Plant, Cell and Environment (2000) 23, 1109–1118
Stomatal responsiveness to leaf water status in common
bean (Phaseolus vulgaris L.) is a function of time of day
M. MENCUCCINI,1 S. MAMBELLI2 & J. COMSTOCK1
1
Boyce Thompson Institute for Plant Research at Cornell University, Tower Road, Ithaca, 14853, NY USA and 2Department
of Integrative Biology, University of California Berkeley, 94720, CA, USA
ABSTRACT
Stomatal response to leaf water status was experimentally
manipulated by pressurizing the soil and roots of potted
common bean plants enclosed in a custom-built root pressure chamber. Gas exchange was monitored using a wholeplant cuvette and plant water status using in situ leaf
psychrometry. Bean plants re-opened their stomata upon
pressurization, but the extent of re-opening was strongly
dependent on the time of day when the soil was pressurized, with maximum re-opening in the morning hours and
limited re-opening in the afternoon. Neither leaf nor xylem
abscisic acid concentrations could explain the reduced
response to pressurization in the afternoon. The significance of this phenomenon is discussed in the context of
circadian rhythms and of other recent findings on the
‘apparent feed-forward response’ of the stomata of some
species to vapour pressure deficit.
Key-words: abscisic acid; common bean; endogenous
rhythms; hydraulic signalling; leaf water stress; stomatal
conductance.
Abbreviations: A, net assimilation rate (mmol m-2 s-1); gs,
stomatal conductance (mol m-2 s-1); Dw, Dws, leaf-to-air and
leaf-to-leaf surface vapour pressure difference (mmol
mol-1); ca, ci, atmospheric and intercellular CO2 concentration (mmol mol-1); E, transpiration rate per unit leaf area
(mmol m-2 s-1); Ys, Yl, Ygc, Yepi, soil, bulk leaf, guard cell
and epidermal water potential (MPa); Pgc, Pepi, guard cell
and epidermal turgor pressure (MPa).
INTRODUCTION
Stomatal responses to air humidity have received considerable attention in the last few decades (e.g. Cowan 1977;
Losch & Tenhunen 1981; Grantz 1990; Schulze 1993;
Monteith 1995). Despite these efforts, considerable uncertainties still exist on many important aspects. Although our
knowledge is rapidly progressing in identifying the molecCorrespondence (present address): Maurizio Mencuccini, Institute
of Ecology and Resource Management, University of Edinburgh,
The Kings Buildings, Mayfield Road, EH9 3JU, Edinburgh, Scotland UK. Fax: +44 1316620478; E-mail: [email protected]
© 2000 Blackwell Science Ltd
ular and bio-chemical mechanisms behind stomatal movements, a consensus on the ecophysiological determinants of
stomatal opening has not been reached, especially in relation to responses to air and soil humidity (Cowan 1995).
More recently, the role of chemical signals transported by
the transpiration stream has been explored in some detail
and hypotheses have been put forward to explain stomatal
responses to water stresses on their basis (Davies & Zhang
1991). While the discussion on the respective roles of leaf
versus root metabolite production has dominated the literature of the past decade, relatively little attention has been
paid to the role of leaf water status, once thought to represent one of the most important variables in stomatal
control. Recently, a number of papers have challenged the
view that leaf water status plays no role in stomatal regulation by showing that stomata can be made to re-open
under various circumstances after direct manipulation of
leaf water status (Saliendra, Sperry & Comstock 1995;
Fuchs & Livingston 1996; Comstock & Mencuccini 1998).
Leaf water status was manipulated by pressurizing the soil
and plant roots enclosed in large root pressure chambers.
During experiments designed to test whether changes in
leaf water status could explain stomatal closure due to root
chilling, soil drought and leaf-to-air vapour pressure difference, we observed that the extent of re-opening under constant environmental conditions varied, depending on when
in the day the pressurization was performed. This observation led to a further set of experiments specifically designed
to test the hypothesis that stomatal responsiveness to manipulations of leaf water status was a function of time of day.
MATERIALS AND METHODS
Experimental material
Two common bean (Phaseolus vulgaris L.) cultivars of the
‘pinto bean’ type were selected for analysis, based on previous work by Comstock & Ehleringer (1993). The two cultivars, G4523 and G5201, have both been selected for use
under rain-fed conditions, but have different centres of
diversity for their germoplasm (i.e. the geographical location where their wild ancestor had probably originated, cf.
Singh, Gepts & Debouck 1991). Accordingly, they differ in
their gas exchange and hydraulic properties (Mencuccini &
Comstock 1999).
1109
1110 M. Mencuccini et al.
Greenhouse propagation
The plants used for this experiment were grown between
October 1996 and August 1997 in the greenhouses of Boyce
Thompson Institute for Plant Research (Ithaca, NY, USA).
After the seeds had germinated, the seedlings were transplanted into 5 L pots filled with an amended mix of fritted
clay, silica sand, pasteurized soil, vermiculite and peat.
Plants were kept well watered and fertilized throughout the
experiment. More details about plant propagation can be
found in Mencuccini & Comstock (1999).
Plants received supplementary lighting using a combination of Na-vapour and metal halide lamps. Photoperiod was
12 h starting at 0600 h. Day/night time conditions were
approximately 30/20 °C, 40/80% relative humidity, and
375/390 mmol mol-1 CO2. A set of several rotating fans continuously stirred the air during growth.
Gas exchange and root pressurization
Measurements started when plants were 4-week-old (from
emergence). Plants were first loaded inside a large custombuilt root pressure chamber. The whole canopy was
enclosed inside a 0·6 m ¥ 0·5 m ¥ 0·5 m water-jacketed
cuvette lined with Teflon film. The speed of leaf temperature control was increased by locating two radiators in the
air flow pathways of the cuvette. Internal mixing fans generated air movement 10–100 times the rate of air flow
through the cuvette for gas exchange measurements, giving
an average wind speed of ~ 0·5 m s-1 in the cuvette. The
design of the whole gas-exchange plus root pressurization
system was as described in Comstock & Mencuccini (1998),
except for a hinged frontal access panel, which was added
to facilitate sampling of plant material during experiments.
The access panel was equipped with Teflon cuffs sealed
internally to the cuvette walls. When the panel was opened
for sampling leaf material, the cuffs could be quickly closed
around the arms of the operator to avoid contamination of
cuvette air with external air.
