| The art of growing plants under (close to) optimal conditions in order to gauge their potential growth rate, or under experimentally manipulated suboptimal conditions to study the effect of the environment is not simple. Various potential pitfalls may await the experimentator. A number of recommendations to avoid these pitfalls have been given in an electronic appendix to Hannemann et al. (2009) Plant Cell & Environ 32: 1185-2000. To make these more easily available to the scientific community, these recommendations are given here below as well. |
| T1 | Light quantity |
| T2 | Light quality |
| T3 | Air humidity |
| T4 | Elevated CO2 |
| T5 | Temperature |
| T6 | Nutrients |
| T7 | Water |
| T8 | References |
| Background: |
|
For the wide range of wavelengths intercepted by leaves of plants, the most relevant for
photosynthesis is radiation in the wavelength between 400 and 700 nm. This is
photosynthetically active radiation (PAR) that is able to excite the photosystems to provide
the energy for carbon fixation. In relation to photosynthesis and C-budgets, light availability
is best described by the amount of quanta in the 400-700 nm range that fall on a horizontal
plane for a given amount of time. Under natural conditions this may vary from 0 at night till
2200 µmol m-2 s-1 in full sunlight at low latitudes. Defining the radiation energy in terms of
the amount of energy m-2 s-1 is not adequate for photosynthesis studies, as blue light has a ca.
70% higher energy per quantum than red light, yet is equally useful for the plant in terms of
photosynthesis. However, radiation energy in the whole wavelength range is a more relevant
descriptor in analyses of the heat balance is of an organ.
The light intensity that saturates photosynthesis strongly depends on the light availability the plants were grown at, as well as other environmental conditions like temperature and nutrient supply. The relative growth rate (RGR) of the plant is not as much driven by the light intensity at a particular moment, but more by the total amount of quanta integrated over the day (Poorter & Van der Werf, 1998; Granier & Tardieu 1998). Daily quanta input in nature varies from approximately 1 mol quanta m-2 day-1 in deep shade, to more than 50 on clear days in summer and in (sub)tropical areas. It is important to note that irradiation outside the range of PAR can have a major impact on plants; in addition to signalling (see next section) this includes the stressful impact of UV, and the impact of irradiation on the heat balance. For this reason, the full spectrum should be measured; there can be large differences in the intensity and distribution of irradiance in the UV and infrared regions of the spectrum, even at the same PAR. |
| Controlled environments: |
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In growth cabinets, light is provided by fluorescent tubes, high intensity discharge (HID)
lamps or, in a recent development, light emitting diodes (LED’s). Several issues are important
to consider. First, the question of how equal is the light intensity distributed over the growth
room needs to be considered. This is particularly the case when point sources such as HID
lamps are used, where variation of more than 50% may occur locally. Densely spaced
fluorescent tubes or LED’s provide generally better distributed lights; however, even when
such lamps are used, light intensity can be 20 – 50% less close to the walls of a growth room.
This can be reduced by using reflective mirrors on walls (a white coating is less effective).
Circulating plants throughout the growth room during the experiment is a solution to mitigate
environmental heterogeneity. Light intensity generally increases the closer to the light sources
(i.e. with increasing plant height) and, horizontally, towards the centre of the cabinet, but
variability increases as well. Hence, heterogeneity increases both horizontally and vertically
for plants with tall stems. Vertical gradients are aggravated when plants are not spaced apart
and shade each other. Finally, light output generally also decreases with the age of the lamps.
Measuring at least two times during the experiment or at least once a month in longer
experiments over 10-20 places in the growth room is recommended to characterize light
quantity. Growth is generally limited by light levels lower than 20 mol m-2 day-1. When light
levels become low for the species of interest, one can consider increasing the photoperiod.
