CSIRO PUBLISHING
Functional Plant Biology, 2010, 37, 1105–1116
www.publish.csiro.au/journals/fpb
Path of water for root growth
John S. Boyer A,C, Wendy K. Silk B and Michelle Watt C,D
A
College of Earth, Ocean and Environment (formerly College of Marine Studies),
University of Delaware, 700 Pilottown Road, Lewes, DE 19958, USA.
B
Department of Land, Air and Water Resources, University of California, Davis,
CA 95616-8627, USA.
C
CSIRO Plant Industry, GPO Box 1600, Canberra, ACT 2601, Australia.
D
Corresponding author. Email: [email protected]
Abstract. Do roots obtain water for their growth directly from soil surrounding the growth zone or indirectly, via phloem,
from water absorbed elsewhere? Wheat (Triticum aestivum L.) was studied with time-lapse imaging of seminal axile roots,
growing in soil and air in a custom-made laboratory rhizotron, before and after excision. The growth data were combined with
a theoretical estimate of the amount of water that could be supplied from the phloem. Roots readily extended into air,
providing strong evidence that they obtain a portion of their growth-sustaining water internally. The time-lapse experiments
indicated that in moist soil, internal sources provided 26–45% of the water for root growth, but the rest came externally from
the soil surrounding the growth zone. From the theoretical analysis, the phloem could supply, on average, 64% of the total,
accounting for all the internal sources. This indicates that phloem water could be used when root tips cannot access external
water, such as in cracks or pores, or regions of dry soil. The distribution of phloem-delivered water for root growth should be
considered in whole-plant modelling of root systems. Maximising phloem flux at root tips might confer more efficient use of
soil water by crops.
Additional keywords: branch, extension, phloem, soil, solutes, stress.
Introduction
Most of the water absorbed by roots is sent to the shoot in the
xylem, but some is retained for the functioning of root cells and
the growth of apical cells that require water to extend into new soil
volumes (Boyer and Silk 2004; Wiegers et al. 2009). To a first
approximation, the cell grows by taking up water and stretching
under the resultant turgor pressure; the mass of young root cells is
more than 90% water (see ‘Results’). The water could be coming
internally from xylem or with solute delivered by phloem.
Alternatively, the water could be entirely or partially supplied
by external water absorbed directly from the soil surrounding the
growing tissue. The importance of these paths is suggested by the
observation that, as soil water is depleted around the mature root
tissues of the transpiring plant, the moister soil around the tip can
provide the water needed for continuing growth into the moister
soil layers (Kramer and Boyer 1995). Conversely, after rainfall,
irrigation, or hydraulic lift, the internal pathway would allow
surface roots to supply water to roots in deeper, drier layers.
Despite much study of plant water relations, there is little
information about the pathway of water for root growth. Water
potential gradients favour water movement towards the root tip
even though the growing tip itself has no functional xylem (Sharp
and Davies 1979; Westgate and Boyer 1985; Zwieniecki et al.
2003). Substantial negative water potentials have been observed
in growing root tips (Sharp and Davies 1979; Westgate and Boyer
1985; Rygol et al. 1993) and water potential gradients within the
CSIRO 2010
growing tip were predicted by Wiegers et al. (2009) using a threedimensional mathematical analysis. Although the tips would
require these gradients in order to move water through the
tissue (Zwieniecki et al. 2003; Wiegers et al. 2009), water
could be moved towards this region from behind the tip in
maturing xylem. Closer to the tip, mature phloem is functional
and would deliver water along with photosynthetic products
(Heimsch 1951; Esau 1953; Popham 1955). Bret-Harte and
Silk (1994) concluded from mathematical analysis that up to
81% of tip growth could be supplied from this source. Pritchard
and his co-workers (Pritchard 1996; Pritchard et al. 2000) studied
the dilution of phloem-delivered K+ in the growing tip cells of
barley roots and concluded that as much as 78% of the water was
delivered by the phloem. This suggests that external water
absorbed directly from the local soil around the tip might
contribute little to root elongation. Indeed some roots grow in
air in the absence of soil in maize (Zea mays L.), orchids and
certain Ficus species.
However, when water surrounds the root tip in soil, there
would seem to be nothing to prevent its direct uptake by the tip.
Because this possibility appears unexplored experimentally, we
hypothesised that the soil surrounding the root tip can be an
important source of water for growth. If the hypothesis is correct,
(1) root tips should grow more slowly in air than in soil, and (2)
should continue to grow after excising the tip in moist soil, where
water would continue to be available. These two tests were
10.1071/FP10108
1445-4408/10/121105
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J. S. Boyer et al.
employed in the following study, and the results indicated that
although roots grow in the absence of water immediately around
the tip, growth was always slower than when the tip had an
external local supply. Moreover, without an external supply, the
slower tip growth could be supported by the amount of water
delivered by phloem even if no water came from the xylem. This
finding supports the complementary hypothesis that water
delivered by phloem is centrally important for roots growing
in drying soils.
