulation remain unclear.
To aid in providing a comprehensive overview of photosynthesis and
respiration in vivo, we determined the PQ pool redox state, the
reduction level of NAD-NADP, and the concentration of organic acids
such as succinate in a range of strains lacking one electron flow
pathway or multiple electron flow pathways. Our findings indicate that
succinate levels, the PQ pool redox state, and the NADP reduction state
are interrelated, with the succinate concentration and SDH activity
being much more important factors than has been realized thus far.
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MATERIALS AND METHODS |
Growth conditions.
Wild-type and mutant cultures of
Synechocystis sp. strain PCC 6803 were grown in liquid BG-11
medium (20) at 30°C. Where indicated, 5 mM glucose was
added for photoheterotrophic growth. Cultures were grown at a light
intensity of 40 to 50 µmol of photons m
2
s1, except when the cultures were grown at a low light
intensity (3 to 5 µmol of photons m2 s1).
Cells were grown in ambient air or, if indicated, in air enriched with
CO2 (bubbling with air containing 3% CO2).
Cells used for organic acid analysis, quinone (Q) redox state
determination, and [NAD] or [NADP] quantifications were harvested
at an optical density at 730 nm (OD730) of 0.5, as
determined with a Shimadzu UV 160 spectophotometer. This corresponds to
mid-exponential phase.
Deletion mutant construction and segregation.
The NDH-1
deficient strain, a gift from T. Ogawa, carries an insertional mutation
of the ndhB gene. The two sdhB genes
(sll1625 and sll0823) were deleted from the
wild-type, NDH-2-deficient, and Cyd-deficient/CtaII-deficient
strains by using plasmids p
sll1625 (Cmr) and psll0823
(Kmr), which were described previously (5).
For deletion of the two sdhB genes from the PSI-deficient
and CtaI-deficient strains, the antibiotic resistance cassette of the
psll1625 and psll0823 plasmids, respectively, was replaced with
the PvuII-PvuII fragment from pZeol
(8), which contains the zeocin resistance
(Zor) cassette. To create a PSII-deficient/SDH-deficient
strain, the SDH-deficient strain (sll0823
sll1625) was used as the background strain for deletion
of PSII by transformation with the pDICEmr plasmid, in
which an erythromycin resistance cassette replaces part of the
psbDIC operon.
Segregation analysis was performed by PCR with primers specific for the
sequence of the flanking regions of the gene being deleted. In
addition, PCR was performed by using primers recognizing sequences
inside the wild-type sequence that was deleted in the mutant. The
exclusive presence of a band corresponding to the inactivated gene in
the first PCR and the absence of a product in the second PCR were taken
as evidence of full segregation.
Organic acid isolation and analysis.
Organic acids were
isolated, purified, derivatized, and analyzed by gas
chromatography-mass spectrometry as previously described (5). One liter of cells to be harvested for organic acid
analysis was grown to an OD730 of 0.5. The levels of malate
and isocitrate were determined as described for succinate and fumarate
(5): derivatized standards were run, and mass spectral
fragmentation patterns coupled with retention time were used to
identify and quantitate the derivatized organic acid of interest.
NADP-NADPH and NAD-NADH extraction.
Extraction and
determination of NADP/NADPH and NAD/NADH ratios and concentrations were
carried out by two separate methods. In the first method, extracts to
be used for enzymatic cycling reactions were prepared by a modification
of the procedure used by Wagner and Scott (26) for
erythrocytes. One liter of cells was grown to an OD730 of
~0.5 and harvested by centrifugation. The pelleted cells were
resuspended in approximately 0.8 ml of breakage buffer (20 mM
nicotinamide, 20 mM NaHCO3, 100 mM
Na2CO3) precooled to 4°C. The resuspended
cells were frozen in liquid N2 and thawed quickly in a
water bath at 20°C. Glass beads (70 to 100-µm diameter;
approximately one-third of the volume of the cell suspension) were
added, and the cells were broken by using a mini-beadbeater (BioSpec
Products) (4 × 30 s). Cells were incubated on ice for 1 min
between breakage cycles. Complete breakage was achieved when the
chlorophyll contents (measured by determining OD663) when
extracted with 80% acetone and with methanol were nearly identical;
80% acetone does not efficiently extract chlorophyll from intact
cells. Following breakage, all subsequent steps were carried out in
darkness to avoid photodegradation of the pyridine nucleotides. The
cell debris was spun at 14,000 rpm in an Eppendorf 5415 microcentrifuge, and the supernantant was removed to a new tube. To
determine [NADP]total, 1 µl of the supernatant was
added to a glass test tube containing 0.9 ml of ice-cold NADP cycling buffer (100 mM Tris-HCl [pH 8.0], 0.5 mM thiazolium blue [MTT], 2 mM phenazine ethosulfate [PES], 5 mM Na4EDTA, 1.3 IU of
glucose-6-phosphate dehydrogenase [G-6-PDH] per ml) and incubated in
the dark at 37°C for 10 min. Following incubation, 100 µl of
glucose-6-phosphate was added and the spectrophotometric changes at 570 nm were monitored for 100 s. The reaction is termed a cycling
reaction since NADP+ is reduced by G-6-PDH to NADPH, which
is then oxidized by PES, which, in turn, is reoxidized by MTT, yielding
a color change reaction (12). None of the samples remained
in cycling buffer for more than 15 min to avoid degradation.