In glasshouses, frames and lamps hanging over the plants take away circa 30% of the incoming radiation. Still, radiation can be very high in clear summer days (> 1100 µmol m-2 s- 1), and very low in winter or with cloudy weather (< 100 µmol m-2 s-1). Additional lamps can help in the winter season to extend the light period to a more or less fixed duration, and also add extra quanta for plant growth. As mentioned before, peak irradiance is a poor characterization of a glasshouse light environment. It should at least be supplemented with daily quantum input averaged (e.g. over the experimental period). |
| Light quantity as a treatment: |
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To use light quantity as a treatment, plants are generally shaded by nettings. Make sure
that the netting intercepts all wavelengths to an equal extent; otherwise light quality will also
be affected. Strong compartmentation of the growth room, for example by shade cloth may
block the air stream, causing secondary effects. Other secondary effects will almost inevitably
occur: a high light environment may warm up the leaves, but also increases the transpiration
rate of the leaves and speeds up growth. This implies that those plants have greater demands
for water and nutrients, and therefore run into water and nutrient limitations earlier during
growth than low-light grown plants. A reddish shade due to anthocyanin accumulation, first
visible at the lower side of the leaves is an indicator thereof.
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| Further reading: |
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| Background: |
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The PAR part of the light spectrum (see above) represents roughly half the irradiance in
normal sunlight. The other half is infrared (700-3000 nm) and ultra violet (UV; 10-400 nm).
To this comes the long-wave radiation from surrounding objects. Plants modify this spectrum
by selectively absorbing PAR and UV and part of the infra-red. Hence, light transmitted and
reflected by green tissue is depleted in red (660 nm; R) relative to far-red (730 nm, FR)
(Smith 1984). Phytochrome, a family of photoreceptors with a common chromophore,
perceives the R to FR ratio of light and thus the proximity of neighbours, leading to a
multitude of photomorphogenetic responses collectively called “shade avoidance”. There are
also other photoreceptors active in plants, cryptochrome and phototropin, that show
absorption in the blue region of the spectrum (Jiao et al. 2007). These are not involved in the
perception of the daylight spectrum, but their absorption characteristics can make their action
relevant in growth chambers with artificial lighting. The spectral differences with daylight can
make photomorphogenesis in growth chambers substantially different from that in daylight. It
depends on the research question to what extent correction is necessary.
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| Controlled environments: |
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Most lamps used in growth chambers have not been designed for imitating the daylight
spectrum. Some are specifically designed for supplemental lighting in greenhouses, where
their spectrum is mixed with that of daylight. The most important differences in the emission
spectra of these lamps compared to daylight are that their short-wave emission is largely in
the PAR region, resulting in a much higher R to FR ratio. Their emission in the PAR region
may furthermore be depleted in red and/or blue, and the presence of emission peaks at
specific wavelengths may be relevant. HID lamps in particular have a very high emission of
long-wave infra-red as a result of their high temperatures
Growth in far-red depleted light results in reduced extension growth, and may alter day length responses. Correction is mostly done by adding incandescent lamps. A disadvantage is their low efficiency and thus the need for a large wattage to raise the R to FR ratio to daylight values; although they have the added advantage of correcting a possible depletion in R. The excessive thermal radiation of HID and incandescent lamps cause a heat load on substrate and plants, particularly shoot tips, which can raise their temperature substantially above the controlled air temperature at the low turbulence in most growth chambers (Cummings et al. 2007). The problem is reduced by mounting the lamps in compartments separated by an infrared absorbing window. This does, however, not solve the problem sufficiently, since reradiation from this window can still result in elevated surface temperatures. Evidently, the magnitude depends on irradiance levels. Recently, far-red emitting diodes have been used for the correction and manipulation of the R to FR ratio in growth chambers (Pierik et al. 2005). Such LED’s have the potential advantage of a high efficiency and wavelength precision. High output white LED’s are becoming available as well. They have not been used in growth chambers yet, but have an evident potential for that purpose. LED’s have a higher efficiency and a reduced thermal radiation per unit of PAR. Their emission spectra can be adjusted and/or a mixture of LED’s with different emission spectra can be used for plant growth purposes. |
| Light quality as a treatment: |
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When studying photomorphogenesis, spectral quality of the light is of particular concern.
