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Tree Physiology 34, 229–240
doi:10.1093/treephys/tpu013
Research paper
Steady sucrose degradation is a prerequisite for tolerance
to root hypoxia
Satoshi Kogawara1,2,3, Takashi Yamanoshita1,2, Mariko Norisada1,2 and Katsumi Kojima1,2
1Asian
Natural Environmental Science Center, The University of Tokyo, Tokyo 113-8657, Japan; 2Japan Science and Technology Agency, CREST, Tokyo 113-8657, Japan;
author ([email protected])
3Corresponding
Received July 29, 2013; accepted February 6, 2014; published online March 18, 2014; handling Editor Heinz Rennenberg
We investigated the role of glycolysis and sucrolysis in the difference in tolerance to root hypoxia between two Myrtaceae
tree species, Melaleuca cajuputi (which shows superior tolerance to root hypoxia) and Eucalyptus camaldulensis (which does
not). Analysis of the adenylate energy charge (AEC) in roots subjected to a 4-day hypoxic treatment (HT) in hydroponic culture
revealed that the interspecies difference in tolerance corresponds to the ability to maintain energy status under root hypoxia:
AEC was reduced by HT in E. camaldulensis, but not in M. cajuputi. The energy status in HT roots of E. camaldulensis was
restored by feeding of glucose (Glc) but not sucrose (Suc). These data provide evidence that low substrate availability for
glycolysis resulting from an impairment of sucrolysis suppresses ATP production under hypoxic conditions in this species.
Measurements of the rates of O2 consumption and CO2 production in roots indicated that E. camaldulensis, but not M. cajuputi, failed to activate fermentation in HT roots. These results cannot be attributed to enzymatic dysfunction, because no
inhibition of main glycolytic and fermentative enzymes was observed in both species, and Glc feeding had a beneficial effect
on AEC of HT roots of E. camaldulensis. The impairment of sucrolysis was demonstrated by inhibited soluble acid invertase
activity in HT roots of E. camaldulensis. In contrast, there was no inhibition in all sucrolytic enzymes tested in HT roots of
M. cajuputi, suggesting that steady Suc degradation is essential for maintaining high energy status under root hypoxia. We
conclude that root sucrolysis is one of the essential factors that determines the extent of tolerance to root hypoxia.
Keywords: flooding, glycolytic flux, invertase, oxygen deficiency, sucrose synthase.
Introduction
Reforestation is urgently needed to restore biomass production
on degraded lands. Degraded lands, including wetlands such as
swamps and marshes, where flooding impairs the biomass productivity of planted trees, exist in various locations worldwide
(Pezeshki 2001, Effler and Goyer 2006). Reforestation of
degraded wetlands requires flood-tolerant tree species. Soil
flooding results in root hypoxia because of the low diffusion
rate of gases in water and therefore low O2 supply (Armstrong
1979). Since O2 deficiency leads to reduced ATP production by
oxidative phosphorylation in mitochondria, root hypoxia is likely
to cause energy deficiency in plant roots (Ricard et al. 1994,
Vartapetian and Jackson 1997). Flood-tolerant tree species are
assumed to possess metabolic mechanisms to prevent energy
deficiency during root hypoxia (Pezeshki 2001).
One possible mechanism for maintaining the energy status
under hypoxia is the activation of fermentation (Tadege et al.
1999, Ismond et al. 2003), which involves induction of the activity
of enzymes such as alcohol dehydrogenase (ADH; EC 1.1.1.1) and
pyruvate decarboxylase (EC 4.1.1.1). Previously, we ­compared the
responses to root hypoxia (by using hydroponic culture with N2
bubbling) in two Myrtaceae tree species, Melaleuca cajuputi
Powell and Eucalyptus ­camaldulensis Dehnh. (Kogawara et al.
2006). We found that the growth of E. camaldulensis, but not
© The Author 2014. Published by Oxford University Press. All rights reserved. For Permissions, please email: [email protected]
230 Kogawara et al.
M. cajuputi, was inhibited, although ADH activity was induced
similarly under hypoxia in the roots of both species (Kogawara
et al. 2006). We speculated that factors other than activation of
fermentative enzymes determine the difference in root hypoxia
tolerance in these species.
For plants to take advantage of hypoxia-induced activation
of fermentative enzymes, glycolytic flux must be sustained or
enhanced to keep up with the demand for substrate (Bouny
and Saglio 1996), which may be achieved by an increase in the
activities of glycolytic enzymes (Rivoal et al. 1989, Fox et al.
1995, Bouny and Saglio 1996). In maize roots, hypoxic treatment (HT) increases hexokinase activity, which improves the
energy status and survival rates of root tips under anoxia
(Bouny and Saglio 1996). Using 13C-labeling, we demonstrated
that HT increased the incorporation of photoassimilates into
the water-soluble carbohydrate fraction of M. cajuputi roots,
but did not change the total carbon pool size of this fraction,
which indicates an increased carbohydrate turnover in the
respiratory metabolism (Kogawara et al. 2006). Therefore, the
high tolerance of M. cajuputi to root hypoxia may depend on an
enhanced glycolytic flux into fermentation.
In plant roots, glycolysis substrates, namely glucose (Glc),
fructose and UDP-Glc, are supplied mainly via sucrose (Suc)
degradation, in which invertase (INV; EC 3.2.1.26) and sucrose
synthase (SuSy; EC 2.4.1.13) play a pivotal role. Both INV and
SuSy have several isoforms with different subcellular locations,
which together regulate Suc partitioning between respiration,
biosynthesis of structural carbohydrates and carbohydrate
storage (Sturm and Tang 1999). Anoxic or hypoxic stress influences Suc degradation through inhibition of INV activity and
promotion of SuSy activity, and stress sensitivity differs
between isoforms (Germain et al. 1997, Zeng et al. 1999,
Albrecht and Mustroph 2003, Gharbi et al. 2007); anoxia also
affects the location of a SuSy isoform in maize roots (Subbaiah
and Sachs 2001). These reports suggest that hypoxic or anoxic
stress may alter substrate availability for glycolysis. Whether
the stability of substrate supply for glycolysis by INV and SuSy
isoforms under root hypoxia is associated with interspecies
differences in hypoxia tolerance remains to be established.