To determine the [NAD]total in the extracts, 1 µl of
the extract was added to 0.9 ml of NAD cycling buffer (100 mM Tris-HCl [pH 8.0], 0.5 mM MTT, 1 mM PES, 0.2 mg of alcohol dehydrogenase per
ml, 1% [wt/vol] bovine serum albumin). The principle of the cycling
reaction for NAD determination is the same as for NADP, except that
NAD-specific alcohol dehydrogenase is used rather than NADP-specific
G-6-PDH. To initiate the reaction, 100 µl of 30% ethanol was added
and the spectrophotometric changes at 570 nm were monitored for
100 s.
An aliquot of the cell extract was heated for 30 min at 60°C in a
glass test tube in the dark. One microliter of this heated sample was
then assayed for NAD and NADP as described above. The heating step
denatures the oxidized, but not the reduced, form of the pyridine
nucleotides, and the rates observed in the cycling reaction represent
the NADH and NADPH concentrations. [NAD+] and
[NADP+] were calculated by subtracting the reduced value
from the [NAD]total and [NADP]total.
A second method by which to extract and detect NADP-NADPH and
NAD-NADH involved fluorescence-based high-pressure liquid
chromatography (HPLC) using a method derived from that of
Klaidman et al. (14). First, 1 liter of cells
(OD730 = 0.5) was pelleted and resuspended to 1 ml in
a mixture containing 0.06 mM KOH, 0.2 mM KCN, and 1 mM
bathophenanthrolinedisulfonic acid. In this solution, the oxidized forms of NAD and NADP will be derivatized with CN, making the oxidized
form visible by fluorescence (emission at 460 nm upon excitation at 330 nm) at an efficiency nearly equivalent to that of the reduced form
(14). Glass beads were added to a total volume of 1.5 ml,
and the cells were broken as described above. Samples were spun at
14,000 rpm in an Eppendorf 5415 microcentrifuge to remove the insoluble
matter, and the supernatant was further rinsed with 0.5 volume of
chloroform to ensure the removal of all lipid material. Samples were
spun through a 0.45-µm-pore-size microcentrifuge spin filter.
Concentrations and ratios of the oxidized and reduced forms of NAD and
NADP were confirmed by fluorescence-based HPLC analysis as described by
Klaidman et al. (14) using an HP1100 LC with an Agilent
1100 fluorescence detector and a Waters Spherisorb 5 µm ODS1 column
(4.6- by 250-mm analytical C18 column).
Isocitrate dehydrogenase and malate dyhydrogenase coenzyme
specificities.
An extract of soluble enzymes was prepared from 10 liters of wild-type Synechocystis sp. strain PCC 6803: Cells
were harvested by centrifugation and resuspended in a buffer similar to
that described for thylakoid isolation (5), with the
exception that the MgCl2 and CaCl2
concentrations were increased to 25 mM MgCl2 and 50 mM
CaCl2. Cells were broken with glass beads as described earlier, and the cells debris was removed by centrifugation. Solid ammonium sulfate was added in small aliquots while stirring at 0 to
4°C until the solution was 85% saturated (
516 mg of ammonium sulfate per ml). The solution was centrifuged at 48,000 × g for 30 min, and the supernatant was discarded. The wet pellet
was resuspended in 50 µl of 80 mM potassium phosphate buffer (pH
7.0). Assays were carried out by addition of 2 µl of resuspended
protein to one of the following solutions (kept at 37°C).