Its control should be more rigorous than for general plant growth purposes, and it must be
possible to manipulate it experimentally. Spectral quality can be manipulated by filters that
absorb in specific spectral regions, or by supplemental lighting. Typically, irradiance of PAR
should be kept constant between treatments, thus avoiding an effect on photosynthetic rate.
Filtering in the PAR spectral region should then be compensated by increasing irradiance.
This manner of spectral manipulation has an additional potential problem created by the
reduced ventilation and turbulence caused by the filter sheets. When the R to FR ratio is the
experimental variable, PAR irradiance is most easily kept constant by manipulating irradiance
in the far-red. Filtered incandescent lamps were used for that purpose in the past, but a high
wattage is required to decrease the R to FR ratio, resulting in a thermal radiation problem that
should for instance be corrected by a water filter. The far-red emitting diodes mentioned
above are now the preferred choice. Also other parts of the spectrum can be manipulated by
LED’s.
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| Further reading: |
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| Background: |
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The amount of water transpired by a plant depends on air humidity, leaf temperature, the
quantity of light intercepted by the plant, stomatal conductance and leaf area. Air humidity is
characterised by the difference between the partial pressures of water vapour in the air at
saturation (e.g. fog) and in the air near the leaves. This difference is termed vapour pressure
deficit (VPDair, kPa), which varies from near 0 in saturated air to 4 kPa in a very dry air. The
effect of light operates via an increase in leaf temperature, which increases the amount of
water held in the air of the intercellular spaces. The driving force of transpiration can
therefore be characterised as the difference in water vapour pressure between the intercellular
spaces (saturated and at leaf temperature) and the open air around the leaf (at air temperature),
termed VPDla. A measure more commonly used is the Relative Humidity (RH) which is the
ratio, instead of the difference, between the partial pressures of water vapours at saturation
and in the air. However, RH by itself is not an accurate description for evaporative demand,
because the evaporative demand increases with temperature at a given RH (e.g. more than
doubles from 16 to 30°C at a RH of 70%). VPDair and VPDla can be easily calculated from
relative humidity and air (respectively leaf) temperature. A good measurement device
depends on both accurate gauging of temperature and water in the air, which is not trivial. It is
essential that the RH sensor is shaded, and if possible ventilated. High quality growth
chambers offer this possibility. Many of the cheaper RH meters will give merely an
approximation if they are not shaded and ventilated, because they provide information about
the sensor temperature instead of the air temperature.
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| Controlled environments: |
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Humidity in a growth cabinet generally increases with the amount of evapotranspiration
of the plants and pots, and thus depends on size and number of plants. Part of the transpired
water will disappear with the vented air. However, most of the air in the growth room is
circulated, and especially if the radiation load in the growth room is high, strong cooling has
to be applied, which acts as a cold finger and extracts most of the air water. In that case an
extra evaporation step in the air treatment is necessary to increase the humidity again. Note
that air is drier during winter than summer, so low humidities are more easily obtained in
winter. Too low humidity (VPDair >3 kPa) will cause stomatal closure of the leaves in some
species, too high humidity (VPDair <1 kPa) will inhibit transpiration and may increase the risk
of diseases. It can also cause serious problems to equipment in the growth chamber.
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| Air humidity as a treatment: |
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Air humidity (VPD) affects both transpiration rate and leaf growth rate in most species.
In well watered conditions, it can therefore be an experimental way to manipulate
transpiration, or to affect leaf growth rate without change in soil water status. In addition, it is
a component of the water deficit, which is defined by both soil offer and evaporative demand
(see below). An experiment with varying VPD usually needs a chamber with a strong air
desiccation, or can otherwise be carried out in winter by using open air. Conversely, it should
be remembered that any experiment which affects air temperature will also affect
transpiration, via VPD. Precise controls are therefore necessary to avoid confusions of effects
between high temperature and high evaporative demand. Carrying the temperature
comparisons during the dark period is a way to avoid this problem. Automatically controlling
VPD is another solution.