Materials and methods
Growing conditions
Wheat (Triticum aestivum L. cv. Janz) plants were grown in a
rhizobox that allowed roots to be photographed and accessed
through a transparency covering the face (Fig. 1). The rhizobox
was constructed from black Perspex (ACT Plastics, Fyshwick,
Canberra, ACT, Australia) 6 mm thick and had internal
dimensions 250 mm wide 500 mm high 10 mm deep. Two
rails divided the rhizobox into three open top compartments
containing soil with two plants in each. A stainless steel mesh
provided a support for the soil above, which had a depth of
~15 cm. Below the mesh was air kept saturated by a strip of thick
wet filter paper to simulate conditions in an air gap in soil. The
1-mm openings in the mesh allowed the roots to grow through
from soil to air because root diameters averaged only 0.55 mm
(Table 1, Line 4).
The rhizobox was first assembled without soil. After placing
wet filter paper in the lower compartments, fresh transparencies
(Transparency Film PP2500, 3M, Minneapolis, MN, USA) were
laid on the front and were supported by the rails outlining the sides
Moist soil
Seminal root axis
Wire mesh
Saturated air
Petrolatum
10 cm
Fig. 1. Root rhizobox containing soil and wheat (Triticum aestivum L.)
plants. Roots are visible in the soil and below the soil in the saturated air. Air
was kept saturated by wet filter paper. Petrolatum held a root in air in the centre
compartment. Transparent tape covered the access flap through which the root
was cut.
and bottom and separating the three compartments. Each
transparency was fixed to the front with bolts that ran through
a white plastic strip, through the transparency, then to the back of
Table 1. Measured and calculated properties of roots growing in soil
Calculations use measured properties from this table and averages from Table 2, as described in text
Line no.
1
2
3
4
Measured
Average
Range
0.0103
0.0093–0.0120 (n = 3)
0.173
0.168–0.178 (n = 3)
1.21
0.85–1.48 (n = 20)
0.275
0.273–0.293 (n = 58)
Equation
Calculation
Persistent DW DLength DPersistent DW
¼
Length
hour
hour
(0.0103) (1.21) = 0.0125
DPeristent DW Total DW Delivery Total DW Delivery
¼
hour
DPersistent DW
hour
(0.0125) (1.89) = 0.0236
Total DW Delivery Phloem Water Delivery Phloem Water Delivery
¼
hour
Total DW Delivery
hour
(0.0236) (5.3) = 0.125
FW
DW
Water Content
¼
Length Length
Length
0.173–0.0103 = 0.163
Water Content DLength DWater Content
¼
Length
hour
hour
(0.163) (1.21) = 0.197
Persistent DW
ðmg mm1 Þ
Length
FW
ðmg mm1 Þ
Length
DLength
ðmm h1 Þ
hour
Radius (mm)
Calculated
5
Accumulation of persistent
DW (mg h–1)
6
Delivery of total DW
by phloem (mg h–1)
7
Volume delivered by
phloem (mg h–1) or (mm3 h–1)
8
Root water content
(mg mm–1)
9
Rate of water accumulation
(mg h–1) or (mm3 h–1)
10
Rate of root volume growth
(mm3 h–1)
pr2
DLength DVolume
¼
hour
hour
(3.14) (0.275)2 (1.21) = 0.287
Path of water for root growth
the rhizobox where each was tightened with a nut (Fig. 1). The
strips held each transparency tightly and ensured tight seams. Any
open seams between transparencies were taped to eliminate air
leaks. A small drainage slit at the bottom allowed excess water to
drain from the rhizobox.
The rhizobox was weighed without soil, soil was added
through the open top, and the whole unit was re-weighed. The
soil was a mix of 50% river sand and 50% recycled potting
soil (organic matter, loam and sand supplemented with lime,
14.4% N, 6.6% P and 5% K). A sample of soil was dried for at
least 24 h to determine the dry weight (DW) of soil in the
rhizobox. After loading with soil, modified half strength
Hoagland’s nutrient solution (in mg L–1; N 212, P 32, K 236,
Ca 161, Mg 48, Na 7.9, S 70, Cl 0.14, Fe 5.0, B 0.11, Co 0.005, Mn
0.11, Cu 0.013, Zn 0.02, Mo 0.012) was added until the soil
drained. After an hour or two for drainage, the rhizobox was
weighed again. These measurements gave the weight of dry soil
and the maximum water content of the soil when drained under
the force of gravity (0.23 g water per g dry soil). The high sand
content ensured rapid drainage and gave a high air-filled porosity
of 20% after draining in the rhizobox. With water removal by the
plants, the air-filled porosity increased. By adding small amounts
of the nutrient solution to the soil surface by weight, water
contents could be kept between 60 and 90% (0.14–0.21 g
water per g dry soil) of the maximum water content of the soil
when drained under the force of gravity. This ensured that no part
of the soil was water saturated during the measurements and
the high soil porosity allowed air to reach the roots. The bulk
density of the soil mix was low (1.18 0.03 g cm–3). The soil
strength of the drained moist soil also was low, and the average
penetrometer resistance of the soil within the compartment and
the soil at the interface between the transparency and the
compartment was 0.3 0.07 MPa.