For isocitrate dehydrogenase activity measurements, the cuvette
contained 1 ml of the reaction solution (10 mM MnCl2,
isocitrate [0 or 10 mM], either NADP [0 or 0.1 mM] or NAD [0 or
0.1 mM], and 80 mM potassium phosphate [pH 7.0]). Assays were
conducted on an HP8452 diode array spectrophotometer by monitoring
changes in A340 over time after addition of the
protein extract.
Malate dehydrogenase activity measurements were carried out similarly,
with the exception that malate replaced isocitrate as the enzyme
substrate in the reaction mixtures.
Respiration measurements.
Oxygen uptake was measured as
described previously (9), with the exception that for 2 min prior to the measurement, samples were exposed to saturating white
light in the presence of DCMU [(3,3-dichlorophenyl)-1,1-dimethylurea]
to ensure complete PQ pool oxidation at the beginning of the
measurement. Without PSII activity, which is inhibited by DCMU, and in
the presence of PSI activity at high light intensity, the PQ pool
becomes oxidized very quickly. All cells were grown under
photoheterotrophic conditions.
Q electrode.
The Q electrode apparatus was assembled and
operated as described previously (4, 6, 28). Cells were
harvested in mid-exponential phase (OD730 = 0.5) and
resuspended to an OD730 of 4.0 in 10 mM HEPES-NaOH buffer
(pH 7.4)-50 mM KCl. A 10-ml volume of the cell suspension was placed
in a cuvette that was kept at 28°C and that contained a glassy carbon
working electrode poised at +360 mV with respect to an Ag-AgCl
reference electrode, as well as a platinum auxiliary electrode.
Following initial electrode equilibration, exogenous ubiquinone-1
(UQ-1) was added to the cuvette to a final concentration of 0.2 µM.
This Q acts as a redox mediator between the PQ pool in the cells and
the electrode surface. The current created by the oxidation of the Q at
the electrode surface was measured as a function of time using a CV-50W
voltammeter (Bioanalytical Systems Inc., West Lafayette Ind.). The
instrument's sensitivity was set at 100 nA V1.
Illumination was provided by a halogen lamp, and light was passed through a filter that kept wavelengths of less than 430 nm from reaching the sample. The light intensity was modulated with a combination of reflectance and wire mesh filters.
Cells were incubated in darkness for 5 min before traces were recorded.
The light intensity was increased in a stepwise manner. To obtain full
reduction of the PQ pool, cells were incubated with 0.2 mM KCN in
darkness, which fully blocks oxidation of the PQ pool by terminal oxidases.
Chlorophyll fluorescence measurements.
The relative
chlorophyll fluorescence yield during incubation in darkness was
monitored by using a Walz fluorometer (PAM 101, 102) in order to help
determine the kinetics of PQ pool reduction indirectly. The intensity
of the measuring light was kept minimal (<0.01 µmol of photons
m2 s1). The time constant of the instrument
response was 960 ms. Cells were harvested in mid-exponential growth
phase (OD730 = 0.5), spun down, and resuspended in 10 mM HEPES-NaOH (pH 7.0) buffer at a chlorophyll concentration of 10 µg
ml1. The fluorescence level (F0)
was measured for 15 s with the measuring light on, the measuring
light was turned off, and KCN was injected to a final concentration of
1 mM. The measuring light (<0.01 µmol of photons m2
s1) was turned on again for 5 s at 10- to 35-s
intervals for the duration of the measurement. The measuring light
itself did not have a noticeable actinic effect.
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RESULTS |
PQ pool redox state measurements.
To determine the effects of
individual redox-active complexes on the redox state of the PQ pool in
the thylakoid membrane of Synechocystis sp. strain PCC 6803, we have utilized a set of mutant strains that lack one or more of the
enzymes catalyzing electron transport into or out of the PQ pool in the
thylakoid membrane. These mutants were analyzed for the relative redox
state of the thylakoid PQ pool by means of a Q electrode. This
apparatus has been used with plant mitochondria (6), pea
chloroplasts (4), Rhodobacter membranes
(28), and most recently with intact cells of
Synechocystis sp. strain PCC 6803 (10). In
PSI-containing strains, the PQ pool was fairly reduced in darkness and
became increasingly oxidized at increasing light intensities (Table
1). The reason for this is that PSI is
very abundant in Synechocystis sp. strain PCC 6803 and is a
highly efficient and high-capacity electron acceptor in the light. In
contrast, PSI-deficient strains became very reduced at increasing light
levels (Table 1). The other major electron acceptor is cytochrome
oxidase (CtaI), and indeed, in darkness the PQ pool was fully reduced
when CtaI was absent (Table 1; Fig. 1).