Adaxial and abaxial leaf boundary layer conductances
(mol m-2 s-1), were determined at varying heights inside the
cuvette, using leaf replicae made out of filter paper sealed
on one side with sticky Teflon film (values between 0·9 and
3·7 mol m-2 s-1). Stomatal ratio was assumed 0·3 for both
cultivars based on previous work (Comstock & Ehleringer
1993). Leaf temperature was calculated as the average of
five type-E thermocouples inserted into leaves in different
parts of the canopy. Gas exchange calculations followed
von Caemmerer & Farquhar (1981) and Comstock &
Ehleringer (1993). Gas exchange and environmental parameters in the cuvette were automatically logged every
two minutes to provide complete time courses of the experiments. Values could also be manually logged when steadystate conditions were reached after pressurization/
depressurization.
The cuvette buffering volume was evaluated following
step changes in inflow environmental conditions; 99% of
the cuvette response was typically achieved in less than four
minutes.
Abscisic acid collection and analyses
For each plant, three leaflets were cut at the beginning of
the day to expose petioles for sap collection. Each petiole
was then fitted with a rubber gasket. After steady-state gas
exchange was obtained under ambient conditions, roots
were pressurized 0·2–0·3 MPa above the final equilibrium
pressure. The excess pressure caused emission of sap from
the cut petioles. Once the bleeding had continued for about
5 min, a vacuum line was used to thoroughly dry off the
petiole before a vial was fitted around it. The previously
positioned rubber gaskets were used to create a seal around
the petiole preventing evaporation during sap collection.
Sap collection continued for about 5 min until each vial was
almost full. Since the entire root system was under pressure,
this procedure maximized the chances that only true xylem
sap was sampled. After sampling, vials were immediately
frozen in a - 60 °C freezer pending analysis.
Whole leaf abscisic acid concentration (ng g-1 fresh
weight) was measured on four samples taken from four different leaves per plant. A circular area of about 0·8 cm2 was
cut out of each leaf using a hole puncher, immediately
sealed in a vial containing 200 mL of extraction solvent
(methanol 20%, acetic acid 1%, v/v) and frozen in a - 60 °C
freezer. Attention was paid to avoid sampling leaf patches
containing large veins. The leaf samples were homogenized
and extracted in darkness at 4 °C for 24 h, then centrifuged
at 5000 g for 5 min.The pellet was re-extracted with another
200 mL of extraction medium and the supernatants were
pooled.
C18 chromatography was used to purify abscisic acid
(ABA), eliminating lipid-like components and soluble
sugars by partitioning of the crude leaf extracts and xylem
sap samples. The ABA-containing fraction was eluted in
120 mL of solvent (methanol 55%, acetic acid 1%, v/v),
vacuum dried and re-dissolved in 120 mL of Hepes buffered
saline solution (50 mM Hepes, 1 mM MgCl2, 10 mM
NaCl, 0·02% (w/v) NaN3, pH 7·5). Aliquots from this solution were assayed for ABA by indirect enzyme-linked
immunosorbent assay (ELISA) (Walker-Simmons 1987;
Ober et al. 1991) using commercially available monoclonal
antibodies (Idetek, San Bruno, CA). (+)ABA concentration
was determined by calculations based on (+)ABA standard
of 0·01–2·5 pmol well-1. Leaf extracts and xylem sap
samples showed no significant interference in a dilutionspike test (Jones 1987).
Xylem ABA fluxes (nM m-2 s-1) were calculated multiplying transpiration rate E and measured xylem [ABA]
(nM m-3).
Xylem water potentials
Xylem water potentials were measured using in situ temperature-compensated leaf psychrometers (leaf hygrome-
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
Stomatal responsiveness to leaf water status 1111
ters, Plant Water Status Instrument Inc., Guelph, Ontario,
Canada) (Dixon & Tyree 1984). Two to three psychrometers were positioned on leaves located in different parts of
the canopy. To improve speed of water vapour equilibration
between internal leaf spaces and psychrometer chamber, a
patch of leaf surface was gently brushed for about 1 min
with a very fine paint brush imbibed with Cellite“ dissolved
in double-distilled water. The leaf surface was then thoroughly rinsed with distilled water and dried out. A 2 cm
square of sticky Teflon with a circular hole in the centre was
then applied onto the leaf surface. The Teflon had the function of avoiding direct contact between the leaf surface and
the compound used to create a vapour-tight seal around the
psychrometric chamber, while allowing free vapour movement in and out of the psychrometer chamber through the
central hole. Dental compound was used to create a vapourtight seal (Reprosil; Dentsply Intl. Inc, Milford, DE, USA).
Once positioned, the psychrometer holder was covered
with four to five layers of aluminium foil to minimize radiative exchanges. Temperature gradients were normally
below 1 mV and frequently below 0·5 mV. Preliminary
experiments were conducted to assess both the accuracy of
water potential measurement by the leaf psychrometers
and the speed of vapour equilibration after step changes
in environmental conditions (normally between 10 and
15 min).
The area covered by the psychrometer holder was a few
times larger than the area of the chamber itself, but still a
rather small fraction of the total leaf blade area. Consequently, transpiration from the patch was effectively suppressed (because of the vapour seal created around the
chamber) and the leaf psychrometer was likely reading a
value of water potential in close equilibrium with the veins
of the fully transpiring leaf.
Experimental protocol
On the morning of the experiment, the plant was exposed to high irradiance levels (about 1·6 mmol m-2 s-1
photosynthetic active radiation at chamber mid-height),
favourable leaf temperatures (25 °C), ambient CO2 of
360 mmol mol-1, and stable leaf-to-air humidity gradients,
Dw (mmol mol-1).These conditions were then kept constant
throughout the day.The level of Dw was intentionally varied
among plants (i.e. among different experiments) between
about 9 and 22 mmol mol-1, to test the effect of this parameter on the diurnal response to pressurization. After
reaching the equilibrium point, the soil was pressurized to
promote stomatal re-opening while the environmental conditions were kept constant. Pressurization and depressurization was carried out at a rate of less than 0·1 MPa min-1.
The pressure used varied between 0·3 and 0·6 MPa, and
was chosen to maximize re-opening in accordance with the
level of vein water potential measured by the in situ
psychrometers.