Previously, we found that Suc accumulated in hypoxic roots
of E. camaldulensis, whereas hexoses but not Suc accumulated
in M. cajuputi (Kogawara et al. 2006). These accumulation patterns may be derived from species traits in sucrolysis and/or
glycolysis under hypoxic conditions: it is presumed that an
­inhibition of sucrolysis may decrease substrate availability for
glycolysis in hypoxic roots of E. camaldulensis but not of
­
M. cajuputi.
We hypothesized that the ability to maintain sucrolytic activity associated with enhanced glycolytic activity determines the
difference in tolerance to root hypoxia in these species. We
first investigated the effect of hypoxia on root energy status
and then examined the effects of feeding sugars, Glc or Suc, to
Tree Physiology Volume 34, 2014
the root on its energy status in hypoxia. We found that feeding
of Glc but not Suc recovered the energy status of hypoxic roots
of E. camaldulensis to the level of the control roots.
Subsequently, we evaluated the response of sugar consumption rate (SCR), as an index of glycolysis/fermentation flux, to
hypoxia by measuring the CO2 production and O2 consumption
of roots. In order to investigate the metabolic background of
the observed results, we examined the effects of hypoxia on
the activities of sucrolytic and glycolytic enzymes and found
that inhibition of sucrolysis may be a major factor in the energy
status decline under hypoxia in E. camaldulensis.
Materials and methods
Plant materials and HT
Seeds of M. cajuputi were obtained from a lowland forest in
Narathiwat Province (lat. 6°30′N, long. 101°45′E), Thailand.
Seeds of E. camaldulensis (lot 19708) were kindly provided by
the Australian Tree Seed Centre, CSIRO Forestry and Forest
Products (Canberra, ACT, Australia). The seeds of each species
were sown in acid-washed sand and germinated in a temperature-controlled greenhouse (30 °C for 16 h and 25 °C for 8 h;
natural light). After germination, seedlings were grown for ~3
months with a nutrient solution supply as described in our previous study (Kogawara et al. 2006). Seedlings of a similar size of
10 cm in height were gently taken out from the seedling beds,
and their roots were washed to remove the sand. The seedlings
were transplanted into hydroponic culture in 12-l plastic containers filled with nutrient solution described previously (Kogawara
et al. 2006). The containers were covered with PVC lids with
holes to allow the shoots to protrude. The nutrient solution was
aerated at 150 ml min−1 and replenished every 3 days.
After acclimatization to the hydroponic culture for 1 month,
seedlings were transplanted into new containers for HT. The
sizes of the seedlings at the time of transplanting were 30 and
20 cm for M. cajuputi and E. camaldulensis, respectively. The O2
concentration in the nutrient solution was controlled as previously described (Kogawara et al. 2006). Hypoxic treatment
was initiated by bubbling N2 through the solution at 150 ml min−1.
The nutrient solution for control seedlings was aerated as during the acclimatization period. The gas tightness of the containers was enhanced with sponges around the root collars of each
seedling and with rubber tapes between the lids and the containers. The O2 concentration was measured every day with a
galvanic electrode (9520-10D, Horiba, Kyoto, Japan) connected
to a dissolved O2 meter (D-55, Horiba), and was within the
same range as in our previous study, i.e., <0.25 mg l−1 in HT
and >8.0 mg l−1 in the control.
Energy status of HT roots with or without sugar feeding
We analyzed the energy status of HT roots and then evaluated
the effects of sugar feeding on root energy status to i­nvestigate
Steady sucrose degradation in hypoxic roots 231
whether substrate availability is the limiting factor for energy
production in roots under hypoxia. Glucose or Suc was used
for sugar feeding. Six containers, each holding five seedlings
per species, were provided for the experiment. Among them,
two containers (one per treatment) were used for non-feeding,
and four containers (two per treatment) were used for sugar
feeding. For each treatment with sugar feeding, one container
was used for Glc feeding and the other one for Suc feeding.
After 3 days of HT, Glc (2 mM) or Suc (1 mM) was dissolved in
the nutrient solution for sugar feeding. To inhibit microbial
propagation, the nutrient solution was circulated in a hose loop
through a UV sterilizer (Turbo-Twist 3X, Energy Savers, Carson,
CA, USA) for 15 min every hour.
At 4 days after HT, i.e., 1 day of sugar feeding, the lateral
roots of each seedling were cut off, immediately frozen in liquid
N2, freeze-dried and weighed. The samples (10–20 mg) were
ground into powder with a mortar and pestle under liquid N2,
followed by further grinding in 1.2 ml of 120 mM 5-sulfosalicylic acid. The homogenates were centrifuged at 17,500g at
4 °C for 10 min. The supernatants were neutralized with
0.95 M KOH (107 µl) and centrifuged again. Debris and
­proteins in the supernatants were excluded by ultrafiltration on
UFC3 LGC 00 filters (Millipore, Billerica, MA, USA). Filtrate
­aliquots (100 µl) were mixed with 100 mM KH2PO4 (600 µl),
and adenine nucleotides were derivatized to 1,N6 -ethenoadenine
nucleotides by the addition of 45% chloroacetaldehyde (60 µl).
