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Plant Physiol. (1999) 120: 293-300
Membrane Lipid Integrity Relies on a Threshold of ATP Production
Rate in Potato Cell Cultures Submitted to Anoxia1
André Rawyler*,
Danijela Pavelic,
Christian Gianinazzi,
Jacques Oberson, and
Roland Braendle
Pflanzenphysiologisches Institut, Universität Bern,
Altenbergrain 21, CH-3013 Bern, Switzerland
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ABSTRACT |
In this paper we report on our study
of the changes in biomass, lipid composition, and fermentation end
products, as well as in the ATP level and synthesis rate in cultivated
potato (Solanum tuberosum) cells submitted to anoxia
stress. During the first phase of about 12 h, cells coped with the
reduced energy supply brought about by fermentation and their membrane
lipids remained intact. The second phase (12-24 h), during which the
energy supply dropped down to 1% to 2% of its maximal theoretical
normoxic value, was characterized by an extensive hydrolysis of
membrane lipids to free fatty acids. This autolytic process was
ascribed to the activation of a lipolytic acyl hydrolase. Cells were
also treated under normoxia with inhibitors known to interfere with
energy metabolism. Carbonyl-cyanide-4-trifluoromethoxyphenylhydrazone did not induce lipid hydrolysis, which was also the case when sodium
azide or salicylhydroxamic acid were fed separately. However, the
simultaneous use of sodium azide plus salicylhydroxamic acid or
2-deoxy-D-glucose plus iodoacetate with normoxic cells
promoted a lipid hydrolysis pattern similar to that seen in anoxic
cells. Therefore, a threshold exists in the rate of ATP
synthesis (approximately 10 µmol g 1 fresh weight
h1), below which the integrity of the membranes in anoxic
potato cells cannot be preserved.
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INTRODUCTION |
O2 deprivation becomes a frequent stress for
plants submitted to unpredictable heavy rainfalls and flooding. The
diffusion of O2 to their submerged underground
organs is severely limited, so that plants must cope with hypoxic or
even anoxic conditions. Whereas ATP is produced with a high efficiency
by respiration in nongreen cells, its synthesis is much lower under
anoxia, with fermentation as the sole energy provider. For most higher
plants, the latter condition eventually becomes lethal. The
multifarious effects of O2 deprivation stress on
plants sensitive and resistant to anoxia are fairly well understood and
have been extensively reviewed in this decade (Armstrong et al., 1994 ;
Sachs, 1994; Ratcliffe, 1995; Crawford and Braendle, 1996; Drew, 1997;
Vartapetian and Jackson, 1997).
Aside from the obvious interest given to the responses of energy
metabolism, the role of macromolecules, and in particular, of gene
expression and protein synthesis, has received considerable attention
(Sachs, 1994; Drew, 1997). In contrast, the behavior of membrane lipids
under anoxia has scarcely been investigated. This is surprising knowing
how important it is for a living cell to maintain its membrane
integrity. In potato (Solanum tuberosum) tubers,
for instance, changes in membrane lipids have mostly been studied
during aging, and have been related to overall lipid unsaturation, lipid degradation, and peroxidation processes (Knowles and Knowles, 1989; Spychalla and Desborough, 1990a, 1990b; Kumar and Knowles, 1993;
Dipierro and De Leonardis, 1997). Although these effects are not
directly relevant to anoxia (Kumar and Knowles, 1996), they are
probably related to the effects likely to occur under anoxia and
postanoxia. Nevertheless, our knowledge of the mechanisms responsible
for membrane lipid alterations during anoxia is still limited. Lipid
synthesis was shown to diminish and desaturation of acyl chains to be
stopped because of their respective ATP and O2
requirements (Vartapetian et al., 1978; Brown and Beevers, 1987).
Organelle damage, mainly to mitochondria, occurs under anoxia and upon
re-aeration (Andreev et al., 1991).
Potato is an important crop with a high sensitivity to
O2 deprivation, which is mostly studied in
tubers. During the first 6 h of anoxia, the adenylate levels and
the energy charge decrease continuously, and the ATP production rate
becomes too low to sustain the basal metabolic requirement of the tuber
in spite of its ample starch reserves (Sieber and Braendle, 1991).
Membrane damage could be induced by ATP deprivation, which is suggested
by the correlation between the leakage of electrolytes from the cells
and the release of FFAs in anoxic tubers (Crawford and Braendle, 1996).