The sequence of pressurization–depressurization was
then repeated throughout the day. Normally, between two
and four sequences could be completed each day. Altogether, 14 experiments were carried out using 10 different
plants. For a subset of 6 d (four different plants), xylem sap
was collected throughout the day and used to measure the
concentration of abscisic acid (from now on [ABA]). Leaf
[ABA] was also measured.
On three plants, to check for diurnal trends in the
absence of any perturbation, control experiments were performed by keeping conditions constant all day long (i.e.
photosynthetically active radiation, leaf temperature (T),
and leaf-to-leaf surface vapour pressure difference, Dws).
The intercellular CO2 concentration (ci) was kept constant
but the ambient CO2 concentration (ca) was not .
RESULTS
Experimental control over environmental
variables and observed variations in plant gas
exchange parameters
As already described in the previous section, leaf temperature was generally held at 25 °C, ambient CO2 concentration at 360 mmol mol-1, while Dw was intentionally varied
from 9 to 22 mmol mol-1. For each experimental course, on
average, steady-state T was only about - 0·11 °C [25 ± 0·68,
95% confidence intervals (CI)] different from the target
temperature with no systematic difference between unpressurized and pressurized points (t-test for independent samples, t = 1·03, P = 0·31). Corresponding values for
average differences from target ca and Dw were: - 1·77
(360 ± 6·81, CI) mmol mol-1 and - 0·13 (± 1·96,
CI) mmol mol-1, respectively.
Values of stomatal conductance varied largely depending
on the level of the chosen Dw, and to a smaller degree from
plant to plant and from day to day. It ranged from
0·08 mol m-2 s-1 for an unpressurized, early morning point
at Dw = 21·5 mmol mol-1, to 0·44 mol m-2 s-1 for a pressurized, mid-morning point at Dw = 15·2 mmol mol-1. Analogous variability was observed for assimilation rates, with
a range from 6·27 mmol m-2 s-1 (unpressurized, very lateafternoon point) to 17·29 mmol m-2 s-1 (pressurized, midmorning point).
Diurnal responses to pressurization
A typical example of a diurnal course of pressurization
is given in Fig. 1. When the stomatal conductance (gs) had
stabilized at around 0·10 mol m-2 s-1 (Fig. 1b), the soil was
pressurized with 0·6 MPa (Fig. 1a, broken line). The first
closing response by the stomata culminated after about
10 min. A re-opening followed in the next 30 min. Because
of the increased gs and transpiration rate, relative humidity
in the incoming air had to be reduced and [CO2] increased,
which caused the limited closure evident in the next 30 min.
After depressurization, the reverse sequence was observed,
with a first immediate re-opening, soon followed by a
continuous decline to a new steady-state value. This final
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
1112 M. Mencuccini et al.
Time of day (h)
Time of day (h)
Figure 1. Time course of one experiment designed to test the
hypothesis that stomatal responsiveness to root pressurization is
a function of time of day. (a), Roots and soil in a large root
pressure chamber were subjected to three cycles of
pressurization during one day (24 January 1997). The broken line
represents the actual levels of pressurization imposed over time,
while the black points (± SE) give the average value of leaf vein
water potential (MPa) measured with three in situ temperaturecorrected leaf psychrometers. (b), Time course of stomatal
opening and closing upon root pressurization. Upper arrows
indicate times of pressurization, lower arrows those of depressurization. The line is interrupted when system checks were
performed Conditions were: leaf T = 25 °C, Dw = 14 mmol mol-1,
ci = 200 mmol mol-1, soil T = 20 °C.
unpressurized value was slightly greater than the one
obtained at the beginning of the experiment. The second
sequence of pressurization took longer to complete
because, after the initial re-opening, two oscillations followed before a steady-state value was obtained. The final
re-opening was similar to the one in the previous cycle.
However, in the third cycle of pressurization, a lower final
re-opening was apparent, with stomatal conductance now
oscillating a few times around an only slightly elevated
value. For ease of comparison, the three curves of stomatal
response to pressurization during the morning, midday and
afternoon hours are re-plotted in Fig. 2, with values now relative to the equilibrium gs reached before the start of each
pressurization sequence (set at 100). It is apparent that the
afternoon pressurization sequence is quite different from
the previous two (Fig. 2). Similar differences were observed
in the other experiments.
Oscillations were never observed during the morning
hours (i.e. pressurization started before 1030 h), but they
were normal events during midday (i.e. pressurization
started up to 1400 h) and afternoon sequences (i.e. pressurization started only after 1430 h).The oscillation pseudoperiod was 44 min both at midday and in the afternoon.
Although the amplitude of the first oscillation did not differ
between morning and afternoon sequences, the amplitude
of the second was greater in the afternoon (Table 1). The
number of oscillations was also greater in the afternoon
than at midday (Table 1), thereby greatly increasing the
time required to obtain equilibrium.
Xylem water potential (Fig. 1a) also followed a typical
pattern. After having declined to a stable value at around
- 0·58 MPa, it responded to root pressurization by rising to
about - 0·16 MPa. Soon after, a progressive decline was
apparent following stomatal re-opening, until a stable value
at around - 0·37 MPa was obtained, soon before depressurization. The same pattern was followed during the
second cycle of pressurization, with only a subtle tendency
to show an oscillatory behaviour. During the third cycle
however, xylem water potential rose to about - 0·16 MPa
upon pressurization, displayed one clear oscillation and
then stabilized at a much higher value than in the two preceding cycles.
A summary of the steady-state gs and net assimilation
rate (A) data for both cultivars collected in the 14 diurnal
sequences is presented in Figs 3 and 4. In the unpressurized
condition, gs (relative to the maximum value of each plant
in each day) showed evidence of a diurnal course, with
slightly lower values in the early morning, stable values
throughout most of the day and an accentuated decline in
the late afternoon, especially after 1630 h (Fig. 3a). Assimilation rate showed little evidence of lower values during
Time since beginning of pressurization (h)
Figure 2. Blow-up of the time course of Fig. 1(b) for the portions
involving stomatal responses to root pressurization. The three
cycles of pressurization–depressurization are plotted relative to
their own initial stable value of stomatal conductance set at 100,
to increase legibility of the kinetics of re-opening. , morning; ,
midday; —, afternoon.