The reactions were performed at 95 °C for 10 min. The
­ erivatives were quantified by high-performance liquid chrod
matography as described by Yamanoshita et al. (2005). Each
derivative (ATP, ADP, AMP) was separated by using an analytical reversed-phase column (5 µm ODS-C18, 150 mm × 6.0 mm
internal diameter, Shimadzu, Kyoto, Japan), detected by a fluorescence detector (RF-10A, Shimadzu), using an excitation
wavelength of 285 nm and an emission wavelength of 410 nm,
and quantified on the basis of its peak area on the chromatogram. Adenylate energy charge was calculated as
AEC = ( ATP + 0.5ADP ) / ( ATP + ADP + AMP ) .
Root respiration
Three seedlings of each species per treatment were prepared.
At 4 days after HT was begun, the entire root system of each
seedling (HT and control) was excised, and respiration
was measured with an open portable photosynthesis system
(LI-6400, LI-COR, Lincoln, NE, USA), which controlled the flow
rate and CO2 concentration in the inflow and measured the CO2
concentration in the outflow. Figure 1a shows the outline of
the measurement system. Respiration of HT roots was measured under hypoxic conditions, and that of control roots was
measured under normoxic and hypoxic conditions. Each
excised root (150–200 mg dry weight) was immediately
immersed in 150 ml of the nutrient solution in a 180-ml widemouth glass jar, which was kept at 30 °C in a water bath.
Figure 1. ​Outline of the measurement system for analyzing root CO2 production rate (a) and O2 consumption rate (b). IRGA, infrared gas analyzer;
DO, dissolved oxygen.
Tree Physiology Online at http://www.treephys.oxfordjournals.org
232 Kogawara et al.
The mouth of the jar was sealed with a rubber stopper, through
which two gas tubes (for inflow and outflow) were inserted.
The jar was set in the flow line between the pump and the
infrared gas analyzer connected to the photosynthesis system.
The tip of the inflow tube was immersed into the nutrient solution to aerate the solution, while the tip of the outflow tube was
set above the solution surface. The inflow rate and CO2 concentration were controlled by the photosynthesis system to be
300 µmol s−1 and 350 µmol mol−1, respectively. The air was
supplied to the pump from a 50-ml tube to control the O2 concentration in the inflow. For hypoxic measurements, N2 gas
was injected into the tube at 500 ml min−1, resulting in 0.4–
0.5 mg l−1 O2 in the inflow; no N2 injection was used for normoxic measurements. The outflow from the jar passed through
a cylinder filled with silica gel before entering the infrared gas
analyzer. The CO2 concentration in the inflow and outflow was
measured over 20 min, and the CO2 production rate was calculated from the difference and the flow rate. The difference in
CO2 concentration between the inflow and the outflow ranged
from 5 to 10 µmol mol−1.
Immediately after the CO2 production rate was measured,
the jar was detached from the flow line and filled with the nutrient solution to its maximum capacity and O2 consumption rate
was measured. Figure 1b shows the outline of the measurement system. The jar was sealed with a sponge, through which
the dissolved O2 meter was inserted into the solution. The
decrease in O2 concentration in the nutrient solution was monitored over 10 min at 30 °C. The O2 consumption rate was calculated by linear regression at 0.4–0.5 mg l−1 O2. During the
measurement, the solution was stirred with a magnetic stirrer
to equilibrate the CO2 and O2 concentrations. After all measurements, each excised root was immediately microwaved
and then oven-dried at 80 °C for 48 h to determine the dry
weight. Carbon dioxide production and O2 consumption rates
are expressed as units per weight.
Assuming that CO2 production from fermentation coincides
with that from ethanol fermentation, we calculated the SCR of
the respiratory metabolism from the O2 consumption rate and
the CO2 release rate of each seedling as described previously
(Albrecht et al. 2004). In the generation of CO2 by Glc oxidation via glycolysis, we assumed that the tricarboxylic acid
(TCA) cycle generates six molecules, and ethanol fermentation
generates two molecules of CO2 per Glc molecule.
Simultaneously, six molecules of O2 are consumed in the electron transport chain coupled to the TCA cycle. Thus, the fermentative CO2 production rate and SCR in Glc basis were
calculated as
[Fermentative CO2 production] = [CO2 ]total – [O2 ]total (1)
[SCR ] = [Fermentative CO2 production] / 2 + [O2 ]total / 6 (2)
Tree Physiology Volume 34, 2014
where [CO2]total is the CO2 release rate and [O2]total is the O2
consumption rate of total respiration.
Glycolytic and fermentative enzyme assays
Two containers (one for each treatment), each with five seedlings
per species, were used. At 4 days after HT was applied, the lateral roots (HT and control) were cut off, immediately frozen in
liquid N2 and stored at −80 °C until enzyme extraction. Glycolytic
and fermentative enzymes were extracted from the frozen samples with 1.5 ml of extraction buffer containing 50 mM HEPES–
KOH (pH 7.5), 15 mM MgCl2, 100 mM 2-mercaptoethanol,
2.5 mM dithiothreitol (DTT), 1 mM phenylmethylsulfonyl fluoride
(PMSF), 15% (v/v) glycerol, 0.05% Triton X-100, 0.5% EDTAfree Complete protease inhibitor cocktail (Roche Diagnostics,
Mannheim, Germany) and 6% (w/v) polyvinylpolypyrrolidone
(PVPP). The homogenates were centrifuged at 17,500g at 4 °C
for 10 min, and the supernatants were desalted by centrifugation
on Sephadex-G25 columns equilibrated with a buffer containing
50 mM HEPES–KOH (pH 7.5), 15 mM MgCl2, 2.5 mM DTT,
1 mM PMSF and 15% (v/v) glycerol.