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TABLE 1.
Relative reduction state of UQ-1,a
reflecting the redox state of the PQ pool, in different strains at
various light intensities as measured with a Q electrode
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FIG. 1.
Sample trace acquired with the Q electrode by using
wild-type (a), NDH-2/SDH-deficient (b), and CtaI-deficient (c) strains
upon illumination and upon addition of KCN. Light level changes (upper)
and total time (lower) are indicated on the horizontal axes. The
vertical axes show the current in nanoamperes. Under our experimental
conditions, a fully reduced UQ-1 pool yields a current of about  4 nA;
a fully oxidized pool corresponds to about 2 nA.
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SDH, NDH-2, and NDH-1 are potential respiratory donors to the PQ pool.
Deletion of any of these three donors led to a much more oxidized PQ
pool in darkness (Table 1), whereas an even more pronounced effect was
seen upon deletion of both SDH and NDH-2 (Fig. 1 and Table 1). In all
strains, a fully reduced PQ pool was observed when KCN was added and
the light was turned off, just as was observed in the CtaI-deficient
strain in darkness without KCN
{[UQH2]/([UQ]+[UQH2]) = 1.00}
(Table 1). However, note that in a mutant lacking SDH and NDH-2, this
KCN-induced reduction in darkness was much slower than in a strain
retaining these dehydrogenases (Fig. 2).
Not surprisingly, the quinol oxidase-deficient strain (Cyd/CtaII
deficient) did not behave significantly differently from the wild type
(Table 1): the quinol oxidases have not been shown to be active in the
thylakoid membrane of Synechocystis sp. strain PCC 6803 under the growth conditions used here (9).
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FIG. 2.
Changes in chlorophyll fluorescence yield in darkness
after addition of 1 mM KCN. Variable fluorescence (arbitrary units
[A.U.]) in intact cells of the wild-type ( ), SDH-deficient ( ),
NDH-1-deficient ( ), and NDH-2/SDH-deficient () strains was
measured by using very weak illumination that had no actinic effect.
KCN was added at time zero. Curves were normalized so that a value of 1 on the y axis corresponds to the maximal variable
fluorescence yield obtained upon illumination in the presence of DCMU.
Error bars are derived from three replicates each on separately grown
cultures.
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Upon removal of SDH in all background strains, the PQ pool became more
oxidized than that in the parent strains, consistent with significant
SDH activity occurring under essentially all conditions. However, a
fully reduced PQ pool was observed in darkness in the CtaI-deficient
strain even if SDH was absent, indicating that in the absence of CtaI,
sufficient electron donation to the PQ pool can occur via other
pathways (such as NDH-1 and NDH-2) to provide a fully reduced PQ pool
(Table 1).
Physiological and metabolic effects of removal of respiratory
complexes.
Removal of SDH, NDH-1, or NDH-2 led to a more oxidized
PQ pool in darkness. If all three pathways were to feed in electrons at
comparable rates, the effects of single deletions on the PQ redox state
might be expected to be less pronounced, as two pathways remain.
Therefore, it is likely that in some cases, the oxidized state of the
PQ pool is a secondary effect. To check this, we examined the pool size
of key organic acids in central metabolism (succinate, fumarate,
isocitrate, and malate) via gas chromatography-mass spectrometry
(5). Interestingly, the succinate levels in these mutants
were decreased (Table 2), with the level
in the NDH-1-deficient strain being so low that the decreased
respiratory electron flow in the NDH-1-deficient mutant could be the
consequence of low succinate levels rather than the primary lack of
NDH-1 activity. We will return to this later. However, the succinate
level in the NDH-2-deficient strain was higher and should be sufficient to not fully impair SDH activity. Indeed, in the fluorescence analysis
of NDH-2-deficient mutants, the fast phase in fluorescence induction
upon addition of KCN now interpreted to be a consequence of SDH
activity (5) was still present (8). To
further study the effect of NDH-1 on succinate levels, three strains
lacking two ndhD genes each (18) were analyzed.
The ndhD1 ndhD2 mutant exhibited decreased
respiratory rates but normal CO2 uptake levels (18). In contrast, the ndhD3
ndhD4 mutant was impaired in CO2 uptake and
the ndhD5 ndhD6 mutant is indistinguishable
from the wild type (18). In two of the three strains
lacking two NdhD copies (ndhD1 ndhD2 and
ndhD3 ndhD4), the succinate levels were
very low as well, even though they were higher than in the NDH-1-deficient strain.