Wheat seeds were planted 2 cm below the soil surface,
embryo facing down and adjacent and towards the
transparency face. The top of the rhizobox and the shoots were
exposed to outside air and light. The irradiance above the soil
was 500 mmol m–2 s–1 with a photoperiod of 12 h supplied by
a metal halide bulb. Temperatures were 1622C and the RH was
uncontrolled but generally around 40% during the day. Root
growth was measured when the plants were 4–17 days old.
Time-lapse photography
The rhizoboxes were mounted in a frame custom-made for
time-lapse photography of root growth (Fig. 2). The frame
allowed the rhizobox to be tilted to different angles while
maintaining the same position between the transparency face
and the camera, also mounted to the frame. The frame was
adjusted to control whether root growth followed the
transparencies, or grew away from the transparencies. Dim
green safelight was supplied to the face of the rhizobox by
light emitting diodes. A digital camera (Nikon CoolPix 5400,
equipped with a Nikon MC-EU1 remote controller and a 1 Gb
card for photograph storage) was mounted ~20 cm in front of
the transparencies. Time-lapse images (2.5 Mb) were recorded
every 15 min. When an experiment was complete, images were
downloaded to a computer, and the rate of root elongation in each
15 min interval was determined by using image analysis software
Functional Plant Biology
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Rhizobox
Camera
10 cm
Green LED lights
Fig. 2. Apparatus to hold rhizobox and camera parallel to each other for
time-lapse imaging of root growth. In order to protect against exposing the
roots to white light, the entire apparatus was enclosed in a light-tight box with
plant shoots exposed to light overhead. Roots were exposed only to dim green
light from the LEDs inside.
(either Canvas X, Build 885, ACD Systems Inc., Victoria, BC,
nster,
Canada; or analySIS Soft Imaging System, GmbH, Mu
Germany). The length measurements were reproducible within
0.1 mm and the times were indicated to within 1 min by the timelapse camera. In each time-lapse series, lengths were calibrated by
measuring part of the apparatus with known dimensions in the
image, usually one of the bolt heads holding the rhizobox
together.
Hypothesis test 1: Time-lapse measurements of intact
roots in soil and air to measure the contribution of soil
water to growth
The hypothesis implies that intact roots should grow more slowly
in air than in soil. Between the single and two leaf stage, growth of
the seminal roots of the wheat seedlings (generally three per plant)
was measured in soil, and then root tips were checked for passage
through the mesh into the saturated air below, where soil was
absent (Fig. 1). If the roots were overly bent their growth was not
reported. If roots became invisible behind soil particles,
measurements were abandoned. If the roots touched the
transparency when growing in saturated air, there was a
possible water path from the soil along the contact surface,
and measurements were rejected. This gave an overall
rejection rate of ~50% and ensured that the reported growth
was only for reasonably straight, intact roots. Roots were
photographed every 15 min, as described above. A feature in
the soil, such as a cluster of particles, or on the root, such as a root
hair, was used as a position reference for these measurements.
Hypothesis test 2: Time-lapse measurements before
and after excision to measure the contribution
of internal water to growth
For the second test of the hypothesis, if some water for root growth
comes directly from the soil surrounding the growth zone, growth
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Functional Plant Biology
should continue after the root tip is excised in soil. Seminal roots
similar to those in test 1 and shown in Fig. 1 were excised in soil
for this test. When the roots were in soil and their root tip visible, a
sharp scalpel cut cleanly through the transparency and across
the root axis at a position 12–22 mm behind the tip. Because the
elongating zone did not extend beyond 10 mm, the cuts were
always behind the elongating zone. The root tip continued to be
held in place by the soil. Tip elongation was observed every
15 min by time-lapse imaging before and after the excision, as
described above.
For roots in air, a flap was opened in the transparency and
petrolatum was injected between the root and the transparency
and on the other side of the root before imaging began. The
petrolatum provided a gentle holder for the root and kept the root
from touching the transparency. After the petrolatum was in
place, the flap was sealed with transparent tape and time lapse
observations began on the intact root (Fig. 1, middle panel). After
at least 2 h of further elongation, the rhizobox was removed, the
flap was opened, and a sharp miniature scissors was used to cut
17–25 mm behind the tip (i.e. behind the elongating zone) and
above the petrolatum,. The petrolatum continued to hold the root
tip in place after the cut, and the flap was re-sealed with the
transparent tape, usually in less than 1 min. The rhizobox was
replaced in the tilting frame and observations continued. The
observations were made every 15 min by time-lapse imaging
before and after the excision, as described above.