The activities of glucokinase (GK; EC 2.7.1.2), fructokinase
(FK; EC 2.7.1.1), ATP:fructose-6-phosphate phosphotransferase
(PFK; EC 2.7.1.11) and pyrophosphate (PPi):fructose-6phosphate phosphotransferase (PFP; EC 2.7.1.90) were
assayed in reaction mixtures (1 ml) containing 50 µl of the
enzyme extracts according to Bouny and Saglio (1996) with
modifications. For determination of GK or FK activity, the final
reaction mixtures contained 50 mM HEPES–KOH (pH 7.5),
3 mM DTT, 2 mM MgCl2, 1 mM ATP, 0.5 mM NADP+, 1.2 units
ml−1 glucose-6-phosphate dehydrogenase, 4.8 units ml−1 phosphoglucose isomerase and 1 mM Glc or fructose. The reactions
were started by the addition of ATP. For determination of PFK
activity, the final reaction mixtures contained 50 mM HEPES–
KOH (pH 7.5), 5 mM fructose-6-phosphate, 2 mM MgCl2, 1 mM
ATP, 0.2 mM NADH, 1.2 units ml−1 aldolase, 1.3 units ml−1 glycerol-3-phosphate dehydrogenase and 8.4 units ml−1 triose
phosphate isomerase. The reactions were started by the addition of ATP. For determination of PFP activity, the final reaction
mixtures contained 50 mM HEPES–KOH (pH 7.5), 5 mM fructose-6-phosphate, 2 mM MgCl2, 1 mM PPi, 0.2 mM NADH,
5 µM fructose-2,6-bisphosphate, 1.2 units ml−1 aldolase,
−1
1.3 units ml
glycerol-3-phosphate dehydrogenase and
8.4 units ml−1 triose phosphate isomerase. The reactions were
started by the addition of PPi. Pyruvate kinase (PK; EC 2.7.1.40)
activity was measured in the reaction mixtures (1 ml) containing 50 µl of the enzyme extracts and 60 mM Mops–NaOH (pH
7.2), 40 mM KCl, 10 mM MgCl2, 1 mM ADP, 0.1 mM NADH and
4 units ml−1 lactate dehydrogenase (Yamanoshita et al. 2005).
The reactions were started by the addition of ADP. Alcohol
dehydrogenase activity was measured in the reaction mixtures
(1 ml) containing 50 µl of the enzyme extracts and 50 mM
MES–NaOH (pH 6.0), 2 mM DTT, 6 mM MgCl2, 0.1 mM NADH
Steady sucrose degradation in hypoxic roots 233
and 380 mM acetaldehyde (Kogawara et al. 2006). The reactions were started by the addition of acetaldehyde. All reactions
were performed at 30 °C.
Enzyme activities were calculated from NADP reduction or
NADH oxidation, monitored as the change in absorbance at
340 nm on a spectrophotometer (MPS-2450, Shimadzu). The
protein concentration in each extract was determined after
desalting by the Bradford (1976) method, and all enzyme
activities were normalized to total protein.
Sucrose cleavage assays
Two containers (one for each treatment), each with four seedlings per species, were prepared. At 4 days after HT was
applied, the lateral roots of each seedling (HT and control)
were processed and stored as described for glycolytic and fermentative enzymes until enzyme extraction.
For the INV assay, the frozen samples were ground with an
ice-cold mortar and pestle in 1.5 ml of the same extraction buffer as for glycolytic and fermentative enzymes. The homogenates were centrifuged at 17,500g at 4 °C for 10 min. The
supernatants were used for the soluble acid INV assay. The
pellets were washed three times with extraction buffer without
PVPP, resuspended in the same buffer containing 500 mM NaCl
and kept at 4 °C overnight to extract the cell-wall INV (Zhang
et al. 2001). The homogenates were centrifuged at 17,500g at
4 °C for 10 min, and the supernatants were used for the cellwall INV assay. Aliquots of each extract were desalted as above.
Soluble acid INV and cell-wall INV were assayed in a total volume of 100 µl; each reaction mixture contained 100 mM
sodium acetate buffer (pH 4.5 and 4.1, respectively), 20 mM
Suc and 10 µl of the enzyme extract in 1.5-ml microtubes. The
reaction was started by the addition of Suc, incubated for
30 min at 30 °C and then stopped by boiling for 1 min. Reducing
sugars were determined by the Somogyi–Nelson method
(Nelson 1944, Somogyi 1952). Mixtures with no incubation
were used as blanks.
Sucrose synthase was extracted according to the method of
Winter et al. (1997) with minor modifications. Frozen samples
were ground with an ice-cold mortar and pestle in 1.5 ml of
extraction buffer containing 50 mM Mops–NaOH (pH 7.5),
10 mM MgCl2, 150 mM NaCl, 1 mM EDTA, 0.1 µM microcystinLR, 100 mM 2-mercaptoethanol, 2.5 mM DTT, 1 mM PMSF,
15% (v/v) glycerol, 0.05% (v/v) Brij35 (Dojindo Laboratories,
Kumamoto, Japan), 0.5% Complete protease inhibitor cocktail
and 6% (w/v) PVPP. The homogenates were centrifuged at
10,000g at 4 °C for 20 min. The supernatants were transferred
to new microtubes and ultracentrifuged at 100,000g at 4 °C for
1 h (Optima TLX, TLA-100.4 rotor, Beckman Coulter, Brea, CA,
USA), and the supernatants were used as soluble SuSy fractions. The pellets were washed with the extraction buffer and
solubilized in the same buffer plus 1% (v/v) 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate. The ­suspensions
were ultracentrifuged at 100,000g at 4 °C for 1 h, and the
supernatants were used as the SuSy microsomal fractions.
Aliquots of each fraction were desalted by centrifugation on
Sephadex-G25 columns equilibrated with a buffer containing
50 mM Mops–KOH (pH 7.5), 5 mM MgCl2, 1 mM DTT, 0.5 mM
CaCl2, 0.1 mM microcystin-LR and 15% (v/v) glycerol. Sucrose
synthase activity was assayed as described by Zeng et al.