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
Stomatal responsiveness to leaf water status 1113
Property
Midday
(SE)
Afternoon
(SE)
Significance
Posc, (h)
100 Dgs1/gs
100 Dgs2/gs
n
0 : 44
7·95
6·90
1·46
(0 : 05)
(3·07)
(2·08)
(0·13)
0 : 44
42·19
16·98
2·50
(0 : 07)
(13·85)
(1·47)
(0·33)
NS
NS
**
**
Table 1. Main characteristics of the
oscillations observed during midday and
afternoon pressurization cycles. Morning
pressurization cycles are not shown
because oscillations were never observed.
Standard errors of the mean in parentheses
Midday includes cycles started in the period between 1030 to 1400 h. Afternoon includes
cycles started from 1430 onwards. Posc, pseudo-period of the oscillation in hours; 100 Dgs1/gs,
amplitude of the first oscillation relative to gs after pressurizing; 100 Dgs2/gs, amplitude of
the second oscillation relative to gs after pressurizing; n, damping factor, i.e. number of oscillations before final stable gs. Significance level of difference between midday and afternoon
cycles: NS, not significantly different; *, P < 0·05; **, P < 0·01.
Time of day (h)
Time of day (h)
(a)
Pressurised A
(% increase over unpressurized value)
Pressurised gs
(% increase over unpressurized value)
Unpressurised gs
(% of maximum daily value)
Unpressurised A
(% of maximum daily value)
(a)
(b)
Time of day (h)
Time of day (h)
Figure 3. Diurnal courses of stomatal conductance from 14
experiments of pressurization and de-pressurization on 10
different plants of two bean cultivars (, G4523, , G5201). (a),
Variations in steady-state stomatal conductance throughout the
day in the unpressurized condition. Values are relativised to the
maximum stomatal conductance observed each day for each
individual plant (set at 100). (b), Percentage increases in gs
following pressurization during the day. The percentages are
calculated in reference to the gs that would have occurred at the
same moment in time, had the plant roots not been pressurized
[i.e. 100((gs,p/gs)-1)], where gs,p is steady-state pressurized gs. Each
data point is normalized to 0·3 MPa of applied pressure
(Mencuccini & Comstock, in preparation). The inset shows that
the residuals from the regression were not systematic with
respect to the Dw under which the experiment was performed.
(b)
Figure 4. Diurnal courses of assimilation rate from 14
experiments of pressurization and de-pressurization on 10
different plants of two bean cultivars (, G4523, , G5201). (a),
Variations in steady-state net assimilation rate throughout the
day in the unpressurized condition. Values are relativised to the
maximum assimilation rate observed each day for each
individual plant (set at 100). (b), Percentage increases in net
assimilation rates above unpressurized values. The increases are
expressed relative to the value of assimilation that would have
occurred had the plant roots not been pressurized (as for gs in
Fig. 3). The coefficients of a linear regression through the data
were not significantly different from zero.
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
1114 M. Mencuccini et al.
Xylem [ABA] (mmol m–3)
Time of day (h)
Xylem [ABA] flux (pmol m–2 s–1)
the early morning compared with midday, but the late afternoon decline was very clear after 1700 h (Fig. 4a).
Pressurization visibly increased water fluxes above their
equilibrium values, with an overall average rise in gs of
34·5% (± 2·1) and in transpiration rate of 21·8% (± 2·4) for
an applied pressure of 0·3 MPa (Fig. 3b). Response to pressurization also followed a clear diurnal trend, in that
stomata showed a constant percentage re-opening during
the morning and midday but progressively reduced
responses during the afternoon. This time-dependency was
present in similar forms in all experiments performed and
for both cultivars. The insert shows that residuals of the
regression against time were not systematic with respect to
the Dw used in each experiment (Fig. 3b, P > 0·05), indicating that leaf-to-air humidity gradients did not interact with
the temporal sequence of our experiments.
Increases in A following pressurization were much
smaller in magnitude, although a significant increase was
almost always detected (Fig. 4b). No relationship with time
of day was apparent.
(a)
(b)
Xylem [ABA] displayed only a subtle diurnal trend, with a
small peak during the early afternoon hours and a decline
afterwards (Fig. 5a). Neither the peak, nor the subsequent
decline were significant at P < 0·05. Not surprisingly, a
similar pattern was also evident for the xylem ABA flux,
again with non-significant differences among times of day
(Fig. 5b). Whole leaf [ABA] showed a reduction from the
early morning hours to midday, after which an almost constant level was maintained throughout the afternoon
(Fig. 5c). The reduction from morning to midday was significant only at P < 0·10.
DISCUSSION AND CONCLUSIONS
Common bean plants responded to root pressurization by
re-opening their stomata and increasing transpiration rates
well above the levels sustained under normal circumstances. The reduced response of A to root pressurization
(Fig. 4a & b) was not unexpected, given the relatively mild
conditions used for the experiments (i.e. low Dw and high
ci) and the probable small degree of stomatal limitation to
photosynthesis.
The experiment was successful in showing that shortterm changes in leaf water status imposed by pressurization
affect stomatal aperture even under well watered conditions and under relatively mild leaf-to-air vapour pressure
gradients (i.e. from 9 to 20 mmol mol-1). Root pressurization has previously been shown to affect stomatal behaviour in a range of woody species, i.e. western water birch
(Betula occidentalis Hook., Saliendra et al. 1995), Douglas
fir (Pseudotsuga menziesii (Mirb.) Franco) and red alder
(Alnus rubra (Bong.) (Fuchs & Livingston 1996), and
Hymenoclea salsola (T. & G.), a desert semi-woody perennial subshrub (Comstock & Mencuccini 1998). However,
earlier reports on herbaceous species failed to obtain
Leaf [ABA] (ng g–1 FW)
(c)
Abscisic acid concentration and fluxes
Time of day (h)
Figure 5. Diurnal variations in abscisic acid (ABA)
concentration and fluxes for six diurnal courses. (a), ABA
concentration in the xylem sap (nM m-3) was determined when
roots were pressurized to obtain re-opening. To express the sap,
roots were first over-pressurized and, when sampling was
completed, returned back to a lower stable pressurization level
(normally between 0·3 and 0·6 bars). Each point is the average of
six diurnal courses. For each diurnal course, three petioles were
used to extract sap under pressurization. (b), Xylem ABA fluxes
obtained from [ABA] and transpiration rates. (c), Variations in
leaf [ABA] (ng g-1 fresh weight) throughout the day. For each
diurnal course, four leaf punches were taken from leaves in
different positions in the canopy. Each plotted value is the
average of all the points taken at that time of day. No significant
diurnal trend was observed in any of the three measured
parameters.
re-opening by root pressurization after imposing a soil
drought (e.g. Gollan, Passioura & Munns 1986; Schurr,
Gollan & Schulze 1992). It is not clear why previous results
on herbaceous plants differ from the present ones. If the
species used in these earlier experiments displayed the
same kind of interaction with time of day as demonstrated
here for P. vulgaris (Fig. 3), then this previously unrecognized effect may have confounded interpretation.