A link should exist between the energy status of the tuber and its
capability to maintain the integrity of its membrane lipids under
anoxia.
However, because of their compactness, potato tubers are not well
suited for some types of experiments. We have chosen to work with
potato cell cultures as an alternative and practical model for anoxia
studies. Cell suspensions allow an optimal diffusibility of gases and
solutes and present an inherent homogeneity. In addition, the duration
of anoxic treatments can be shortened to about 24 h.
Here we compare the changes in lipid composition and in energy
metabolism between cells under anoxia and cells treated with inhibitors
in the presence of O2. We conclude that a
threshold in the ATP synthesis rate exists, below which potato cells
become committed to hydrolysis of their membrane lipids.
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MATERIALS AND METHODS |
Chemicals
Most chemicals were high-purity products from Fluka or Sigma. Kits
were from Boehringer Mannheim and Lumac (Landgraaf, The Netherlands).
Cells
Potato (Solanum tuberosum L. cv Bintje) cells (kindly
given by Dr. J.L. Coquoz, University of Fribourg, Switzerland) were cultivated as 50-mL suspension cultures on Murashige and Skoog medium
fortified with 10 mg L1 thiamine-HCl, 0.1 mg
L1 kinetin, 3 mg L1
2,4-D, and 90 mM Suc in 250-mL conical flasks.
Cell cultures were incubated in a rotary shaker (120 rpm) at
24°C ± 1°C in the dark and renewed by a weekly inoculation of
3.5 mL of cell suspension in 50 mL of fresh medium. After transfer,
cells were allowed to grow for 4 d under the above conditions. At
this stage, they were in their mid-log phase and reached a
concentration of about 15 mg fresh weight mL1.
Cell suspensions were aseptically pooled, distributed as 6- to 8-mL (or
25-mL) aliquots in 50-mL (or 100-mL) conical flasks, treated, and
incubated as described below. The liquid surface area
available for gas exchange was 1 to 2 cm2 mL1.
Treatments of Cells
O2-free conditions were provided by
incubating and filtrating cells in an anaerobic workbench (model 1029, Forma Scientific, Marietta, OH) filled with 85%
N2, 5% CO2, and 10%
H2. The residual O2 level
(monitored with an O2 analyzer, model LF 700, Toray, Lippke, Neuwied, Germany) was <0.001%. Cells were frozen in
liquid N2 and stored at  80°C.
Modulation of energy metabolism in the presence of
O2 was achieved by aseptically feeding cells with
0.1 to 1 µM FCCP, 2 mM NaN3, 2 mM SHAM, 20 mM
DeOGlc, 2 mM IAc, or 1 to 5 mM NaF, followed by
incubation under normoxia. Ethanol, as a cosolvent for some of these
compounds, had no effect on cell biomass changes or on lipid
composition at least up to 100 mM (approximately 0.6%,
v/v). In all of the cases, cells were separated from their incubation medium by filtration through filter paper (type LS 14, Schleicher & Schuell) under reduced pressure (50-100 mbars).
Lipid Analysis
Frozen cells (60-250 mg fresh weight) were placed in a 7-mL
screw-cap glass tube, and 0.18 mL of 0.5 N HCl, 2.4 mL of
methanol, and 1.2 mL of chloroform were added. After shaking,
incubating for 10 min, and spinning (1000g) for 5 min, the
supernatant was saved. The pellet was re-extracted once as above. We
added 1.2 mL of chloroform and 1.8 mL of 0.5 M
KCl to the supernatants. After shaking, the mixtures were spun down as
described above. Pooled lower phases were filtered on glass wool and
anhydrous Na2SO4 and dried
under N2. The residue, taken up in 1 mL of
chloroform:methanol (8:2, v/v), was the total lipid extract.
Total lipids were measured on a 200-µL aliquot of the total lipid
extract, supplemented with 100 µg of nonadecanoic acid as the
internal standard. FFAs were obtained as follows: The remaining total
lipid extract (800 µL) received 100 µg of heptadecanoic acid as the
recovery standard for FFAs. The mixture was spotted as a 3-cm streak
(Linomat II, Camag AG, Muttenz, Switzerland) onto a TLC glass plate
(10 × 20 cm) coated with a 0.25-mm-thick layer of silicagel 60 (E. Merck AG, Dietikon, Switzerland). After development with petroleum
ether (boiling point = 50°C-70°C):diethylether:acetic acid
(70:30:1, v/v), the plate was briefly air-dried, sprayed with primuline
(0.01% in acetone:water, 4:6, v/v), and viewed under UV light. Bands
corresponding to the FFAs were scraped off and transferred into 7-mL
tubes together with 100 µg of nonadecanoic acid as an internal
standard.