(1998) with some modifications. The reaction was performed in
a mixture (42 µl) containing 27 µl of the enzyme extract and
50 mM HEPES–KOH (pH 7.5), 15 mM MgCl2, 10 mM fructose
and 5 mM UDP–Glc at 30 °C for 30 min, and terminated by the
addition of 42 µl of 30% KOH. To remove the unreacted fructose, we heated the mixtures at 100 °C for 10 min and then
cooled them on ice. Sucrose synthase activity was calculated
from the amount of Suc produced, which we quantified by adding 0.14% anthrone in H2SO4 and measuring absorbance at
620 nm. Mixtures with no incubation were used as blanks.
The protein concentration in each extract was determined as
above, and all enzyme activities were normalized to total
protein.
Statistical analyses
Statistically significant differences between treatments in the
mean values of enzyme activities and root respiration rates
were determined by the t-test. The SCR of control or HT roots
under hypoxic conditions was compared with that of control
roots under normoxic conditions by Dunnett’s test. The statistical differences in AEC and ATP concentrations among all combinations of treatments and sugar-feeding types were determined
using Scheffé’s test. All statistical analyses were performed in
the Excel Toukei software (version 6.0, Esumi, Tokyo, Japan).
Results
Energy status of HT roots with or without sugar feeding
After 4 days of HT without sugar feeding, AEC decreased
<0.72 in E. camaldulensis roots, but did not decrease in
M. cajuputi roots in comparison with normoxic roots (Figure 2a
and b). The root ATP content was decreased by HT in E. camaldulensis but not in M. cajuputi (Figure 2c and d). These results
indicate that M. cajuputi can maintain its root energy status
better than E. camaldulensis upon several days of hypoxia.
To investigate whether low substrate availability for glycolysis is the cause of reduced energy in HT roots of E. camaldulensis, we compared the effects of Glc or Suc feeding on the
energy status of HT and control roots of this species. Control
roots fed with Glc or Suc showed AEC similar to unfed control
roots (Figure 2a), suggesting that the sugar feeding did not
have any effect on the root energy status. Hypoxic treatment
roots fed with Glc showed similar AEC to control roots, but HT
roots fed with Suc showed lower AEC than control roots
(Figure 2a). The difference in the ATP content between unfed
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234 Kogawara et al.
Figure 2. ​Energy status (a and b) and ATP concentration (c and d) were assayed for control (white bars) and hypoxic (gray bars) roots of E. camaldulensis (a and c) and M. cajuputi (b and d). Recovery effects of Glc or Suc feeding on root energy status and ATP concentration were investigated.
Data represent mean values plus standard deviation (n = 5). Different letters indicate that the results are significantly different at P < 0.05 by
Scheffé’s test. DW, dry weight.
control and HT roots was abolished by Glc feeding (Figure 2c).
These results indicate that the shortage of substrate for glycolysis causes the lower energy status in HT roots of E. camaldulensis. In contrast, Suc feeding did not recover AEC to the
level of the control roots. In the roots of M. cajuputi, the AEC
and ATP content of HT roots was similarly high regardless of
the sugar feeding (Figure 2b and d).
Respiration rate and sugar consumption rate
In both species, the O2 consumption rate in control roots analyzed under hypoxic conditions was less than half that under
normoxic conditions (Figure 3a), indicating that mitochondrial
respiration was suppressed during the measurement. In both
species, the O2 consumption rate was similar in HT and control
roots analyzed under hypoxic conditions (Figure 3a). Carbon
dioxide production in control and HT roots was similarly
suppressed during analysis under hypoxic conditions in
­
E. camaldulensis (Figure 3b), whereas HT roots of M. cajuputi
showed a slight (but not significant) increase in CO2 production compared with control roots (Figure 3b). The difference
between the CO2 production rate and the O2 consumption rate
can be interpreted as the rate of CO2 production from fermentation (Eq. (1); Figure 3c). In control roots of both species,
Tree Physiology Volume 34, 2014
fermentative CO2 production increased when measured under
hypoxia (Figure 3c). Fermentative CO2 production in the HT
roots was more than double that in control roots in M. cajuputi
measured under hypoxia, but no such difference was observed
in E. camaldulensis (Figure 3c).
Sugar consumption rate (Eq. (2)) in control roots under normoxia was compared with that in control or HT roots under
hypoxia (Figure 4). In control roots of both species, SCR was
lower under hypoxia than under normoxia. The responses of
HT roots differed between the species. The SCR in E. camaldulensis HT roots was lower than that in control roots under normoxic conditions whereas that in M. cajuputi HT roots was
similar to that in control roots under normoxic conditions.
Activities of glycolytic enzymes
In the roots of E. camaldulensis, the activities of all glycolytic
enzymes investigated were not affected by HT (Table 1). In the
roots of M. cajuputi, the activities of PFK and PK were increased
upon HT, whereas those of the other glycolytic enzymes were
not affected (Table 1). Alcohol dehydrogenase activity in HT
roots was 4.6 times that in the control roots in E. camaldulensis
and 3.1 times in M. cajuputi (Table 1), in line with our previous
report (Kogawara et al. 2006).
Steady sucrose degradation in hypoxic roots 235
Figure 4. ​Effects of root hypoxia on SCR in roots. Sugar consumption
rate was expressed as hexose consumption rate, which was calculated
from the CO2 production and O2 consumption rates (see main text for
calculations). The roots of control seedlings were analyzed under normoxic or hypoxic conditions. The roots of seedlings subjected to HT
were analyzed only under low-O2 conditions. Data are means ± standard deviation (n = 3). †Significant difference at P < 0.05 (Dunnett’s
test) compared with control roots measured under normoxic conditions. n.d., not determined; DW, dry weight.