Destructive measurements to determine the amount
of water and solute deposited in roots growing in soil
Seedlings were grown in soil in the rhizoboxes to the single leaf
stage, and their root positions marked every day for 2–3 days until
they were 100–150 mm long and about to emerge into the
saturated air (6–8 days). The transparency was then removed
and the entire seedling gently lifted to free the roots from most of
J. S. Boyer et al.
the soil. The seedlings were immediately submerged in water and
the roots gently stroked under water with a soft paint brush to
remove adhering soil particles. When clean, the roots were
excised 2 cm below the base. Then, 2 cm of the root tip was
removed so that the remaining segment was mature. The mature
segments were viewed in a dissecting microscope to measure
diameter using an ocular micrometer usually within 2 cm of
the ends of the segment but occasionally at several positions
along the entire length. The total length of these segments was
measured with a submerged ruler. The segments were then
removed from the water, blotted gently, and weighed to give
fresh weight (FW) after which they were dried at 70 for 24 h
in an aluminum foil envelop, cooled in a desiccator and
weighed to give DW. Three roots from each of six plants (total
of 18 roots) were pooled for the weight determinations. Except
where noted, the statistical variation is reported as the range
in order to allow a sensitivity analysis of the range of variation
to be made.
Results
Roots readily grew from soil to air on all plants grown in the
rhizoboxes (Fig. 3). Because nutrient solution was supplied
sparingly (60–90% of field capacity), there were no saturated
soil regions or water flowing over the surface of the roots in air. All
of the experiments were conducted during the day when the plants
were transpiring and the water in the root xylem was under
tension. This could be seen in those roots in air that followed
the transparency face when condensate was present (early
morning). The condensate was removed for ~1 mm around the
entire exposed root and root hairs, indicating that the root was
absorbing water from its surface. When roots were in air without
touching the transparency, no liquid water was seen on the root
surface. In the latter condition, water for growth appeared to come
Fig. 3. Image of root box with roots extending into the saturated air compartment. Arrows show position of
screen separating compartments for soil and air. Roots growing in air did not touch any surface. Note that the
position of the root hair zone varies among different roots that have entered the air at about the same time (circles).
Path of water for root growth
Functional Plant Biology
only from internal sources corrected for any that might come from
or be lost to the air.
Hypothesis test 1: Time-lapse measurements of intact
roots in soil and air to measure the contribution
of internal water to growth
If a root grows in air, soil no longer surrounds its tip, and soil water
cannot contribute directly to its growth. Elongation rate in air
was only 45% of that in soil (Fig. 4a, b, Intact). Six replicate
roots grew at a rate of 1.39 0.23 mm h–1 in soil, and another six
grew at a rate of 0.63 0.17 mm h–1 in air.
Hypothesis test 2: Time-lapse measurements before
and after excision to measure the contribution
of internal water to growth
If the root tips growing in soil were excised to eliminate water
delivery from internal sources, their growth rate of 1.39 mm h–1
decreased to 1.03 mm h–1 or 74% of the intact rate where it
continued for at least 30 min (Fig. 4a, Soil, Intact v. Cut). As a
control, tips were also excised from roots growing in air and
reliant only on internal water sources. Upon excision, the tips
grew at 5% of the intact rate for the next 30 min, which was
statistically insignificant (Fig. 4b, Air, Cut).
Sometimes an intact root grew into the photographic field of
the excised root (Fig. 5a). Cutting one root had little effect on the
growth of the neighbouring intact root that continued to grow at
the original rate (Fig. 5b). Similarly for roots growing in air
(Fig. 6a), another nearby root left intact continued to grow rapidly
after growth nearly ceased in the accompanying, excised one
(Fig. 6b). Close inspection indicates that root hairs often
developed gradually close to the tip in the excised ones but not
in the intact ones (Fig. 6b).
Destructive measurements to approximate the amount
of water and solute deposited in roots growing in soil
Table 1 indicates that the roots grew rapidly in soil, averaging
1.21 mm h–1 when reasonably straight, with a relatively wide
range of rates (0.84–1.48 mm h–1). Compared with this variation,
root DWs (0.0103 mg mm–1, range 0.0093 to 0.0120) and FWs
(0.173 mg mm–1, range 0.168 to 0.178) were quite uniform
because 18 roots were pooled for each measurement. Also,
root radii were uniform, averaging 0.275 mm (range 0.273 to
0.293) with no branches along the segment axis.
(b)
Average elongation
rate (mm h–1)
(a)
Fig. 4. Rates of elongation by intact wheat (Triticum aestivum L.) roots
growing in (a) soil or (b) air. Also shown are rates during 30 min after cutting
the same roots (Cut). Rates are averages s.e. (n = 6 in soil, 6 in air). Roots
growing in air did not touch any surface.
1109
It was possible to use these measured properties to calculate
the amount of water delivered by the phloem to the root tip.
Solute delivered as a solution by the phloem is polymerised
into cell walls and cytoplasmic constituents that account for
most of the persistent DW of the tissues. In addition to this
DW, Table 2 gives pulse-chase results reported by Dilkes et al.