Total lipids and FFAs were methylated in 2 mL of 5%
H2SO4:methanol under
N2 for 60 min at 85°C. After cooling, we added
1.8 mL of pentane and 2 mL of water and shook and then centrifuged the
mixture. The upper phase was washed once and transferred into glass
inserts. Fatty acid methyl esters were isothermally (190°C) separated
on a 25-m × 0.4-mm × 0.2-µm FFA phase-coated
column (Macherey and Nagel, Oensingen, Switzerland) and quantified with an AutoSystem (Perkin-Elmer).
Determination of Respiratory Activity of Cells
We measured the O2 uptake of cells in a
glass cuvette at 24°C using a Clark-type O2
electrode. The reaction medium contained 40 to 80 mg of cell fresh
weight in 5 mL of Murashige and Skoog medium plus 90 mM
Suc.
Measurement of ATP
ATP was extracted at 2°C from 250 mg of frozen cells by
homogenization with 6 mL of 6% perchloric acid for 30 s in a
Potter glass homogenizor. After centrifugation for 10 min at
24,000g at 2°C, the supernatant was neutralized with 5 M ice-cold
K2CO3 in an ice bath.
KClO4 was removed by spinning as described above. We determined ATP immediately with the luciferin-luciferase system, using a biocounter (model 2500, Lumac) as described by Sieber and
Brändle (1991).
Determination of Fermentation End Products
We measured the amounts of ethanol, lactate, and Ala in cells and
in their corresponding filtrates. The ethanol and lactate were
extracted from frozen cells (0.5-2 g) in the presence of 3 to 6 mL of
6% perchloric acid as described above. We placed the filtrates in
tightly closed vessels, inactivated them at 80°C for 15 min, and then
froze and stored them until use. We performed the spectrophotometric
determination of ethanol and lactate at 365 nm with enzymatic test kits
(Boehringer Mannheim). We used the supernatants (centrifuged at
24,000g for 10 min) of cell homogenates and of the
corresponding filtrates for Ala determination. After protein
precipitation, we readied the amino acids for reaction with
phenylisothiocyanate (Bidlingmeyer et al., 1984) and analysis by HPLC
(Davey and Ersser, 1990) on a Novapak C18 column (150 × 3.9 mm; 4 µm, porosity 60 Å), using norleucine as an internal standard.
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RESULTS |
Time Course of Biomass Change
Under normoxic conditions cell biomass increased regularly, almost
doubling over a 24-h period (Fig. 1).
Anoxic cells behaved similarly for up to 6 h, but then their
biomass started to decrease, reaching the initial 100% value after
about 12 h. This decrease continued steadily for up to 24 h
(Fig. 1). The filtration behavior of cells and their aspect changed
dramatically after 12 h of anoxia. Before this time, anoxic cells
were easy to filtrate and the packed cells had the typical yellowish
and granulous aspect of normoxic cells. On the other hand, filtration
of anoxic cells became increasingly tedious after 12 h, and packed
cells appeared dark yellow to gray with a pasty consistency.
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| Figure 1.
Relative change of cell biomass during incubation
time under normoxic ( ) and anoxic ( ) conditions. The initial
(100%) value ranged between 15 and 25 mg cell fresh weight
mL1. Each point is the average of up to five independent
determinations.
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Time Course of Lipid Hydrolysis under Anoxia
When potato cells were incubated under anoxia, FFAs were released
in a time-dependent way. Lipid hydrolysis started after a time lag of
about 10 to 12 h, and reached about 60% 12 h later (Fig.
2). In contrast, the lipid composition of
normoxic cells did not change during this period, as indicated by the
stable low level (approximately 1%) of endogenous FFAs (Fig. 2). To
check whether lipids would be released from cells during the
treatments, we performed control experiments to compare the lipid
content of filtrates obtained from cells incubated for 24 h under
normoxia with those under anoxia. We found a maximum of 1% and
6% of the total cell fatty acids in the filtrates of normoxic and
anoxic cells, respectively. However, most of these lipids could be
ascribed to small cell fragments (especially under anoxia) having
leached through the filter and not to the net release of lipid
molecules into the medium.