The kinetics of stomatal re-opening after pressurization
evident from Figs 1 and 2 are similar to the ones found by
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
Stomatal responsiveness to leaf water status 1115
Guard cell turgor at constant epidermal turgor pressure (MPa)
Stomatal conductance (mol m–2 s–1)
Comstock & Mencuccini (1998) with H. salsola, i.e. a rapid
hydropassive phase lasting in the order of 10 min followed
by a slower re-opening concluded in about 30 min. They are
similar to the ones given by Saliendra et al. (1995) for
western water birch, i.e. 5 and 20 min, respectively, but substantially longer than the ones found by Fuchs & Livingston
(1996) in Douglas fir and red alder, i.e. 1 and 5 min. The
large kinetic differences observed in different experiments
can only partially be attributed to the specific experimental conditions. For instance, the buffering time of our
cuvette (typically less than 4 min for a 99% change)
reduced the accuracy with which we could determine the
initial very rapid hydropassive responses, but it was small
enough to allow us to clearly monitor the subsequent temporal changes.
The differences among experiments observed after the
‘hydropassive’ stage have more relevance. It is likely that
changes in leaf water status initiated a sequence of metabolic events, which eventually led to the activation of the
stomatal ion pumps and re-opening, i.e. leaf water status
was involved in stomatal control, but only by mediating
some kind of metabolic phenomenon. Instead, the kinetics
of Douglas fir and red alder re-opening (Fuchs &
Livingston 1996) were probably too fast for an interpretation based on the activation of a signal transduction
pathway or of enzyme metabolism. Analogously, Whitehead et al. (1996) reported very rapid changes (within one
minute) in gs in the upper crown of a Pinus radiata D. Don
tree upon the sudden shading of the lower canopy with
removable plastic panels.
Comstock & Mencuccini (1998) interpreted stomatal
responses to soil pressurization on the basis of changes in
total epidermal water potential. We extend here that interpretation using concepts of turgor regulation in guard cells
and in the epidermis, which are diagrammatically illustrated
in Fig. 6. Stomatal conductance (strictly, guard cell volume)
can be described as a function of both guard cell turgor
pressure (Pgc), and of epidermal cell turgor pressure (Pepi)
(Raschke, Dickerson & Pierce 1972, Franks, Cowan & Farquhar1998). For each value of Pepi, gs is a curvilinear and
monotonically increasing function of Pgc, with higher values
reached at Pepi = 0 (Fig. 6, dotted lines with black circles
and squares, for Pepi = 0 and 0·4 MPa, respectively).
Maximum gs is obtained at zero Pepi, because no back-pressure is exerted on the stomatal walls in contact with the subsidiary cells and the epidermis mechanical advantage is lost
(Meidner & Edwards 1996). In our experiments, the equilibrium gs before pressurization was likely to be close to the
curve for Pepi = 0 under conditions of full light and medium
evaporative demands (Meidner & Edwards 1996; Klein
et al. 1996), for instance at point A (Fig. 6). Soil pressurization (for instance, 0·4 MPa) may first act to lower stomatal
conductance to point B, where the hydro-passive closure
despite increased Pgc is explained by the shift to a different
curve at higher Pepi. The subsequent active re-opening may
then bring gs from point B to C on the same curve (i.e. with
higher Pgc but at constant Pepi). However, point C cannot
represent the final equilibrium since the increased transpi-
Figure 6. Interpretation of stomatal responses to soil
pressurization based on epidermal turgor relationships. The two
curves (dotted lines with black circles and squares) depict the
relationship between stomatal conductance and guard cell turgor
Pgc at constant epidermal turgor pressure Pepi, set at the two
values of 0 and 0·4 MPa. The plant was supposed to be at
equilibrium point A, before the soil was pressurized with a
pressure of 0·4 MPa. The starting level of Pepi was likely to be
very close to zero (see text). Movement to point B represents the
initial transient hydropassive closure caused by the back-pressure
of epidermal cells on stomata. The final equilibrium point is not
located on the lower curve (for instance at C) because stomata
re-opening increases transpiration rate and depresses Pepi. The
final point E can be located anywhere in the space between A, B,
C and D. If epidermal water relations are homeostatically
maintained (i.e. they are conserved despite the pressurization),
then point E must be located somewhere on the curve joining A
and D.
ration rate would also tend to lower Yepi and Pepi. This will
establish a positive feedback loop, whereby decreased Pepi
would lead to further re-opening and further decreases in
Pepi. This positive feedback loop is counteracted only by the
decline in Pgc brought about by increased transpiration
rates. The actual trajectory would probably resemble more
closely the curve drawn from A to E (Fig. 6), since these
processes will overlap with each other over time.
Comstock & Mencuccini (1998) presented evidence that
re-opening following pressurization was such that final
equilibrium leaf water potential (Y ) fell back to the value
before pressurization, i.e. it showed evidence of a homeostatic-like behaviour. If this were also the case for the epidermis of common bean in this experiment (i.e. Yepi were
to be conserved), then point E would necessarily be located
on the curve between points A and D, therefore minimizing the epidermal back-pressure.