Table 1. ​Effects of root hypoxia (HT) on glycolytic and fermentative
enzymes in the roots of E. camaldulensis and M. cajuputi seedlings.
Enzymes Activity (nmol min−1 mg−1 protein)
Figure 3. ​Effects of root hypoxia on (a) total O2 consumption rate, (b)
total CO2 production rate and (c) fermentative CO2 production rate of
roots excised after 4 days’ HT. Fermentative CO2 production rate was
calculated by subtracting total O2 consumption rate from total CO2 production rate. The control roots (grown under normoxia) were analyzed
under normoxic or low-O2 conditions. The roots of seedlings subjected to
HT were analyzed only under low-O2 conditions. Data are means ± standard deviation (n = 3). *Significant difference between the control and HT
roots at P < 0.05 (t-test). n.d., not determined; DW, dry weight.
GK
FK
PFK
PFP
PK
ADH
E. camaldulensis
M. cajuputi
Control roots HT roots
Control roots HT roots
52 ± 11
113 ± 26
25 ± 4
94 ± 18
60 ± 17
807 ± 330
40 ± 6
30 ± 7
97 ± 12
89 ± 10
35 ± 12
83 ± 20
101 ± 41
172 ± 35
94 ± 63
67 ± 10
3745 ± 535 591 ± 71
34 ± 5
90 ± 15
140 ± 31
243 ± 85
116 ± 39
1860 ± 443
Mean values (n = 4) ± standard deviation are shown. Bold type indicates significant difference at P < 0.05 between control and HT
(t-test).
Activities of sucrolytic enzymes
In both species, the activity of cytosolic SuSy was not altered
by HT (Figure 5a), whereas the activity of membrane-­
associated SuSy was significantly increased in E. camaldulensis,
but not in M. cajuputi (Figure 5b). The soluble acid INV activity
was inhibited by HT in the roots of E. camaldulensis, but not in
those of M. cajuputi (Figure 6a). The cell-wall INV activity was
unaffected by HT in both species (Figure 6b).
Discussion
In our previous study using the same experimental system and
species, we monitored photosynthesis and growth response
during 19 days of HT (Kogawara et al. 2006). By the fifth day
of HT, the photosynthesis of E. camaldulensis was reduced by
stomatal closure. On the 10th day of HT and thereafter, the
photosynthesis and growth of E. camaldulensis were inhibited
almost completely, while no effect of HT was observed in
M. cajuputi throughout the duration of the treatment (Kogawara
et al. 2006). Both species showed induction of fermentation
activity, evaluated by ADH activity, by the first day of HT. And
at the third day of HT, both species showed different effects of
HT on photoassimilate allocation (Kogawara et al. 2006). In
the present study, we investigated the metabolic process that
is responsible for the interspecies difference in tolerance to
root hypoxia. We focused on the fourth day of HT, when metabolism is already affected but photosynthesis is still operating in
E. camaldulensis.
The ability to maintain energy status and ATP content under
root hypoxia was higher in M. cajuputi than in E. camaldulensis
(Figure 2). Low energy status under O2 deficiency can be
Tree Physiology Online at http://www.treephys.oxfordjournals.org
236 Kogawara et al.
Figure 5. ​Activities of (a) cytosolic SuSy and (b) membrane-­associated
SuSy in roots of E. camaldulensis and M. cajuputi. Data are
means ± standard deviation (n = 4). *Significant difference between
treatments at P < 0.05 (t-test). White bars, control; gray bars, HT.
Figure 6. ​ ​Activities of (a) soluble acid INV and (b) cell-wall INV in
roots of E. camaldulensis and M. cajuputi. Data are means ± standard
deviation (n = 4). *Significant difference between treatments at
P < 0.05 (t-test).
responsible for a decrease in ion transport (Colmer et al.
2001) or root hydraulic conductivity (Tournaire-Roux et al.
2004), which is in line with the growth reduction of E. camaldulensis but not of M. cajuputi under root hypoxia (Kogawara
et al. 2006). In the previous study investigating the effects of
flooding on M. cajuputi roots, root AEC showed a drop to 0.7
on the second day of flooding and then recovered to the level
of the control on the fourth day of the treatment (Yamanoshita
et al. 2005), which is in agreement with the result in the present study. Both studies indicate that M. cajuputi is the species
able to maintain normal energy status of roots when hypoxic
stress is imposed.
The results showing that AEC recovered in HT roots of E.
camaldulensis by feeding Glu revealed that substrate availability is the limiting factor of energy production in HT roots of this
species. This is consistent with previous studies reporting that
Glu feeding to hypoxic roots had a positive effect on survival or
growth of root tips (Webb and Armstrong 1983, Bouny and
Saglio 1996), suggesting that carbohydrates are the primary
energy resource whose supply is more important for hypoxic
roots than for well-aerated roots. The importance of carbohydrate reserves has also been proposed in studies on tree
­species, in which relatively flood-intolerant species reduced
soluble sugar concentration (Kolb et al. 2002, Ferner et al.
2012) or starch concentration (Angelov et al. 1996, Gravatt
and Kirby 1998) in flooded roots, but relatively flood-tolerant
species kept them high. In contrast to Glu feeding, Suc feeding
to HT roots of E. camaldulensis did not recover AEC (Figure 2a
and c), implying that limited supply of Glc to glycolysis may be
caused by impaired Suc degradation. Sucrose accumulation in
HT roots in the species observed in the previous study
(Kogawara et al. 2006) is in line with this possibility.