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| Figure 2.
Change in the hydrolysis extent of potato cell
lipids during incubation time under normoxic () and anoxic ()
conditions. The release of FFAs is expressed as a percentage of the
total fatty acid content of cells. Each point is the average of up to
10 independent determinations. The total fatty acid content of potato
cell lipids (100%) was 25.3 ± 1.2 µmol fatty acids
g1 fresh weight of which 76% ± 3% were phospholipids,
14% ± 2% were glycolipids, and 10% ± 1% were neutral, apolar
lipids (mostly triacylglycerols and acylated sterols).
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We also determined the acyl chain composition of total lipids and of
the FFAs over a 24-h period (Fig. 3). The
acyl composition of total lipids was very constant with time and
essentially similar in normoxic and anoxic cells. In all cases, the
three major fatty acyl chains were 18:2 (linoleic), 18:3 (linolenic),
and 16:0 (palmitic) acids, with minor amounts of 18:1 (oleic) and 18:0
(stearic) acids (Fig. 3A). On the other hand, the acyl composition of
the FFAs changed dramatically with the progress of lipid hydrolysis
(see legend to Fig. 2). During the initial time lag of approximately 12 h, the low level of FFAs in the anoxic cells (Fig. 3B) was constant and essentially accounted for by saturated chains (16:0 and
18:0). This was also the case for the normoxic cells. The absence of
lipid hydrolysis was thus reflected by the low DBI of the small
endogenous FFA pool (see legend to Fig. 3) generated by neosynthesis
and elongation (Harwood, 1988). When hydrolysis started in the anoxic
cells, the acyl composition of the FFAs switched rapidly toward
increased unsaturation and became similar to that of the total lipids
(compare with the final fatty acid levels in Fig. 3). In this case,
lipid hydrolysis was reflected by a much higher DBI value for the FFAs
released, which was comparable to that of the total lipids (see legend
to Fig. 3).
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| Figure 3.
Acyl chain composition (in mol %) of total cell
lipids (A) and of the FFAs released (B) in potato cells incubated under
anoxia. Each data point is the average of two experiments (agreeing
within ±1%). The acyl composition of total lipids in normoxic cells
is essentially similar to that of anoxic cells and is therefore not
detailed here. By definition, DBI = ( [N × mol % fatty acid ])/100, where N is the number of
double bonds in each fatty acid molecule. Here, DBI is thus equal to
(0 × mol % 16:0 + 0 × mol % 18:0 + 1 × mol % 18:1 + 2 × mol % 18:2 + 3 × mol % 18:3)/100. In A, the DBI
values were 1.788 at 0 h, 1.804 at 12 h, and 1.986 at 24 h. In B, the DBI values were 0.575 at 0 h, 0.734 at 12 h, and
1.912 at 24 h. The constancy of the fatty acid composition of
total lipids in anoxia-treated cells (A) is in agreement with the
requirement of acyl chain desaturation for O2 (Harwood,
1988).
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Nature of the Enzyme Responsible for Lipid Hydrolysis
We investigated the nature of the enzyme responsible for the lipid
hydrolysis reported in Figure 2 and summarized our results in Tables I
and II. On a molar basis, each phospholipid hydrolyzed gave rise to two
FFAs, and lysophospholipids were not detected (Table
I). When we assayed a cell extract for
esterase activity in a Ca2+-free buffer using
p-nitrophenylpalmitate as a substrate (Galliard, 1971), a
substantial amount of palmitic acid was produced (Table I). Finally,
after significant lipid hydrolysis, the acyl composition of released
FFAs and that of total lipids matched closely (Table II; see also Fig. 3). Together, these
data suggest that the FFA release described in Figures 2 and 3 is due
to a LAH.
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Table I.
Evidence for the occurrence of a lipolytic acyl
hydrolase activity in potato cells
Phospholipids were separated from other lipids using silica gel
cartridges. After loading the total lipid extract and washing with
acetone, phospholipids were eluted with methanol, and either separated
by TLC or transmethylated and analyzed by GC. Soluble cell extracts
were prepared by homogenization of normoxic potato cells (0.2-0.5 g
fresh wt) in 1 mL of 0.1 M potassium phosphate buffer (pH
7.5) using a Potter and spinning at 15,000g for 15 min. The
supernatants were assayed for activity in a Ca2+-free
phosphate buffer (Galliard, 1971).