(2004) and Gregory and Atwell (1991) showing that 33–48% of
the delivered solute is respired by the roots of wheat, and 1–11% is
exuded from the root surface. These values are similar to those of
other grass species (Table 2). Accordingly, the persistent DW of a
wheat root (Table 1) can be considered to be 51–55% of the total
solute delivered by the phloem (Table 2). Taking 53% as the
average for wheat, the total DW delivered by the phloem was 1/
0.53 = 1.89 times the persistent DW delivery, or 0.0236 mg h–1
(Table 1, Lines 5 and 6).
The water delivered with the DW depends on the
concentration of the phloem solution. According to Table 2,
reported average concentrations of the phloem solution in
wheat were 386 mM sucrose (132 g L–1), 192 mM amino acids
(23 g L–1, assuming average molecular weight is 118), and
123 mM K+ (4.8 g L–1), or ~160 g L–1 of phloem solution. With
the assumption that one litre of this solution contained at least
840 g of water (i.e. the density of phloem solution was the density
of water), the phloem could deliver ~5.3 weight units of water
with every weight unit of solute or 0.125 mg of water every hour
(Table 1, Line 7).
For a comparison with the amount of water delivered by
phloem, the average amount of water actually accumulating in
the root can be calculated from the water content of the mature root
tissues (Table 1, Line 8) and the growth rate of the roots
(Table 1, Line 9). Comparing this amount (0.197 mm3 h–1) to
the average amount of phloem-delivered water (0.125 mm3 h–1
from Table 1, Line 7), the water delivered by the phloem could
account for as much as 64% of the total water accumulated by the
root tip as it grows in soil, on average.
But considerable variation exists in this estimate and can be
evaluated from a sensitivity analysis of the calculation. For
simplicity, the basic equations can be combined to
approximate the rate of water delivery by the phloem. From
Table 1, substituting the equation from Line 5 into 6, and Line 6
into 7 gives the rate of phloem water delivery in terms of DW and
root length:
Persistent DW DLength Total DW Delivery
Length
hour
DPersistent DW
Phloem Water Delivery Phloem Water Delivery
¼
Total DW Delivery
hour
ð1Þ
This equation is maximised when all the numerator terms
are at their maximum and all the denominator terms are at
their minimum. The equation is minimised when this
situation is reversed. Maxima and minima were taken from
the range of observations in Tables 1 and 2 calculated in
Table 3. Water delivered by phloem could be as little as
0.030 mm3 h–1 or as much as 0.341 mm3 h–1, or 15–173% of
the 0.197 mm3 h–1 of water observed to accumulate in the
growing root tip (cf. Table 3, last line and Table 1, Line 9).
The maximum delivery rate suggests enough water is provided
1110
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J. S. Boyer et al.
Uncut
(a)
To be
cut
(b)
Uncut Before
cut
Uncut
Uncut Before
cut
After
cut
Uncut
After
cut
5 mm
Change in length (mm)
8
6
4
2
cut
0
0
50
100
150
200
250
300
Time (min)
Fig. 5. Details for wheat (Triticum aestivum L.) roots growing in soil and shown in Fig. 4a (Intact v. Cut). (a)
Two root tips are visible, one to be excised and the other to remain intact. (b) The two root tips in (a) grow.
Horizontal bars in images on right indicate the position of the cut (After cut). Graph at bottom shows details of
growth for the cut (closed circles) and uncut roots (open circles). Note that growth continues for a time after cutting
that deprives the tip of an internal water supply. Data are individual lengths measured from time-lapse images like
those above the graph.
to cause exudation from the root surface; and, indeed,
this is sometimes observed (Passioura 2002; Tumlinson et al.
2008).
It should be noted that this comparison concerns only the water
delivered to the root to build mature root tissue. It does not take
into account the volume of non-aqueous structures occupied by
the persistent DW and the intercellular spaces. Although
considering these additional structures does not alter
the conclusions from the root length measurements, the
nonaqueous structures do contribute volume to the
photographic images of the roots used in the present
experiments. The accumulating water occupied 69% of the
root volume (cf. Table 1, Lines 9 and 10) and non-aqueous
structures occupied the rest.
Path of water for root growth
Functional Plant Biology
1111
(a)
Uncut
To be
cut
(b)
Uncut Before
cut
Uncut Before
cut
Uncut
Uncut
After
cut
After
cut
5 mm
Change in length (mm)
12
10
8
6
4
2
cut
0
0
100
200
300
400
500
Time (min)
Fig. 6. Details for wheat (Triticum aestivum L.) roots growing in air and shown in Fig. 4b (Intact v. Cut). (a) Two
root tips are visible, one to be excised and the other to remain intact. Also shown is the petrolatum around the root
to be cut (top of image). (b) The two root tips in (a) grow. Horizontal bars in images on right indicate the position of
the cut above the petrolatum, which holds the root segment without allowing contact of the segment with the
transparency (although the petrolatum may slide along the transparency). Graph at bottom shows details for the
cut (closed circles) and uncut roots (open circles). Note that, in contrast to Fig. 5b in soil, growth ceases
immediately after cutting in air. Data are individual lengths measured from time-lapse images like those above
the graph.
Discussion
The results of this study support the hypothesis that water for
root growth can come directly from the soil surrounding the tip.