Analysis of fumarate, malate, and isocitrate levels was also
informative regarding possible impairments in the various mutants. As
previously observed (5), the fumarate concentration
in the SDH-deficient strain was much more reduced than that of
succinate, leading us to conclude that the enzyme functions as an SDH
and not a fumarate reductase (Table 2). Indeed, in the NDH-1-deficient mutant, the NDH-2-deficient mutant, and two of the NdhD mutants, the
fumarate level was also very low, which is suggestive of limited SDH
activity. The level of malate in the cells was qualitatively consistent
with that of fumarate in all strains except the NDH-2-deficient strain.
This suggests that this strain is impaired in the conversion of malate.
We will come back to this later. Isocitrate levels did not change by
more than a factor of 3 in the various strains relative to the wild
type (Table 2), but it is of note that most strains carrying NDH-1
mutations had high isocitrate levels, which is suggestive of an
impairment of isocitrate conversion in these strains.
The conversions of isocitrate to 2-oxoglutarate and malate to
oxaloacetate are both dependent on the oxidized form of NAD(P). The
redox cofactor specificity of the two dehydrogenases depends on the
organism. To examine whether the malate buildup in the strain lacking
NDH-2 and the isocitrate accumulation in the strain lacking NDH-1 may
have been due to a lack of the oxidized cofactor, we determined the
pool size and redox state of NADP and NAD in the strains lacking SDH,
NDH-1, and NDH-2 in comparison with the wild type. Whereas the
wild-type and SDH-deficient strains contained significant levels of
NADP, NADPH, NAD, and NADH (Table 3), in the NDH-1-deficient and NDH-2-deficient strains, the oxidized form of
either NAD or NADP was virtually depleted. The NDH-1-deficient strain
essentially lacked the oxidized form of NADP, whereas the NAD/NADH
ratio in this strain was normal (Table 3). In contrast, in the
NDH-2-deficient strain, virtually all NAD was reduced whereas ample
oxidized NADP was present (Table 3). This confirms the NADPH
specificity of NDH-1 and the NADH specificity of NDH-2 and suggests
that NAD and NADP are not in redox equilibrium with each other.
The results presented in Table 3 suggest a reason for the isocitrate
and malate accumulation in the NDH-1-deficient and NDH-2-deficient strains, respectively, if the corresponding dehydrogenases are specific
for NADP and NAD, respectively. Indeed, in Synechocystis sp.
strain PCC 6803, the nucleotide-binding motif in isocitrate dehydrogenase suggests NADP specificity because the negative charge (Glu or Asp) at the end of the second 
sheet in this motif is missing (1). To test the cofactor specificity of
isocitrate dehydrogenase, the activity of this enzyme was determined in
a crude protein extract from wild-type Synechocystis sp.
strain PCC 6803 with either NAD or NADP as a cofactor, and the NADP
specificity of the enzyme was experimentally confirmed (Table
4). As NADP levels are very low in the
NDH-1-deficient strain (Table 3), the isocitrate dehydrogenase activity
may be limited, thus causing isocitrate accumulation and a drastic
decrease in the level of succinate in the cells (Table 2).
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TABLE 4.
Isocitrate and malate dehydrogenase activities in crude
protein extracts from wild-type Synechocystis sp. strain PCC
6803 as a function of cofactor identity and availability
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In contrast, the primary sequence of malate dehydrogenase in
Synechocystis sp. strain PCC 6803 contains a motif
consistent with NAD-binding specificity (a negative charge is present
19 residues from the third Gly of the GXGXXG nucleotide
binding motif). Indeed, a specificity of the enzyme for NAD was
observed when assaying for malate dehydrogenase activity in a crude
protein extract in the presence of either NADP or NAD (Table 4). In the NDH-2-deficient strain, the NAD level is very low and, indeed, malate
accumulation is observed (Table 2).
Respiratory rate.
The results presented thus far suggest an
important role of succinate and SDH in respiratory electron transfer.