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Energy Production Linked to the Formation of Fermentation Products
under Anoxia
We measured the production of ethanol, lactate, and Ala together
with the level of ATP in anoxic potato cells. There was a 45% decrease
in the ATP level after 6 h of anoxia (Table
III). Longer incubation times led to a
further depletion of the ATP pool, which was reduced to 10% of the
initial normoxic value after 12 h. Ethanol accumulated steadily
during 24 h up to a level 4.5-fold higher than that of lactate or
Ala. Lactate rose only during the first 6 h of anoxia and then
remained stable. Ala increased up to 18 h, then stabilized at the
same level as lactate. We calculated the ATP synthesis rates as
described in the legend of Table III. Under normoxia the synthesis
rates were high and insensitive to the method (via invertase or Suc
synthase) of Suc metabolism (Plaxton, 1996). These rates were
drastically reduced during the first 6 h of anoxia to 1.5% and
2.8% (via invertase or Suc synthase, respectively) of the maximum
theoretical values attainable under normoxia; they diminished further
in the next 6 h, and finally showed an essentially constant value
between 12 and 24 h.
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Table III.
Levels of ATP and fermentation products generated
during incubation of potato cells under anoxia, and the
corresponding calculated ATP synthesis rates
Levels were measured as described in ``Materials and Methods'' and
are the average (±SD) of n = 5 determinations. Except for the zero-time samples, the rates of ATP
synthesis were calculated by: (a) assuming an equimolar correspondence
between each fermentation end product and ATP, (b) summing all end
products after subtraction of their zero-time levels, and (c)
expressing results on a 1-h basis. The ATP synthesis rate at time 0 was
calculated from the O2 uptake of normoxic cells (96 ± 11 µmol O2 g fresh wt1 h1,
n = 11) and assuming the production of 38 or 40 ATP
molecules per 6 O2 molecules consumed. Cases a and b
correspond to the feeding of the glycolytic pathway by the invertase or
by the Suc-synthase, respectively (Plaxton, 1996).
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Metabolic Inhibitors Can Induce FFA Release under Normoxia
The results of Table III point to the obvious and important role
played by energy metabolism. We attempted to mimic the effect of anoxia
on membrane lipid degradation (Fig. 2) by treating potato cells under
normoxia with different inhibitors of: (a) the glycolytic pathway
(DeOGlc + IAc), (b) the mitochondrial electron transport in the Cyt
chain (azide) and in the alternative pathway (SHAM), and (c) the
membrane-linked ADP phosphorylation (FCCP). After incubation for 12 and
24 h, we determined the extent of lipid hydrolysis and the
relative change in biomass (Fig. 4). FCCP
had no effect on these parameters (as compared with normoxic controls), although it stimulated the O2 uptake rate by
50%. The DBI of the FFA fraction was <1. However, lipid hydrolysis
occurred in a time-dependent manner when the mitochondrial inhibitor
couple, azide plus SHAM, or the glycolytic inhibitor couple, DeOGlc
plus IAc, was used. The acyl chain composition and the DBI of the FFAs
released under normoxia (Fig. 4) resembled that obtained between 12 and
24 h of anoxia (Fig. 3B) and was accompanied by a decrease in the
relative cell biomass. Glycolytic inhibitors were more efficient than
mitochondrial inhibitors for both parameters (Fig. 4). Furthermore,
both the change in the relative biomass and the extent of lipid
hydrolysis obtained after 12 and 24 h of normoxia in the presence
of inhibitors were consistent with those observed during the anoxic
incubation of potato cells (compare Fig. 4 with Figs. 1 and 2). NaF
(1-5 mM) could not be substituted to the glycolytic
inhibitors. Finally, total lipids were constant, either on a fresh
weight basis or as absolute amounts in a fixed cell-suspension volume.
Lipid hydrolysis was thus achieved without further processing (e.g.
 -oxidation) of the hydrolysis products. We must also point
out that membrane degradation occurred independently in the absence
(Figs. 2 and 3) or presence (Fig. 4) of O2.
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| Figure 4.
FFA release and relative biomass changes (inset)
in potato cells incubated under normoxia for 0, 12, and 24 h (no
addition, open bars), or in the presence of FCCP (0.1 µM,
black bars), azide plus SHAM (2 mM each, light gray bars),
and DeOGlc plus IAc (20 and 2 mM, dark gray bars).
Figures above bars represent the DBI of the corresponding FFA pools.