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| Further reading: |
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| Background: |
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The CO2 concentration in the air is steadily rising and is currently circa 380 µl.l-1. There
is variation over seasons, especially in the northern hemisphere, with values during summer
being circa 5-20 µl.l-1 lower than during winter (Keeling et al. 1996). In some locations, there
can be large short-term changes; for example, CO2 concentration often peaks during periods
of heavy traffic in locations close to roads, or in cities on winter days with temperature
inversion. The CO2 concentration is limiting photosynthesis in the short-term, and thus initial
increases of 20-50 % are found when the CO2 concentration is doubled. Although growth over
the longer term is less responsive to increases in CO2 concentration above 380 µl.l-1,
individually-grown plants are, nevertheless, on average 30 – 60 % heavier at elevated CO2
concentrations, with part of the weight increase due to starch accumulation (Poorter & Navas
2003). Stronger effects on growth are likely to occur when plants are grown at CO2
concentrations below 330 µl.l-1.
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| Controlled environments: |
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In most growth cabinets CO2 concentrations are not controlled, and even not measured,
making it the most ill-defined condition of the aerial environment. If a large amount of
biomass is present in the growth room, CO2 concentrations may go up during the night, and
down during the day. The extent to which this happens will partly depend on how ‘leaky’ the
growth room is towards its outer environment. In buildings, the CO2 concentration is
generally 50 - 200 µl.l-1 higher than outside, due mainly to the human respiratory activity.
With someone walking into a growth room, respiration will locally increase dramatically and
can go up to 1200 µl.l-1 or higher. Consequently, working for a few minutes in a growth room
to sample your plants leads to an unwanted and uncontrolled CO2 enrichment experiment! If
phenotypic variables are being studied that respond rapidly (e.g. stomatal conductance and
photosynthesis), it may therefore be necessary to exhale into piping that leads the CO2 into an
absorber or outside the chamber.
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| CO2 concentration as a treatment: |
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CO2 enrichment studies can be carried out relatively easily by measuring the CO2
concentration, and adding CO2 by operating a valve on a CO2 cylinder. It is important to make
sure that the CO2 source of your manufacturer is not biological, as otherwise ethylene may be
present, which is a strong plant hormone. CO2 control is easiest during the stage that there is
considerable biomass in the growth room, which causes a strong drawdown. CO2 control is
less easy in the beginning of an experiment, especially at ambient concentrations, because
concentrations can be too high. When sub-ambient concentrations are of interest, molecular
sieves that catch the CO2 in the ingoing air stream are necessary.
High CO2 concentrations close the stomata, and so water availability is generally not an issue in such experiments. However, there is a strong interaction of plant growth stimulation due to CO2 with nutrient availability (Geiger et al., 1999; Poorter & Navas 2003), and larger and fast-growing plants require more nutrients than smaller slow-growing plants. Care should therefore be taken that nutrient supply in a CO2 enrichment experiment is even better controlled than usual. |
| Further reading: |
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| Background: |
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In nature, plants frequently experience considerable spatial and/or temporal temperature
variability, with profound consequences for their physiology, development and overall
growth. Temperatures decline by approximately 1°C for each 2° increase in latitude or 100 m
increase in altitude. Temperatures of flowers, leaves, stems and roots may differ substantially,
with leaf temperatures often being up to several degrees higher or lower than that of the
surrounding air depending on radiation, air movement and stomatal conductance. Large daily
and seasonal variations in temperature are also common in many habitats. Finally,
temperatures are changing as a result of global climate change, with night-time temperatures
raising more than daytime temperatures.