To determine whether this is reflected in electron transport rates,
respiration was monitored as O2 consumption in darkness by
the wild-type and SDH-deficient strains grown photoheterotrophically
before the measurement. Wild-type cells respired at a rate of 34 ± 8 µmol of O2 mg of chlorophyll1
h1 in darkness. This rate was inhibited greater than 90%
by addition of 1 mM KCN and was about 50% inhibited upon addition of 5 mM malonate, an inhibitor of SDH. In the SDH-deficient strain, the respiratory rate in darkness was 9 ± 5 µmol of O2
mg of chlorophyll1 h1, reflecting 76%
inhibition relative to the wild type.
Chlorophyll fluorescence as a probe of PQ reduction rates following
KCN addition.
Chlorophyll fluorescence yield measurements can be
used as a tool with which to monitor (over)reduction of the PQ pool
indirectly. When the PQ pool becomes fully reduced, some
QA (the reduced form of the first
quinone-type electron acceptor in PSII) is formed due to reverse
electron flow (3). The midpoint potential of
QA/QA is about 80 mV more
negative than that of PQ/PQH2 (15), and therefore, the redox equilibrium constant between QA and PQ
is 20 to 30. This formation of QA when the PQ
pool becomes fully reduced can be visualized as an increase in the
chlorophyll fluorescence yield of PSII when excited by a weak
(nonactinic) measuring beam. Upon addition of KCN to the cell
suspension in the dark, the pathway of electrons out of the PQ pool to
the terminal oxidases is blocked and the donation of electrons to the
PQ pool via respiratory pathways can be measured by observing the
increase in fluorescence yield as a function of time. The SDH-deficient
strains lack the fast phase of QA formation
following KCN addition in the dark, which is a prominent phase in the
wild type (2). After the fast phase of QA
reduction, a slow phase appears that is independent of SDH and that was
attributed to PQ reduction by a strong reductant such as NADPH or NADH
(2). As the midpoint potential difference between
succinate and QA is not in favor of QA
reduction by succinate, only partial QA
formation via the fast (SDH) pathway can occur. To determine the
pathway(s) of electron flow that is responsible for the slow phase, we
examined the rise in chlorophyll fluorescence yield in various strains
lacking one or more of these pathways. The initial rate of the rise in
the SDH- and NDH-2-deficient strain was about half of that seen in the
SDH-deficient strain and approximately 1/10 of the initial rise
observed in wild-type cells (Fig. 2). In a first approximation, the
difference between the rates in the SDH-deficient strain and the SDH-
and NDH-2-deficient strain may be attributed to NDH-2 activity, and the
activity remaining in the SDH- and NDH-2-deficient strain may be
attributed to NDH-1 activity. However, in the NDH-1-deficient strain,
not only was the fast phase of the rise essentially absent (consistent
with the absence of succinate in this strain) but also the rate of the
slower phase was reduced.
To test whether the absence of the fast phase in the NDH-1-deficient
strain is due to a lack of succinate, cells were incubated with 1 mM
succinate for different times prior to measurement. A fast phase of
chlorophyll fluorescence yield increase was recovered (Fig.
3), indicating that the rapid loss of
electron donation to the PQ pool in darkness in this mutant results
from the lack of succinate. A further confirmation of this assessment
was provided when malonate was added, in addition to succinate, to the
NDH-1-deficient cells; no KCN-induced variable fluorescence was
obtained under these conditions (Fig. 3).
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FIG. 3.
Effect of succinate addition on chlorophyll fluorescence
yield as a function of time after the addition of KCN (at time zero) in
the NDH-1-deficient strain. Variable fluorescence (arbitrary units
[A.U.]) was monitored in samples without (closed symbols) and with
(open symbols) addition of 1 mM succinate or with both 1 mM succinate
and 5 mM malonate (*) 5 min prior to KCN addition. The
NDH-1-deficient culture was incubated with or without succinate for 2 (,  ) or 10 ( ,  ) min while bubbling with 3%
CO2.
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DISCUSSION |
The PQ pool redox state is affected by several respiratory
complexes.
Data collected by using the Q electrode are
enlightening with respect to the steady-state redox state of the PQ
pool under various irradiance conditions. The PQ pool of
Synechocystis sp. strain PCC 6803 is much more oxidized in
the light than in darkness. This differs from chloroplasts but is
consistent with the relative overabundance of PSI in the cyanobacterial
thylakoid membrane. Furthermore, when PSII is functionally deleted, the
steady-state PQ pool redox state is not dramatically different from
that in the wild type at any of the light intensities studied, which is consistent with a low ratio of PSII relative to PSI (23)
in Synechocystis under our growth conditions.