During our experiments, we found evidence of diurnal
changes in stomatal opening and assimilation rate under
constant environmental conditions (Figs 3a & 4a). Control
experiments performed under constant conditions and
without pressurization confirmed that this was the case
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
1116 M. Mencuccini et al.
(data not shown). This is not unexpected, since P. vulgaris
has frequently been employed to study circadian rhythms
(e.g. Hopmans 1971; Holmes, Sager & Klein 1986; Cardon,
Berry & Woodrow 1994) and is known to have stomatal and
metabolic rhythms around the day–night cycle (Hennessey
& Field 1991; Freeden, Hennessey & Field 1991). The
diurnal responses to root pressurization however, i.e. a
reduced stomatal responsiveness to leaf water status in the
afternoon gave rise to a number of questions.
The lack of response to pressurization was quite distinct
from the daily changes in gs under constant conditions.
While unpressurized gs started declining only after 1630 h,
a reduced re-opening upon pressurization was evident soon
after 1400 h. Because the response to pressurization in the
afternoon was so much slower (Table 1), estimating a reference value for gs in the unpressurized condition was not
simple in some cases (e.g. see Fig. 1). However, inspection
of Fig. 2 reveals that, despite the longer time required to
reach equilibrium, presence or absence of re-opening could
be predicted within 20 min from the start of the pressurization sequence, i.e. in a period within which one could
safely assume that reference gs had remained constant.
Endogenous circadian rhythms may appear as rhythmic
changes in stomatal opening and photosynthesis under constant conditions or as rhythmic changes in sensitivity of the
response to some environmental factor, e.g. light (Freeden
et al. 1991; Hennessey & Field 1991), or temperature (Hennessey & Field 1991). Given the importance of Pgc and Ygc
in determining stomatal aperture, it is hardly surprising that
the responsiveness to water status may also be controlled
by some sort of internal biological clock. However, since
our experiments were only designed to provide systematic
observations on this phenomenon, final conclusions about
a biological clock cannot be drawn from our data.
Our data cannot be explained by a variable leaf orientation through the day. Leaves of common bean are known
to undergo both nastic and tropic movements (e.g. Dubetz
1969; Satter 1979; Berg & Hsiao 1986). However, nyctinastic ‘sleep’ movements rapidly take place toward sunset,
immediately before it gets dark, whereas paraheliotropic
movements occur in response to changes in the direction of
vectorial blue light and are reversible (Koller 1990). Temperature, nutrient availability, soil water stress and intensity
of radiation load, factors that have been shown to modulate this second response (Wainwright 1977; Shackel & Hall
1979; Fu & Ehleringer 1989, 1991; Kao & Forseth 1991),
were also kept constant in our experiments.
The simplest hypothesis to explain the lack of response
to leaf water status, i.e. that there was an afternoon increase
in the concentration or the amount of ABA delivered
through the xylem sap or present in the leaf mesophyll, was
disproved. Both xylem [ABA] and ABA fluxes fluctuated
only slightly around a daily mean, whereas total leaf [ABA]
tended to decline from morning to afternoon. Our values
of xylem [ABA] (30–50 nM) and xylem ABA fluxes
(2–4 pmol m-2 s-1) are broadly similar to those previously
reported for P. vulgaris by Trejo & Davies (1991) and Trejo
et al. (1995).
It is possible that other factors, such as the leaf metabolizing and compartmentalizing capacity for ABA also have
diurnal components that are hitherto unrecognized. Sap pH
is known to fluctuate daily with consistent increases in
the afternoon in Ricinus communis L. (Schurr & Schulze
1995), such that a lower mesophyll sequestration could be
expected at constant xylem [ABA]. Stomatal sensitivity to
[ABA] has also been reported to vary with sap pH in
Heliantus annuus L. (Schurr et al. 1992). Unfortunately, sap
pH could not be measured in these experiments.
How do we explain that unpressurized gs showed no sign
of decline until well after 1630 h, but the responsiveness to
shoot water status was altered throughout the afternoon?
If we assume that metabolic changes in the mesophyll or in
the epidermal apoplast acted to increase the local [ABA]
above xylem [ABA], this should have also determined a
decline in unpressurized gs. That is, here we present a
phenomenon that is different from the more commonly
observed afternoon decline in stomatal conductance and
possibly precedes it.
One possibility is that slight changes occurred in the relationship between guard cell Ygc and pressure (Pgc) via
altered osmotic regulation, such that a slightly lower Pgc
was obtained at constant Ygc in the afternoon. If such
changes were limited to the upper end of the Y–P relationship, then re-opening upon pressurization would have
differed through the day, even though unpressurized gs
appeared unaffected.
Alternatively, it is also possible that changes occurred in
the mechanical properties of the cell wall during the day
such that the response to pressurization changed from
morning to afternoon. Such an event need not be mechanical in nature, e.g. acidification of the cell wall has been
shown to alter the guard cell wall elasticity (Bittisnich,
Entwisle & Neales 1987). Again, changes should have been
limited to the upper end of the turgor response curve. This
is possible, since stomatal opening during the Motorphase
exhibits a two-stage behaviour, according to whether the
cell wall behaves isotropically or anisotropically (Sharpe,
Wu & Spence 1987).
The absence of oscillations during the morning (Table 1)
also deserves some comments. Oscillations in stomatal
aperture have frequently been shown to occur in common
bean (e.g. Hopmans 1971). Theoretical models have been
proposed and successfully used to predict occurrence
of oscillations (Cowan 1972; Delwiche & Cooke 1977;
Haefner, Buckley & Mott 1997), although there is still
uncertainty as to which mechanism is more likely to occur.
In all models however, the source of the oscillation has
been traced to the difference in time-constants of two
co-occurring phenomena, either they being completely
hydraulic (Cowan 1972; Delwiche & Cooke 1977) or partly
bio-chemical (Hafner et al. 1997). Our data cannot be
used to distinguish between the different alternatives,
but they show that the property responsible for creating
oscillations had a component that varied throughout the
day.
Whether the presence of the oscillations and the reduced
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
Stomatal responsiveness to leaf water status 1117
re-opening in the afternoon truly represented a component
of a circadian rhythm or whether it was affected by the temporal sequence of the experimental conditions or by the
accumulated photosynthetic products during the day
cannot be told from our data and will require further experimentation. The observation that the responsiveness to
manipulations of leaf water status did not changed abruptly
at the end of the day, but rather gradually throughout the
whole afternoon would indicate that a biological clock was
not involved (I. Forseth, personal communication).