The roots always grew faster when this external supply
accompanied internal sources. The experiments indicated that
in moist soil internal sources provided 26–45% of the total, and
the soil surrounding the growth zone supplied the rest
(summarised in Table 4, Lines 1 and 2). At least some of the
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Functional Plant Biology
J. S. Boyer et al.
Table 2. Reported properties of whole root systems and phloem solution
Root properties are DW fractions from 14C pulse-chase experiments, with average and (range of values). Phloem properties are average concentrations of exudate
from aphid stylets, with (range of values)
Persistent DW
Whole-root system
Respired DW
Exuded DW
0.55
0.51
0.57
0.51
0.46
0.52 (0.46–0.57)
0.33
0.48
0.42
0.47
0.41
0.42 (0.33–0.48)
0.11
0.01
0.01
0.03
0.13
0.06 (0.01–0.13)
Sucrose (mM)
Phloem solution
Amino acids (mM)
K+ (mM)
251
409 (342–476)
278 (270–287)
452
538 (512–564)
–
386 (251–564)
262
86 (62–110)
148 (140–156)
56 (52–60)
138 (133–143)
460 (134–1037)
192 (52–1037)
Reference
Crop
Pulse conditions
Dilkes et al. (2004)
Gregory and Atwell (1991)
Farrar (1985)
Gregory and Atwell (1991)
Nguyen et al. (1999)
Average (range)
15–19 days wheat
50 days wheat
16 days barley
49 days barley
35 days maize
20 h after pulse
24 h after pulse
CO2 release, 1.25 h pulse
24 h after pulse
92 h after pulse
Stylet position
299
40 (38–41)
–
30 (25–34)
–
–
123 (25–299)
Hayashi and Chino (1986)
Fisher (1987)
Fisher and Gifford (1986)
Fisher (1987)
Fisher and Gifford (1986)
Gattolin et al. (2008)
Average (range)
Pre-anthesis wheat
Anthesis wheat
Post-anthesis wheat
Anthesis wheat
Post-anthesis wheat
Pre-anthesis wheat
Night, leaf sheath
Night, peduncle
Night, peduncle and grain
Day, peduncle
Day, peduncle and grain
Day, leaf sheath
Table 3. Sensitivity analysis of the contribution of phloem-delivered water to the growth of wheat roots
Persistent DW
ðmg mm1 Þ
Length
DLength
ðmm h1 Þ
hour
Total DW Delivery
ðmg mg1 Þ
DPersistent DW
Phloem Water Delivery
ðmg mg1 Þ
Total DW Delivery
Phloem Water Delivery
ðmg h1 Þ or ðmm3 h1 Þ
hour
Minimum
Maximum
Comments
0.0093
0.012
Table 1, Line 1
0.85
1.48
Table 1, Line 3
1/0.55
1/0.51
Wheat, Table 2
2.1
9.8
ð0:0093Þð0:85Þð2:1Þ
¼ 0:030
0:55
ð0:012Þð1:48Þð9:8Þ
¼ 0:341
0:51
Sucrose, amino acids, K+, Table 2
Eqn 1
Table 4. Summary of internal and external rates of water delivery for root growthA
Line no.
1
2
3
4
5
Experiment
B
Comparison: growth in air versus soil in intact roots
Comparison: growth in soil before and after excisionC
Growth calculated from average water delivery by phloem, wheatD
Maximum from theoretical calculation, maizeE
Maximum, dilution of inorganic ions from phloem, barleyF
% Internal
% External
45
26
64
81
78
55
74
36
19
22
A
% internal + % external = 100%.
This study, intact root rates in Fig. 4; % internal = 100 (rate in air intact/rate in soil intact).
C
This study, rates in soil before and after excision in Fig. 4; % internal = 100 – 100 (rate in soil excised/rate in soil intact).
D
This study Table 1, % internal = 100 (calculated average rate of water delivery from phloem/actual rate of total water delivery
to root).
E
Bret-Harte and Silk (1994).
F
Pritchard et al. (2000).
B
internal sources always operated because the phloem
delivered solute essential for building new root, and water
delivered with the solute would contribute to the water supply.
That each supply was important was demonstrated by a control
test that excised the roots growing in saturated air. Without an
internal water supply from the plant or an external supply from the
Path of water for root growth
soil, growth was statistically indistinguishable from zero (e.g.
Fig. 6b).
Water from internal and external sources
By using time-lapse imaging, we were able to make precise
measurements in situ with intact, undisturbed roots in soil and
in air, before and after excision. The imaging showed
unequivocally that all seminal axes of wheat can grow in
saturated air, at ~45% of the rate in soil. The roots were
unlikely to take up water directly from the air because the root
tissues were slightly warmer than the air due to metabolic heat
(also seen in thermocouple psychrometry, where the heat must be
measured and controlled (Boyer 1995)). With the roots slightly
warmer than the air, the vapour pressure of root water could be
greater than in the air. It is likely that the bulk air was at or very
close to 100% saturation because condensate was observed on
the transparency face close to the soil-mesh interface in the
mornings when the soil temperature was likely cooler than the
air in the compartment. Because roots did not change in length
significantly when excised in this air, we concluded that the
roots neither lost nor gained water from the saturated air. Roots
leaving the soil lost direct contact with nutrients that may also act
as signals to induce changes in gene expression. However when
tissues are deprived of mineral nutrients, genes require at least
10–30 min to change expression (Wang et al. 2002; Schachtman
and Shin 2007). Additional time is needed for these changes to
be expressed in the phenotype.