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| Controlled environments: |
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As mentioned above, radiation by lamps can result in leaf temperatures being
substantially higher (1-6°C) than that of the surrounding air. To minimize heterogeneity of
temperature from one side of the cabinet to another, air flow should be vertical. Vertical air
flow also helps reduce, but not eliminate, radiative heating of leaves at the top of canopy. In
older cabinets, temperatures are often controlled using on-off system cycles, with the result
that temperatures could vary by as much as 5°C above/below the set point temperature. In
modern cabinets, this variability is reduced by matching the rate of heating/cooling to the
extent to which the temperature has deviated from a set point temperature. When growing
plants at low temperature, ice needs to be removed from the cooling coils at regular intervals
using a defrosting cycle; in some cabinets, defrosting may result in a transient increase in air
temperature (e.g. usually once a day when lamps are off). As maintaining low temperatures
places a heavy load on refrigerating systems, it is advisable to select cabinets designed to
maintain temperatures several degrees below the set point temperature, rather than expect
reliable operation at the engineering limits of the cabinet.
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| Temperature as a treatment: |
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When using temperature as a treatment, consideration needs to be given to whether the
aim is to achieve a set point leaf and/or air temperature. In the event that leaf temperature is of
interest, then adjustments to air temperature and air flow rates may be necessary. In
treatments where cabinet day- and night-time temperatures differ, consideration may need to
be given to the fact that the temperature of the rooting media will lag behind that of the
leaves. Many plant species exhibit maximal growth rates when air temperatures are 3-10°C
higher during the day than at night, and thus differential day/night temperatures are often
used. Where it is necessary to control shoot and root temperatures independently, set point
root temperatures should be maintained separately (e.g. by inserting temperature coils into
solid rooting media). Be also aware that increasing temperature results in an increase in
evaporative demand.
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| Further reading: |
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| Background: |
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Plants require a wide range of macro- and micronutrients for their growth. De novo
uptake by the roots is generally only required for the construction of new biomass. This
implies that nutrient demand of a plant follows growth (Ingestad, 1982), and as growth of
most plants studied in controlled environments is close to exponential, it implies that nutrient
demand increases exponentially as well! In the aerial environment, factors are relatively easily
controlled by keeping their values constant (such as temperature and humidity) or resupplying
additional resources (such as CO2 due to diffusion). However, nutrients are
different and if not well controlled, plants that started of with a luxurious supply of nutrients
during the seedling stage will hit stronger forms of nutrient stress later during growth. This is
especially the case if the plants are supplied with nutrients in a form that is easily leached out,
or are grown in small pots. Another potentially major source of error can arise in experiments
that use a treatment (e.g. changes in the amount of irradiance or temperature, or comparison
of different genotypes) to alter the rate of growth. Unless the nutrient supply is luxurious, the
differing rate of growth can result in secondary changes in the nutrient status of the plant, and
the effects of this may modify or even mask the effects of the original treatment. For example,
elevated CO2 concentrations lead to only a marginal increase in growth but a marked decrease
in organic N if an N supply is used that is just about sufficient for growth in ambient CO2,
whereas elevated CO2 concentrations increase growth and have little or no effect on the
organic N content when plants are supplied with supra-optimal high N (Geiger et al., 1999).
The levels of nutrients such as N, P, K, S or Fe should be regularly determined in potting soils that are used in routine. However, the nutrient availability is not always easily measured. For example, phosphorus may be found to be present in the soil in rather large amounts, but may be bound to the soil so strongly, that it is not available for plant growth. Furthermore, availability depends strongly on the volume of soil that is covered by roots. In nature, higher demands of larger plants are partially covered by large plants exploring larger soil volumes. However, plants in pots may quickly have their roots pot-bound. |
| Controlled environments: |
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Plants can be grown in a variety of substrates. In the simplest way plants are grown in
pots on a soil-like substrate. Generally, pot size is chosen relatively small, to accommodate
large amounts of plants in a generally limited amount of table space, but this increases
vulnerability of unwanted nutrient stress. Be aware that most potting mixes contain very
limited amount of nutrients, which implies that nutrients have to be added, slow-release
fertilizer being a good option. Nevertheless, even with the addition of slow-release fertilizer,
which provides nutrients more or less linearly, exponential demand of the plant is not met,
and such plants may develop nutrient stress over time.