It is interesting to note the similarity in the interpretation of our results with the one put forward by Franks,
Cowan & Farquhar (1997) to explain the ‘apparent feedforward response’ of some species to Dw, i.e. the irreversible
decline of transpiration rate at high Dw. During previous
experiments with common beans on individual leaves
(Comstock & Ehleringer 1993) and subsequent experiments on whole plants (Mencuccini and Comstock in
preparation), we observed a similar behaviour, and a
decline of transpiration rate at high Dw could not be replicated in the short term (cf. Mott & Parkhurst 1991 for
similar observations on bean plants).
From the experiments reported in Franks et al. (1997), it
is clear that a time dependency of stomatal response to Dw
is present, at least in some species, which is similar to our
observations on the response to pressurization. Franks et al.
(1997) put forward two hypotheses to explain their results:
(a) that the time constant of some hydraulic component in
the pathway from the mesophyll to the epidermis was
larger than the equilibration times normally employed in
cuvette experiments, or (b) that exposure to high Dw sets in
motion a sequence of metabolic events that cannot be interrupted by subsequent exposure to low Dw (Cowan 1995).
It is worth noting that in our case reduced stomatal
re-opening upon pressurization was equally observed at all
the Dw employed during this experiment (see inset in
Fig. 4a), although we intentionally avoided Dw higher than
25 mmol mol-1. Altogether, the two works suggest that
stomatal water relations are affected by processes, possibly
of metabolic origin, which have a time dependency.
Although it was not a feature we observed in all plants,
we also encountered afternoon declines in A at constant ci.
Mott & Parkhurst (1991) commented on the association
between ‘apparent feed-foward response’ and reductions in
A at constant ci, and the possible role of stomatal patchiness in explaining both phenomena. In our experiments,
only in some plants was there evidence that the lack of
response to pressurization was accompanied by a lower A
at constant ci. However, as Mott & Buckley (1998) pointed
out, the absence of a decline in A at constant ci is not sufficient proof to conclude that patchiness was not present
and that it was not interacting with the response to
pressurization.
ACKNOWLEDGMENTS
We wish to thank Rich Raba, who conducted some of the
preliminary gas exchange work and helped in the setting up
of the experiment. The work was supported by USDA grant
#95–37100–1640.
REFERENCES
Berg V.S. & Hsiao T.C. (1986) Solar tracking: light avoidance
induced by water stress in leaves of kidney bean seedlings in the
field. Crop Science 26, 980–986.
Bittisnich D.J., Entwisle L.O. & Neales T.F. (1987) Acid-induced
stomatal opening in Vicia faba L. & the role of guard cell elasticity. Plant Physiology 85, 554–557.
Cardon Z.G., Berry J.A. & Woodrow I.E. (1994) Dependence of
the extent and direction of average stomatal response in Zea
mays L. & Phaseolus vulgaris L. on the frequency of fluctuations
in environmental stimuli. Plant Physiology 105, 1007–1013.
Comstock J. & Ehleringer J. (1993) Stomatal response to humidity
in common bean (Phaseolus vulgaris): implications for
maximum transpiration rate, water-use efficiency and productivity. Australian Journal of Plant Physiology 20, 669–691.
Comstock J. & Mencuccini M. (1998) Control of stomatal conductance by leaf water potential in Hymenoclea salsola (T. & G.), a
desert subshrub. Plant, Cell and Environment 21, 1029–1038.
Cowan I.R. (1972) Oscillations in stomatal conductance and plant
functioning associated with stomatal conductance: observations
and a model. Planta 106, 185–219.
Cowan I.R. (1977) Stomatal behaviour and environment. Advances
in Botanical Research 4, 117–228.
Cowan I.R. (1995) As to the mode of action of guard cells in dry
air. In Ecophysiology of Photosynthesis (eds E.-D. Schulze &
M.M. Caldwell), pp. 205–229, Springer, Berlin, Germany.
Davies W.J. & Zhang J. (1991) Root signals and the regulation of
growth and development of plants in drying soil. Annual Review
of Plant Physiology and Molecular Biology 42, 55–76.
Delwiche M.J. & Cooke R.J. (1977) An analytical model of the
hydraulic aspects of stomatal dynamics. Journal of Theoretical
Biology 69, 113–141.
Dixon M.A. & Tyree M.T. (1984) A new stem hygrometer, corrected for temperature gradients and calibrated against the pressure bomb. Plant, Cell and Environment 7, 693–697.
Dubetz S. (1969) An unusual photonastism induced by drought in
Phaseolus vulgaris. Canadian Journal of Botany 47, 1640–
1641.
Franks P.J., Cowan I.R. & Farquhar G.D. (1997) The apparent feedforward response of stomata to air vapour pressure deficit: information revealed by different experimental procedures with two
rainforest trees. Plant, Cell and Environment 20, 142–145.
Franks P.J., Cowan I.R. & Farquhar J.D. (1998) A study of stomatal
mechanics using the cell pressure probe. Plant, Cell and Environment 21, 94–100.
Freeden A.L., Hennessey T.L. & Field C.B. (1991) Biochemical correlates of the circadian rhythm in photosynthesis in Phaseolus
vulgaris. Plant Physiology 97, 415–419.
Fu Q.A. & Ehleringer J.R. (1989) Heliotropic leaf movements in
common bean controlled by air temperature. Plant Physiology
91, 1162–1167.
Fu Q.A. & Ehleringer J.R. (1991) Modification of paraheliotropic
leaf movements in Phaseolus vulgaris by photon flux density.
Plant, Cell and. Environment 14, 339–343.
Fuchs E.E. & Livingston N.J. (1996) Hydraulic control of stomatal
conductance in Douglas fir (Pseudotsuga menziesii (Mirb.)
Franco) and alder (Alnus rubra (Bong.) seedlings. Plant, Cell and
Environment 19, 1091–1098.
Gollan T., Passioura J.B. & Munns R. (1986) Soil water status
affects the stomatal conductance of fully turgid wheat and
sunflower leaves. Australian Journal of Plant Physiology 13,
459–464.
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118
1118 M. Mencuccini et al.
Grantz D.A. (1990) Plant response to atmospheric humidity. Plant,
Cell and Environment 13, 667–679.
Haefner J.W., Buckley T.N. & Mott K.A. (1997) A spatially explicit
model of patchy stomatal responses to humidity. Plant, Cell and
Environment 20, 1087–1097.