Data represent the mean value of two independent experiments.
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The Relation between Lipid Hydrolysis and ATP Synthesis Rates
One can see from the results in Figure 2 and Table III that lipid
hydrolysis occurred under anoxia only after a threshold time of
approximately 12 h, when the energy production was rather low. We
have explored this relation over a wide range of ATP synthesis rates,
and the results are presented in Figure
5. Under anoxia, lipid hydrolysis did not
occur provided that the rates were higher than about 10 µmol
g1 fresh weight h1,
depending on how Suc could be metabolized. Below this threshold value
anoxic cells became committed to lipid hydrolysis. When we treated
potato cells with inhibitors under normoxia, we calculated the
ATP-synthesis rates from the residual O2-uptake
rates. As shown in Figure 5, these rates were indeed reduced by SHAM,
FCCP, and azide, but each of the three compounds failed to induce lipid degradation when applied alone. In contrast, the combination of azide
plus SHAM could elicit lipid hydrolysis in normoxic cells; the
glycolytic inhibitors were even more efficient in this respect. We
obtained similar results with 12 µM antimycin A instead
of sodium azide (data not shown).
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| Figure 5.
Relation between calculated ATP synthesis rate
(logarithmic scale) and the extent of lipid hydrolysis (linear scale)
in potato cells. In the first series of experiments ( ), cells were
incubated for 0, 6, 12, 18, and 24 h under anoxia and the levels
of ethanol, lactate, and Ala were determined together with lipid
hydrolysis. From these data, ATP synthesis rates were calculated
assuming that Suc is hydrolyzed via the invertase pathway, which gives
a net yield of two ATP per hexose (continuous line), or via the
Suc-synthase pathways, which can yield between three (dashed line) and
four ATP (dotted line) per hexose, and plotted against the
corresponding extent of lipid hydrolysis. In the second group of
experiments (large symbols), cells were treated for 12 h under
normoxia with 2 mM azide ( and  ), 2 mM
SHAM ( and  ), or azide + SHAM (2 mM each,  and
 ), DeOGlc + IAc (20 and 2 mM, and ,
respectively), and FCCP (0.1 and 1 µM,  and  ,
respectively). ATP synthesis rates were then calculated from the
O2 uptake rates (for SHAM and/or azide, for DeOGlc + IAc
and for FCCP) measured after 12 h of incubation, and plotted
against lipid hydrolysis extents. These calculations were based on an
O2-to-hexose ratio of 6 and on the maximal theoretical
yield of ATP:hexose derived from metabolic pathways. Filled and open
symbols correspond to the two extreme Suc utilization scenarios (see
above). In the case of the combined azide + SHAM treatment, italic
numbers correspond to the number of mitochondrial "phosphorylation
sites" used for the calculation. FW, Fresh weight.
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DISCUSSION |
We studied the changes in biomass, lipid composition, fermentation
end products, ATP levels, and synthesis rates in cultivated potato
cells submitted to anoxia stress. We then compared the changes with
those obtained in cells treated in the presence of O2 with inhibitors that interfered with energy
metabolism. The cells exhibited a two-phase behavior with respect to
anoxia. The first phase (0-12 h) reflects the survival of cells that
temporarily coped with the stress condition by switching to
fermentation. There was no net loss in cell biomass, and cell viability
was maintained (Fig. 1), although both the level and the rate of ATP synthesis were depressed to lower values because of fermentations (Table III). Cell membranes were still intact, as suggested by the
absence of lipid degradation (Figs. 2 and 3B). The second phase was an
autolytic process starting approximately 12 h after the onset of
anoxia. It was characterized by a decrease in cell biomass (Fig. 1), a
further reduction of ATP levels and production rates (Table III), and a
massive accumulation of FFAs (Figs. 2 and 3B) at the expense of
phospholipids (Table I).
The release of two fatty acids per phospholipid and the absence of
lysophospholipids (Table I) suggest that an LAH was responsible for the
extensive lipid degradation observed in Figure 2. This was confirmed by
the close similarity of the acyl compositions of total lipids and FFAs
(Table II; see also the 24-h point in Fig. 3B). This would not be the
case if fatty acids had been released from a single position of the
glycerol by a phospholipase A1 or A2 because 16- and 18-carbon fatty
acids show a preferential localization on the
sn2 and
sn1 carbon atoms, respectively, of
the glycerol backbone of plant phospholipids (Galliard, 1973).