The effect of respiratory complexes on the redox state of the PQ pool
is best seen in darkness, as under this condition PSI does not draw
electrons out of the PQ pool. A relatively surprising result presented
in this paper (Table 1) is that deletion of any one of the three
respiratory pathways feeding electrons into the PQ pool (SDH, NDH-1, or
NDH-2) resulted in a much more oxidized PQ pool in darkness. However,
according to reference 5 and results presented in Fig. 2,
the capacity of SDH to feed electrons into the PQ pool is much larger
than that of either of the NDH complexes. Therefore, the effects of
NDH-1 and NDH-2 deletion on the redox state of the PQ pool are unlikely
to be primary effects.
Organic acid and NAD(P) reduction levels.
The effects of
deletion of SDH, NDH-1, and NDH-2 on organic acid accumulation levels
are extensive. The mutant lacking NDH-2 accumulates NADH and malate,
suggesting that the malate-to-oxaloacetate step of the tricarboxylic
acid (TCA) cycle may have been blocked by NAD+ limitation.
Indeed, malate dehydrogenase from Synechocystis sp. strain
PCC 6803 was found to be an NAD-specific enzyme (Table 4).
Interestingly, the succinate concentration in the NDH-2-deficient strain has been reduced about 10-fold compared to that in the control,
suggesting a feedback inhibition of enzymes earlier in the pathway or a
lack of flux through the TCA cycle. Sufficient succinate remains (Table
2) for SDH to impact the redox state of the PQ pool in the short term
(8), but the steady-state SDH flux is low in the
NDH-2-deficient mutant, as the fumarate level in this strain is low.
This provides a reason why the NDH-2-deficient strain has an oxidized
PQ pool in darkness.
The NDH-1-deficient mutant accumulated isocitrate and essentially
lacked acids that are downstream of isocitrate in the TCA cycle. This
suggests an inhibition of isocitrate dehydrogenase. A plausible reason
for this apparent inhibition is a lack of an electron acceptor, as the
NDH-1-deficient strain essentially lacked oxidized NADP+
and isocitrate dehydrogenase appears to be a NADP-dependent enzyme in
Synechocystis sp. strain PCC 6803. The results presented in Fig. 3 and Table 2 present a plausible explanation for the oxidized PQ
pool in darkness in the absence of NDH-1 activity: in the absence of
NDH-1, the succinate level in the cell is very low and the SDH-mediated
reduction of the PQ pool is inhibited due to a lack of substrate.
Indeed, reduction of the PQ pool resumes upon addition of succinate
(Fig. 3).
It is interesting that in Synechocystis sp. strain PCC 6803, the redox state of the NAD and NADP pools is able to fluctuate independently, as evidenced by the lack of oxidized NADP in the NDH-1-deficient strain and of oxidized NAD in the NDH-2-deficient strain, while the redox state of the other pyridine dinucleotide is
fairly close to that in the wild type. This implies that
there is no significant transhydrogenase activity in this
cyanobacterium, although open reading frames have been identified in
the Synechocystis sp. strain PCC 6803 genome sequence
that appear to code for the A and B subunits of pyridine nucleotide
transhydrogenase (slr1239 and slr1434)
(13).
Another interesting finding is that the total level of NAD(H) and
NADP(H) was variable, depending on the strain studied and the
conditions under which it was grown. The reason for this, presumably,
is that these compounds are produced via pathways starting at TCA
cycle intermediates. NAD(P) is produced from the cyclization of
glycerol and aspartate to form quinolinate. Aspartate is synthesized
from oxaloacetate via aspartate aminotransferase or possibly from
fumarate via fumarate dehydratase. Therefore, reduced levels of
fumarate and oxaloacetate are likely to lead to lower levels of NAD(P).
A final observation about the cellular levels of NAD and NADP is that
there is an order of magnitude more NADP than NAD, as determined by two
independent methods. This difference in concentration probably is
related to the fact that many important cellular reactions in the
cyanobacterium are NADP specific rather than NAD specific and that NADP
is used for storage of reductant generated by photosynthesis.
Relative activities of the respiratory complexes in the thylakoid
membranes.