Hennessey T.L. & Field C.B. (1991) Circadian rhythms in photosynthesis. Plant Physiology 96, 831–836.
Holmes M.G., Sager J.C. & Klein W.H. (1986) Sensitivity to far-red
radiation in stomata of Phaseolus vulgaris L. rhythmic effects on
conductance and photosynthesis. Planta 168, 516–522.
Hopmans P.A.M. (1971) Rhythms in stomatal opening of bean
leaves. In Mededelingen LandBouwhogeschool. 71–3, p. 86. H.
Veenman & Zonen N.V., Wageningen, The Netherlands.
Jones H.G. (1987) Correction for non-specific interference in competitive immunoassays. Physiologia Plantarum 70, 146–154.
Kao W.Y. & Forseth I.N. (1991) The effects of nitrogen, light and
water availability on tropic leaf movements in soybean (Glycine
max). Plant, Cell and Environment 14, 287–293.
Koller D. (1990) Light-driven leaf movements. Plant, Cell and
Environment 13, 615–632.
Klein M., Cheng G., Chung M. & Tallman G. (1996) Effects of
turgor potential of epidermal cells neighbouring guard cells on
stomatal opening in detached leaf epidermis and intact leaflets
of Vicia faba L. (faba bean). Plant, Cell and Environment 19,
1399–1407.
Losch R. & Tenhunen J.D. (1981) Stomatal responses to humidity
– phenomenon and mechanism. In Stomata Physiology (eds P.G.
Jarvis & T.A. Mansfield), pp. 137–161, Cambridge University
Press, Cambridge, UK.
Meidner H. & Edwards M. (1996) Osmotic and turgor pressures of
guard cells. Plant, Cell and Environment 19, 503.
Mencuccini M. & Comstock J. (1999) Variability in hydraulic architecture and gas exchange of common bean (Phaseolus vulgaris)
cultivars under well-watered conditions: interactions with leaf
size. Australian Journal of Plant Physiology 26, 115–124.
Monteith J.L. (1995) A reinterpretation of stomatal responses to
humidity. Plant, Cell and Environment 18, 357–363.
Mott K.A. & Buckley T.N. (1998) Stomatal heterogeneity. Journal
of Experimental Botany 49, 407–417.
Mott K.A. & Parkhurst D.F. (1991) Stomatal responses to humidity in air and helox. Plant, Cell and Environment 14, 509–515.
Ober E., Setter T.L., Madison J.T., Thompson J.F. & Shapiro P.S.
(1991) Influence of water deficit on maize endosperm development. Enzyme activities and RNA transcripts of starch and zein
synthesis, abscisic acid and cell division. Plant Physiology 97,
154–164.
Raschke K., Dickerson M. & Pierce M. (1973) Mechanics of stomatal responses to changes in water potential. In: Plant Research
pp. 155–157. MSU/AEC, Plant Research Laboratory, Michigan
State University, East Lausing.
Saliendra N.Z., Sperry J.S. & Comstock J. (1995) Influence of leaf
water status on stomatal response to humidity, hydraulic conductance, and soil drought in Betula occidentalis. Planta 196,
357–366.
Satter R.L. (1979) Leaf movements and tendril curling. In Physiology of Movement. Encyclopedia of Plant Physiology (N S ), Vol
7 (eds W. Haupt & M.E. Feinleib), pp. 442–484, Springer-Verlag,
Berlin.
Schulze E.-D. (1993) Soil water deficits and atmospheric humidity
as environmental signals. In Water Deficits. Plant Responses from
Cell to Community (eds J.A.C. Smith & H. Griffith), pp. 129–145,
Bios Scientific Publisher, Oxford.
Schurr U. & Schulze E.-D. (1995) The concentration of xylem sap
constituents in root exudate, and in sap from intact, transpiring
castor bean plants (Ricinus communis L.). Plant, Cell and Environment 18, 409–420.
Schurr U., Gollan T. & Schulze E.-D. (1992) Stomatal response to
drying soil in relation to changes in the xylem sap composition
of Helianthus annuus. II. Stomatal sensitivity to abscisic acid
imported from the xylem sap. Plant, Cell and Environment 15,
561–567.
Shackel K.A. & Hall A.E. (1979) Reversible leaflet movements in
relation to drought adaptation of cowpeas, Vinga unguicolata
(L.) Walp. Australian Journal of Plant Physiology 6, 265–276.
Sharpe P.J.H., Wu H. & Spence R.D. (1987) Stomatal mechanics. In
Stomatal Function (eds E. Zieger, G.D. Farquhar & I.R. Cowan),
pp. 91–114, Stanford University Press, Stanford, CA, USA.
Singh S.P., Gepts P. & Debouck D.G. (1991) Races of common bean
(Phaseolus vulgaris, Fabaceae). Economic Botany 45, 379–396.
Trejo C.L. & Davies W.J. (1991) Drought-induced closure of Phaseolus vulgaris L. Stomata precedes leaf water deficit and any
increase in xylem ABA concentration. Journal of Experimental
Botany 42, 1507–1515.
Trejo C.L., Clepham A.L. & Davies W.J. (1995) How do stomata
read abscisic acid signals? Plant Physiology 109, 803–811.
von Caemmerer S. & Farquhar G.D. (1981) Some relationships
between the biochemistry of photosynthesis and the gas
exchange of leaves. Planta 153, 376–387.
Wainwright C.M. (1977) Sun tracking and related leaf movements
in lupine Lupinus arizonicus. American Journal of Botany 64,
1032–1034.
Walker-Simmons M. (1987) ABA levels and sensitivity in developing wheat embryos of sprouting resistant and susceptible cultivars. Plant Physiology 84, 61–66.
Whitehead D., Livingston N.J., Kelliher F.M., Hogan K.P., Pepin S.,
McSeveny T.M. & Byers J.N. (1996) Response of transpiration
and photosynthesis to a transient change in illuminated foliage
area for a Pinus radiata D. Don tree. Plant, Cell and Environment 19, 949–957.
Received 8 January 2000; received in revised form 1 June 2000;
accepted for publication 1 June 2000
© 2000 Blackwell Science Ltd, Plant, Cell and Environment, 23, 1109–1118