Furthermore, cell extracts hydrolyzed
p-nitrophenylpalmitate; Ca2+ was not
required for this activity, in contrast to phospholipases A. As shown
in Table I, the extract from 1 g of cell fresh weight would form
5.3 µmol of palmitic acid from p-nitrophenylpalmitate in
12 h. In anoxic cells an equivalent FFA release would correspond to a hydrolysis extent of about 20%, which falls within the range of
values shown in Figure 2. The autolytic phase of anoxic potato cells
involves the activation of an LAH. This enzyme had already been
implicated in the response of plant cells to mechanical or pathogenic
wounding (Racusen, 1984; Slusarenko et al., 1991; Farmer and Ryan,
1992) and in the formation of autophagic vacuoles after Suc starvation
(Aubert et al., 1996).
Because FFAs account for up to 60% of total lipids after 24 h of
anoxia (Fig. 2), they must arise mainly from phospholipids and
glycolipids, which make up 76% and 14%, respectively, of total cell
lipids. This suggests that lipid degradation eventually affects most
cellular membranes. The very sharp DBI change in the FFAs (Fig. 3B) is
more illustrative of a general hydrolytic process than of selective
degradation, such as that shown by cell membrane autophagy under Suc
starvation (Aubert et al., 1996). FFAs remain in the membrane in which
they were generated (Roelofsen, 1982); our control experiments agree
with this view. Aside from the detergent/fusogen properties of FFAs,
there is another consequence of LAH action: removal of the polar
headgroups of membrane lipids, which dissolve in the aqueous medium.
Clearly, membrane properties must be strongly altered by these
structural changes. In potato tubers the anoxia-induced accumulation of
FFAs was correlated with an increased electrolyte leakage, which
reached almost 100% when the FFA level was 6% to 7% of total lipids
(Crawford and Braendle, 1996). The change in the aspect of cell pellets
after 12 h of anoxia may well reflect these alterations in
membrane composition and structure.
Cell behavior during anoxia depends on how the energy requirements are
fulfilled when the energy supply is exclusively supported by
fermentation. The metabolic survival strategy of cultivated cells
includes several aspects. ATP-consuming processes of lower priority can
be suppressed, as illustrated by the arrest in biomass increase after
6 h of anoxia (Fig. 1). Entering into anaerobic retreat can slow
down the ongoing ATP-consuming processes (Pradet and Raymond, 1983).
The metabolism of Suc via Suc synthase rather than via invertase, for
example, can improve residual ATP production (Stitt and Steup, 1985;
Sachs, 1994), resulting in an increase in the ATP net yield of 1.5- to
2-fold, according to pyrophosphate availability (Mertens, 1991; Stitt,
1998). Finally, the efficiency of ATP processes can be enhanced, e.g.
by raising the H+-to-ATP ratio of plasmalemma and
tonoplast H+-ATPase pumps (Slayman, 1980). This
strategy met with some success during the first 12 h of anoxia,
because the cells did not undergo any membrane damage during this
period, although both the ATP levels and the synthesis rates were
reduced (Table III).
The ATP content of a cell cannot sustain itself for more than 1 to 2 min (Roberts et al., 1984). On a time scale of hours, the main
determinant of energy balance must be the rate of ATP synthesis rather
than the ATP level (Tadege et al., 1998). There is a threshold in
fermentation and ATP synthesis rates below which survival is prejudiced
(Xia et al., 1995). Our results are in agreement with this view and
suggest that the integrity of membrane structure relies on a low
threshold value of the ATP production rate of approximately 10 µmol
g1 fresh weight h1.
This value is about 1.5% of the maximal theoretical ATP synthesis rate
under normoxia (Table III; Fig. 5). It is generally agreed, however,
that only about half of the mitochondrial H+
electrochemical potential is used to drive ATP synthesis. Thus, the maximum reachable ATP yield per Glc is probably closer to 20 than to 38. The threshold of 10 µmol ATP
g1 fresh weight h1
would then be 3% of the maximum ATP synthesis rate. The most important
aspect of this threshold is not the sharpness of its absolute value,
but rather its existence and its low value compared with control rates.
However, this seemingly low value is higher than the cost of membrane
maintenance in nongrowing plant cells, estimated to be
approximately 1.7 mg Glc g1 dry weight
d1 by Penning de Vries (1975), which is
equivalent to approximately 0.16 µmol ATP g1
fresh weight h1. These modest rates merely
emphasize the sobriety of membrane maintenance mechanisms.