If factors other than water supply are constant between the
soil and air compartments, then it is reasonable to conclude that
the ratio of air to soil rates is an estimate of the contribution from
internal sources (Table 4, Line 1). However, we are aware of the
decrease of root elongation rate with soil resistance (Passioura
1991). We emphasise that in the present study the penetration
resistance was low, and the density of the bulk soil and the
interface between the soil and the transparency were very low
compared with those bulk densities known to affect root growth
in controlled experiments (Greacen and Oh 1972; Bengough and
Mullins 1991) and in the field (Brady 1990). Certainly, our results
show clearly that roots can extend in saturated air for days by
relying on internal water sources.
When the internal water supply was removed by excising the
root tips in soil or air, growth responded instantaneously. We
attribute this result to a need for a continuing supply of the
incompressible liquid as shown by Boyer et al. (1985) and
Matyssek et al. (1988, 1991) in hypocotyls and stems.
Excision also removed the carbohydrate supply but we assume
carbohydrates were not depleted for several minutes. This
assumption is supported by continued growth for ~30 min post
excision in soil, after which elongation generally diminished.
Excision also may cause artefacts due to touch, wounding, and
removal of hormones and other small signals travelling into the
root tips, causing changes in gene expression. Expression of
new gene products and subsequent growth rate modulation
would take longer than a half hour (Massa and Gilroy 2003;
McCormack et al. 2006). However, touch (mechanical
stimulation) does cause immediate effects on plant function.
Phloem transport is slowed within minutes of stimulation with
a vibrating probe (Jaeger et al. 1988). This short-term effect is
Functional Plant Biology
1113
not a problem for our interpretation, as cessation of phloem
transport in the excised root would halt the residual internal
water supply to the growth zone. The most serious challenge
to our interpretation of the excision experiments comes from
reports that mechanical stimulation rapidly induces growth
cessation in several species (reviewed by Coutand 2010).
Growth rate is slowed in tissue distant from the site of
mechanoperception (Coutand and Moulia 2000). The growth
inhibition often lasts for hours and begins on a time scale of
minutes after application of a short mechanical stimulus. This
phenomenon casts doubt on our interpretation that excised
roots grow slowly as a direct result of removal of the phloem
water supply. However, recent theories of the mechanism of
thigmomorphogenesis support our conclusion. First, it is
becoming evident that plants respond to strain (tissue
stretching) rather than the stress imposed during mechanical
stimulation (Coutand and Moulia 2000; Coutand et al. 2009).
In our excisions with sharp scissors, tissue strain was minimised;
also both excised and control roots were subject to some
thigmostimulation from soil. Thus, we expect growth effects
from the mechanical stimulus of excision are less than the
impact of the removal of phloem water.
For the roots in soil, a simple model of three outcomes seems
possible for these experiments. As shown in Fig. 7 for the first
30 min after excision, root growth is maintained at the same rate
after excision because all the water is from the external source in
the soil (Fig. 7a), or root growth ceases because all water is from
internal source (Fig. 7b), or a portion of water is internally
supplied while another portion is externally supplied (the
hypothesis of this study, Fig. 7c). Figure 7c is what was
observed and is the origin of the 26% estimate for growth
originating from internal sources and 74% supplied externally
by the soil (Table 4, Line 2).
Water from phloem
Water moving into the non-vascularised root tip may travel
towards the tip on cell surfaces, in mature xylem behind the
tip, or in the mature phloem. Surface flows might occur along
longitudinal grooves between cells, outside of the root or within
the root (J. Passioura, pers. comm.). Outer groove flow was
unlikely because the soil water above the mesh was too dry to
provide free water, and intercellular spaces between epidermal
cells in soil-grown roots are filled with a gel biofilm of polymers
of root and microbial origin (McCully 1999; Watt et al. 2006).
Water in intercellular spaces in roots (Watt et al. 1996) could be
drawn to expanding tip cells to provide some water for extension
if longitudinal spaces are continuous and free of polymers.
Nevertheless, because the surface water was under tension in
the transpiring plants, as shown by Westgate and Boyer (1984,
1985) for maize roots, most surface water would be absorbed and,
thus, was unlikely to contribute significantly. By contrast, xylem
supplies the water for transpiration far in excess of the
requirement for growth. Functional xylem develops basal to
the growth zone and clearly would be capable of moving to
the base of the growth zone (but not into the growth zone) all the
water required by the tip.