To investigate whether the activation of fermentation in HT
roots differs between the two species, we measured root CO2
production and O2 consumption under hypoxia. Fermentative
CO2 production rate was similar in control and HT roots of
E. camaldulensis, whereas activation of fermentation was
observed in HT roots of M. cajuputi (Figure 3c). Tolerance to
O2 deficiency can be improved with increasing ethanol production, which represents the increased fermentation (Bouny and
Saglio 1996, Huang et al. 2003, Gharbi et al. 2007). The difference in the activation of fermentation in HT roots is the likely
cause of the difference in tolerance to root hypoxia between
E. camaldulensis and M. cajuputi.
These interspecies differences in fermentation can be interpreted in terms of sugar consumption. In roots grown under
normoxic conditions, a sudden imposition of O2 deficiency on
them leads to a net decrease of respiratory sugar consumption
because the aerobic respiration is restricted and fermentation is
yet to be induced. This was shown in the SCR of the control
Tree Physiology Volume 34, 2014
Steady sucrose degradation in hypoxic roots 237
roots measured under hypoxic conditions, in both M. cajuputi
and E. camaldulensis (Figure 4). Along with the de novo synthesis of fermentative enzymes, which is a part of the plant adaptive response to low O2 (Chang et al. 2000), SCR is expected to
increase with the growing demand for fermentation substrate.
This is the case for HT roots of M. cajuputi, in which SCR was
restored to the normoxic level (Figure 4) with the increased
fermentation (Figure 3c). In contrast, E. camaldulensis failed to
restore SCR in HT roots (Figure 4), which reflects the lack of an
increase in fermentation (Figure 3c). Glycolytic flux in HT roots
of E. camaldulensis seems not high enough to produce ATP
through fermentation. Theoretically, low glycolytic flux in HT
roots of E. camaldulensis could have resulted from an enzymatic
impairment or substrate shortage. The recovery of energy status and ATP content in HT roots of this species by Glc feeding
(Figure 2a and c) revealed that substrate shortage but not
enzymatic impairment is the key to the low glycolytic flux in HT
roots of this species. No inhibition of the activities of glycolytic
enzymes in HT roots is in line with this conclusion (Table 1).
Since we measured the enzymatic activities in vitro, some regulation in planta may affect the response manner of the enzyme
activities to HT. However, this should not have limited the glycolytic flux in HT roots of E. camaldulensis, which was evidenced
by the results of the Glc feeding.
The activities of several glycolytic enzymes increased in
M. cajuputi roots (Table 1), similar to observations in other plant
species (Rivoal et al. 1989, Fox et al. 1995, Bouny and Saglio
1996). An increased turnover of carbohydrate incorporated
into the soluble fraction in HT roots of this species, suggested
by 13C experiments in the previous study (Kogawara et al.
2006), is consistent with this result. Nevertheless, the SCR
value of HT roots under hypoxia was not superior to that of the
control roots under normoxic conditions (Figure 3). These
results imply that the glycolytic flux into pathways other than
ethanol fermentation increased in M. cajuputi roots under
hypoxia. In many plant species, the glycolytic flux into lactic
acid fermentation is well known to increase in early response
to O2 deficiency (Davies et al. 1974, Roberts et al. 1984, Rivoal
and Hanson 1994). Lactic acid fermentation plays a role in
regenerating NAD+ to supply for glycolysis under O2 deficiency,
as well as ethanol fermentation (Greenway and Gibbs 2003).
However, there are only a few cases in which lactate dehydrogenase activity persists for a long period after the induction of
ethanol fermentation (Hoffman et al. 1986, Good and Crosby
1989). As for M. cajuputi roots, lactate dehydrogenase activity
was not induced during the first week, while it was slightly
induced during the second week (Yamanoshita et al. 2005).
Thus, lactic acid fermentation is unlikely to be the reason for
the higher tolerance of M. cajuputi. Alanine, a product of
­pyruvate metabolism, is also a major metabolite accumulated
under O2 deficiency (Ricoult et al. 2005, Kato-Noguch 2006,
Kreuzwieser et al. 2009). While it has been suggested that
alanine production during anoxia may contribute to mitigate
cytoplasmic acidification by competing with lactate production
for pyruvate (Ricoult et al. 2005), evidence that alanine production directly affects energy status during O2 deficiency has
not been reported so far.
Since the glycolysis–fermentation pathway produces less
ATP than aerobic respiration, a decrease in ATP production
through inhibition of the TCA cycle cannot be compensated for
by this pathway unless sugar consumption is increased (Gibbs
and Greenway 2003). Despite no increase in SCR, HT roots of
M. cajuputi sustained an energy status comparable to that of
the control roots (Figure 2b). It is possible that the energy
requirement for maintenance is reduced in HT roots of M. cajuputi, similar to anoxia-tolerant plant tissues such as rice coleoptiles (Colmer et al. 2001) and storage roots of red beet (Zhang
and Greenway 1994). Generally in plant species, O2 deficiency
reduces protein synthesis, biosynthetic processes that have a
high energy cost (Edwards et al. 2012), but often simultaneously increases the net synthesis of specific proteins (Sachs
et al. 1980, Mocquot et al. 1981, Ricard and Pradet 1989),
so-called anaerobic proteins (ANPs). Many of the ANPs are
enzymes of glycolysis and fermentation, presumably related to
induced activities of these enzymes as observed in M. cajuputi
(Table 1). In addition to these classical ANPs, some other proteins whose synthesis is enhanced under anoxia were detected
in organs that can grow under anoxia (Ishizawa et al. 1999,
Huang et al. 2005), including enzymes involved in the PPidependent metabolism and scavenging system of reactive oxygen species (Huang et al. 2005). Such specific protein
synthesis allows plant cells to use energy efficiently for maintenance, resulting in a reduction of energy requirement
(Greenway and Gibbs 2003). This could also be the case for
HT roots of M. cajuputi.