The reduction kinetics of the PQ pool in darkness upon
addition of KCN were much slower in SDH-deficient strains (Fig. 1 and 2). The fast phase of the increase in the chlorophyll fluorescence yield was completely absent in the SDH-deficient strain (Fig. 2). From
the estimated redox midpoint potential of
QA/QA versus PQ/PQH2,
the amount of PQ, the amount of chlorophyll, and the rate of
fluorescence yield increase upon KCN addition, the rate of reduction of
the PQ pool in the wild type upon KCN addition was calculated to be 80 to 100 µmol of electrons (mg of chlorophyll)1
h1 (5), with a considerable potential
underestimation because of the time it takes for KCN to diffuse in and
to act. The observed respiratory rate (O2 uptake) in
wild-type Synechocystis in the dark is about 120 to 200 µmol of electrons (mg of chlorophyll)1 h1
[30 to 50 µmol of O2 consumed (mg of
chlorophyll)1 h1] (9). The
rate of PQ reduction in the SDH-deficient strain was calculated to be
10 to 30 µmol of electrons (mg of chlorophyll)1
h1 (5), and therefore, the SDH complex
accounts for the majority of the respiratory electrons donated to the
PQ pool upon dark respiration. This implies that the remaining
complexes (NDH-1 and NDH-2) account for less than half, and perhaps
only a small fraction, of the respiratory electrons reaching the oxidases.
In line with this, initial results indicated that in the
NDH-2-deficient strain, the rate of KCN-induced filling of the PQ pool
was similar to that in the control (8). Indeed, according to the rate of KCN-induced QA reduction in darkness in the
SDH-deficient strain relative to that in the SDH- and NDH-2-deficient
strain (Fig. 2), the rate of PQ pool reduction by NDH-2 complexes is about 10 µmol of electrons (mg of chlorophyll)1
h1 and therefore accounts for only 5 to 10% of the total
respiratory electrons donated to the PQ pool in darkness. This
approximation assumes that NDH-1 activity remains the same in the
presence and absence of NDH-2.
The observation that the rate of PQ pool reduction in the dark is about
half in the SDH- and NDH-2-deficient strain compared to that in the
SDH-deficient strain (Fig. 2) also implies that the rate of electron
donation by NDH-1 (plus any unknown electron donors that may be
present) to the PQ pool is similar to that of NDH-2. Therefore, the
NDH-1 contribution to respiratory electron transport is relatively
minimal (5 to 10% of the total respiratory electron transfer) in the
thylakoid membrane of Synechocystis sp. strain PCC 6803.
The large contribution of SDH in providing electrons to the PQ
pool in darkness is evidenced further by comparing the respiratory rates of the wild-type and SDH-deficient strains. The
SDH-deficient strain grown photoheterotrophically showed very little
KCN-sensitive oxygen consumption; the rate of oxygen uptake in this
strain in darkness was 9 ± 5 µmol of O2 (mg of
chlorophyll)1 h1, whereas in the presence
of KCN, about 5 µmol of O2 (mg of
chlorophyll)1 h1 remained (the latter rate
of KCN-insensitive oxygen uptake is found in all of the
Synechocystis sp. strain PCC 6803 isolates we have analyzed,
and this oxygen uptake appears to be unrelated to respiration). This
again illustrates that the contribution of other respiratory donors,
NDH-1 and NDH-2, is minimal compared to that of SDH activity.
In summary, the SDH complex has a major role in respiratory electron
flux and thereby in redox poising of the PQ pool. On the other hand,
the direct role of the NDH-1 complex in electron donation to the PQ
pool is minor, and changes in respiratory rates upon NDH-1 deletion
need to be reinterpreted in terms of succinate depletion. Finally, as
shown by evidence presented here and previously (8), NDH-2
also has a role in thylakoid PQ pool reduction that seems to be
quantitatively similar to that of NDH-1.
We thank Teruo Ogawa for the gift of the NdhB-deficient (M55) and
NdhD-deficient strains, Mark Hayes and Karl Booksh (Department of
Chemistry and Biochemistry, Arizona State University) for the use of
the CV-50W analyzer, Lokesh Joshi (Department of Plant Biology, Arizona
State University) for the use of the HPLC apparatus with fluorescence
detection, and Crispin Howitt (currently at the Division of Plant
Industry, Commonwealth Scientific and Industrial Research Organisation)
for help with some of the Q electrode experiments. UQ-1 was a gift from
David A. Ward (Department of Chemistry, University of Adelaide).
Support for this research was provided by grants from the Human
Frontiers Science Program (RG 0051/19997M) and the U.S. Department of
Agriculture National Research Initiative Competitive Program (97-35306-4881). Jason Cooley was supported by a Graduate Research Training grant from the National Science Foundation (DGE-9553456).
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Abstract
Introduction