But how much of the internal water could be supplied by
the phloem, which passes through the elongating region? On
1114
Functional Plant Biology
J. S. Boyer et al.
by Pritchard et al. (2000) for elongating roots of wheat,
calculated from dilution of phloem contents upon delivery to
the growing cells (Table 4, Line 5). Part of the difference may be
caused by the assumption of equal densities for water and phloem
solution in the calculation. If the phloem density was heavier than
water (e.g. 386 mM sucrose solution has a density of 1050 g L–1),
a given weight of sucrose delivered to the root tip would carry a
larger volume of water than calculated here, and the phloem
would be judged to be capable of delivering more than 64% of the
total for root growth. In the absence of density data for the phloem
solution, however, the current estimate is conservative and seems
acceptable. The literature gives wide variation in phloem
concentrations obtained from aphid stylets probably because
different environments, species, or organs could have
influenced the estimates. For example, reported concentrations
tended to be lower at night than during the day. But these
conditions were included in the averages, which should then
be reasonably robust.
In addition to the water and solute fluxes already mentioned,
carbon released from root tips as exudates would carry water, and
there may be water efflux to the rhizosphere from the root tips
(Passioura 2002; Tumlinson et al. 2008). There also is recirculation of water and solute typically for expansion of root
hairs or branch roots within the parent root (Lambers et al. 1982).
Also, inorganic ions delivered in soil water can be re-circulated to
the root tips in the phloem, with K+ predominating (details by
Hayashi and Chino (1986)). The experimental data included
effects of these losses and re-circulation, because persistent
DWs and FWs were net values indicating the difference
between influx and efflux to the root tip, and the K+ in the
phloem was included in the analysis.
(a)
0
(b)
Change in length
100
(c)
26
Significance for root architecture
Time
Fig. 7. Model of root elongation before and after cutting at arrow. (a) Cut
makes no difference to elongation. (b) Cut eliminates elongation. (c) Cut
causes slow elongation. In (a), no water needs to be supplied internally in order
for elongation to continue (0), but in (b) all the water for elongation is supplied
internally (100) and in (c) 26% of the water for elongation is supplied
internally (26).
average, calculations indicated the phloem could supply as
much as 64% of the total water for growth (Table 4, Line 3),
but the amounts in the experimental measurements were 45 and
26%, respectively. Therefore, the phloem could be the source of
all the internal water for growth.
Although 64% is a large amount, it is somewhat below
the estimates by Bret-Harte and Silk (1994) for elongating
primary roots of maize, calculated from deposition rates of
carbohydrate with accompanying water (Table 4, Line 4), and
The relative contributions of the external and phloem pathways
probably depend on the environment. Phloem water might be
entirely responsible for root growth across air gaps in soil or in
open air (nodal, prop roots of maize), or when surrounding soil is
dry. This growth would depend on water delivery from distant
roots in wet soil. There is evidence that some of this water can leak
out through the roots into the surrounding soil, creating ‘hydraulic
lift’ that enables the roots to re-hydrate dry soil mostly at night
followed by re-uptake the following day (Richards and Caldwell
1987; Kramer and Boyer 1995). Depending on the distribution of
water in the soil profile, it may be the deep roots that lift this water
to surface roots, or the converse. The growth in dry soil, although
slow, determines root system architecture (Kirkegaard et al.
2007) and is important for models such as the one by
Clausnitzer and Hopmans (1994) or for genetic efforts to
improve root systems with specific gene targets (De Dorlodot
et al. 2007).
With this insight it becomes clear that morphologically thinner
roots could distribute water for extension more effectively than
thicker roots, although recent modelling of the contribution of the
phloem water source to extension indicated that root thinning had
only a small impact on the ability of roots to sustain growth on
phloem water (Wiegers et al. 2009). The study by Wiegers et al.
(2009) also indicated that differentiation of phloem poles closer to
the root tip helps sustain root growth on phloem water sources.
Path of water for root growth
Indeed we observed variation in the growth zone and root hair
differentiation in roots growing in air for a few days (Fig. 3).
Phloem differentiation probably also varied relative to the tip (e.g.
see fig. 7 in Watt and Evans 1999).
In conclusion, roots can grow in air by relying solely on water
from internal sources. The phloem is capable of supplying all of
this water. This indicates that water absorbed in one region of soil
can feed root growth in another, thus, allowing roots to ‘jump’ soil
gaps and extend into dry soil, influencing the overall architecture
of the root system. Nevertheless, root growth is most rapid if
internal water sources are supplemented by water from the
immediate surroundings.
Acknowledgements
We are grateful to the Grains Research and Development Corporation
(GRDC) and to the CSIRO McMaster Foundation for providing funds for
this research, and the visits of WK Silk and JS Boyer to CSIRO, Canberra. We
thank Mike Hauptmann of the CSIRO Scientific Instrumentation Group for
building the time-lapse frame and the rhizo boxes and John Passioura for
calculating the air-filled porosity of the soil.
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Manuscript received 11 May 2010, accepted 10 September 2010
http://www.publish.csiro.au/journals/fpb