Previous reports have indicated that the sustained activity
of carbohydrate-degrading enzymes to ensure hexose entry
into respiratory metabolism is a key for energy production
under O2 deficiency. A difference in tolerance of two oak species to waterlogging could be attributed to their different abilities to maintain glycolysis and fermentation, reflected in the
upregulation of mRNA levels of not only glycolytic and fermentative enzymes but also amylase, which degrades starch to
soluble carbohydrates (Le Provost et al. 2012). Cereal seeds
with dysfunctional amylases cannot germinate under anoxia,
and benefit in this respect from exogenous Glc (Perata et al.
1992), which suggests that the inability to use carbohydrate
reserves for glycolysis under anoxia is critical for energy production for seed germination. Amylase activity has been suggested to play a key role in the maintenance of glycolytic flux
into ethanolic fermentation in elongating rice coleoptiles under
anoxia (Huang et al. 2003), and variation in amylase activity
was correlated with variation in anoxia tolerance among rice
cultivars (Ismail et al. 2009). As well as the role of starch
Tree Physiology Online at http://www.treephys.oxfordjournals.org
238 Kogawara et al.
­ egradation during O2 deficiency, Suc degradation must be
d
sustained to supply substrate for anaerobic energy metabolism, especially when carbohydrate supply depends mainly on
photoassimilate transport from the shoot. This situation was
demonstrated by our present study using intact roots with
sugar feeding (Figure 2), and was the reason for analyzing
activities of Suc-degrading enzymes, SuSy and INV.
To investigate the background of the impaired degradation
of Suc in HT roots of E. camaldulensis, we measured the activities of soluble and insoluble INV and SuSy. Since Suc degradation via SuSy saves ATP in comparison to its degradation via
INV when its products are subsequently metabolized in glycolysis (Stitt 1998, Koch 2004), the induction of SuSy (in particular, its cytosolic form) has been recognized as an adaptive
response to low-O2 conditions (Guglielminetti et al. 1997,
Zeng et al. 1999). However, HT did not affect the activity of
cytosolic SuSy in either species (Figure 5a), while increasing
the activity of membrane-associated SuSy in E. camaldulensis
(Figure 5b). Membrane-associated SuSy plays a role mainly in
directing carbon to polysaccharide synthesis (Amor et al.
1995). The activity of membrane-associated SuSy was
increased and was accompanied by callose deposition in
anoxic maize root tips (Subbaiah and Sachs 2001), or by cellulose deposition in hypoxic wheat roots (Albrecht and
Mustroph 2003). Thus, the induction of membrane-associated
SuSy under hypoxia could reduce substrate availability for glycolysis. However, at least in E. camaldulensis, this effect would
be negligible, since the previous study showed that the incorporation of 13C-labeled photoassimilate into the structural carbohydrate, the corresponding fraction for cellulose and
callose, was not increased during HT (Kogawara et al. 2006).
Thus, SuSy seems not involved in the impaired degradation of
Suc in HT roots of E. camaldulensis.
Soluble acid INV activity in E. camaldulensis was reduced by
HT (Figure 6a), which is in agreement with observations for
other plant species (Zeng et al. 1999, Albrecht and Mustroph
2003). Soluble acid INV is located in the vacuole in most plant
species and plays a pivotal role in hexose entry into cytoplasmic metabolism (Winter and Huber 2000, Koch 2004, Bocock
et al. 2008). Inhibition of soluble acid INV in HT roots of
E. camaldulensis (Figure 6a) may be responsible for the limited
substrate availability for respiratory metabolism in HT roots. The
accumulation of root Suc observed in our previous study
(Kogawara et al. 2006) may result from inhibition of this enzyme.
Sustained activity of soluble acid INV under HT in M. cajuputi
(Figure 6) would provide an advantage to this species by keeping the substrate availability for respiratory metabolism at a sufficient level. While the behavior of SuSy has been extensively
studied as a factor involved in the mechanism of low O2 tolerance (e.g., Guglielminetti et al. 1997, Ricard et al. 1998, Zeng
et al. 1998, 1999, Kumutha et al. 2008), INV has not been
previously reported as a determinant of low O2 tolerance.
Tree Physiology Volume 34, 2014
Because SuSy has a role in saving ATP as mentioned above,
and INV participates in futile cycling of sugars that consume
considerable ATP (Dieuaide-Noubhani et al. 1995, Nägele et al.
2010), INV repression accompanied by cytosolic SuSy induction has an advantage of avoiding energy waste under low O2
conditions. However, our results suggest that, in the case when
cytosolic SuSy is not induced, stability of INV is crucial for maintenance of energy metabolism under low O2 conditions. The
adaptive response of SuSy for anaerobic energy metabolism is
not universal among plant species, as another role of SuSy in
carbon partitioning under low O2 condition has been found
(Subbaiah and Sachs 2001, Albrecht and Mustroph 2003).
Therefore, a comprehensive analysis of SuSy and INV is important for understanding the underlying causes of interspecies
difference in hypoxia tolerance.
In conclusion, our data are consistent with the hypothesis
that the difference in stable sucrolytic activity (mainly due to
soluble acid INV) and hence in substrate availability for glycolysis determines the difference in energy metabolism and in tolerance to root hypoxia between M. cajuputi and E. camaldulensis.
In E. camaldulensis, the decline in substrate availability is likely
to be the main cause of the failure to activate fermentation in
response to hypoxia. Sustained sucrolytic enzyme activities in
HT roots of M. cajuputi should be a prerequisite for maintaining
high energy status under hypoxia.
Acknowledgment
We thank Mr Tomoaki Watanabe (University of Tokyo) for
greenhouse maintenance.
Conflict of interest
None declared.
Funding
This work was supported in part by a Grant-in-Aid for Scientific
Research (Grant No. 18380087) from the Ministry of
Education, Culture, Sports, Science and Technology of Japan
and by Global Environment Research Fund from the Ministry of
the Environment of Japan.
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