Accordingly, treating normoxic cells with metabolic inhibitors should
confirm the involvement of the energy production rate in the
maintenance of membrane integrity. Cells can be fed with an uncoupler
(e.g. FCCP). This will suppress membrane-linked
phosphorylations, whereas substrate-level phosphorylations of the
glycolytic pathway and of the tricarboxylic acid cycle will still occur
and will even be stimulated by the increased O2
uptake rate. Clearly, uncoupling conditions allow sufficiently high
rates of substrate-level phosphorylation (60-144 µmol ATP
g1 fresh weight h1) to
prevent any lipid hydrolysis during the whole incubation period (Figs.
4 and 5).
Azide (or antimycin A) and/or SHAM can inhibit mitochondrial electron
transport pathways (Vanlerberghe and McIntosh, 1997). Membrane lipid
degradation (Figs. 4 and 5) is then strictly dependent on the presence
of both inhibitors, suggesting that degradation occurs only when all
membrane-linked redox reactions in mitochondria are inhibited. DeOGlc
plus IAc can block the glycolytic pathway (Jans et al., 1997). This
treatment is more efficient than one that inhibits respiration (Figs. 4
and 5), probably because of the upstream localization of the inhibition
sites. These inhibitors are usually used for short-term studies, and
with isolated enzymes and organelles rather than with whole cells.
Moreover, IAc is an oxidizing agent of sulphydryl groups. Secondary
effects might contribute to overall inhibition during the 12-h
incubation period and shift the lipolytic response toward higher values
of the rate of ATP synthesis. The lack of effect of 1 to 5 mM NaF as an alternative glycolytic inhibitor is presumably
due to restricted penetration within the cells. Nevertheless, we
ascribe the release of FFAs in normoxic cells treated with glycolytic
and respiratory inhibitors to a threshold in the rate of ATP synthesis,
the value of which is comparable to that observed under anoxia (Fig.
5). In addition, the mere absence of O2 does not
trigger lipid hydrolysis. Therefore, no O2 sensor
is required in this process.
The similarity of the extent of lipid hydrolysis and of the acyl
composition of FFAs obtained under anoxia (Figs. 2 and 3) and normoxia
with inhibitors (Fig. 4) suggests that these hydrolytic processes stem
from a common mechanism. The mechanism is triggered when the rate of
ATP synthesis reaches a low threshold value (Fig. 5) and eventually
activates an LAH. Clearly, membrane lipid integrity is a key factor in
the survival of both anoxia-intolerant (this paper) and resistant
plants (Henzi and Braendle, 1993). Accumulation of new mRNAs and a
shift in protein synthesis (Sachs, 1994) would be useless without an
intact membrane network. Finally, our results emphasize the prime
importance of the events occurring under anoxia, as compared with those
taking place after reaeration (Pfister-Sieber and Braendle, 1994).
Indeed, with the hydrolysis values reported in Figure 2, a cell is
dead. A further degradative process (e.g. peroxidation) would be purely
(bio)chemical, without any physiological relevance for the
actual cell. Overexpressing the key enzymes of the fermentation
pathways can increase the energetic competence of cells. We are
currently establishing potato cell lines with enhanced fermentation
capacity from transformed plants. We hypothesize that such cells will
not undergo lipid degradation altogether or will delay it
significantly.
| |
FOOTNOTES |
1
This work was supported by the Swiss National
Science Foundation.
*
Corresponding author; e-mail andre.rawyler{at}pfp.unibe.ch; fax
41-31-332-2059.
Received December 8, 1998;
accepted February 7, 1999.
| |
ABBREVIATIONS |
Abbreviations:
DBI, double-bond index.
DeOGlc, 2-deoxy-D-Glc.
FCCP, carbonyl-cyanide-4-trifluoromethoxyphenylhydrazone.
FFA, free fatty
acid.
IAc, sodium iodoacetate.
LAH, lipolytic acyl hydrolase.
SHAM, salicylhydroxamic acid.
X:Y, a fatty acyl group containing X carbon
atoms and Y double bonds (cis unless otherwise
specified) .
| |
ACKNOWLEDGMENT |
We are grateful to Urs Kaempfer (Chemistry Department,
University of Bern, Switzerland) for his help with the HPLC analysis of
amino acids.
| |
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Top
Abstract
Introduction