There are several ways to provide a somewhat more ‘controlled’ nutrient supply. Their usefulness depends on the plant, and the biological question. Plants are frequently grown on nutrient agar. The levels of solutions typically supplied in nutrient agar, for example when grown on full strength MS (Murashige & Skoog, 1962), are extremely high, but can also be varied in a precise manner. Nutrient agar can also be used to study the effect of local heterogeneities in the nutrient supply (Zhang & Forde, 1998). This growth system does not allow control of nutrient supply as the plants become larger. It may also negatively affect plants that are sensitive to the low O2 supply in agar. Another option is to grow plants in sand, and supply a defined nutrient solution. The sand needs to be chosen to provide airspaces, while also retaining enough water and nutrient solution to avoid dehydration and support plant growth. Typically, this can be achieved by mixing sand with at least two different particle size ranges. Watering is necessary at frequent intervals, often every day or even more frequently when the plants are larger. The nutrient solution is applied by filling up the pot, allowing the nutrient to run through, and repeating this 2-3 times, in order to replace the original depleted solution. Rather than sand, other substrates like rockwool can be used. These growth systems can be further refined by using continuous or computer-controlled drip irrigation. Alternatively, plants can be grown in a hydroponics system, with roots hanging freely in an aerated solution. Good circulation is the key to avoid depletion zones around the roots or anaerobic patches, but should not be too vigorous, as it may hinder Fe uptake, especially for grasses. The pH of the solution may increase or decrease, depending on whether ammonium or nitrate is present in the solution, but should generally not deviate too much from pH 6. As a rough rule, the concentrations of nutrients that need to be supplied in a hydroponic system are 5-10 times lower than when plants are growing on sand. |
| Nutrient stress as a treatment: |
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Nutrient stress can be applied as an overall decrease of available nutrients, or with one
specific element in short supply. The latter requires plants to receive water in some form of
nutrient solution. Several issues need careful consideration in planning an experiment,
including: (i) whether it may be necessary to observe and / or harvest the roots (this precludes
the use of substrates that form an intimate physical complex with them); (ii) how to maintain
a plant in an intermediate state of nutrient supply for a period of time (rather than shifting it
rapidly from high to limiting nutrient supply); and, (iii) whether the response of root growth
and root architecture forms an integral part of the biological response that is being
investigated.
A hydroponic system is not necessarily the best medium to impose nutrient stress. The rate of nitrate uptake by a root, for example, is virtually constant over a range of 10 ìM to 10 mM. This implies that plants will experience a situation of plenty, until almost all nutrients are taken up, and then suddenly stress will hit the plants severely. A better way is to increase the amount of nutrients available to a plant in an exponential way. This method, initiated by Ingestad and co-workers (Ingestad 1982), provides a more constant level of nutrient availability relative to demand over time. Plants will grow with a growth rate that is determined by the exponential increase in nutrients, which should be lower than set by the maximum RGR of the genotype in the environment under consideration. Hydroponic systems also have the disadvantage that nutrients are continuously replenished at the root surface, and there is no ‘benefit’ from exploring a larger volume. i.e., it does not pay for a plant in such a system to allocate more of its biomass to roots to sequester more nutrients. To allow for this it would be better to daily supply nutrients to sand or another solid substrate, in which a higher allocation to roots implies that a larger volume is explored and more nutrients can be accessed. The impact of low nutrient stress is partly visible as lower growth, but also shows up in a more specific manner as changes in composition. In addition to lower levels of nutrients themselves, low N typically leads to a large decrease of the glutamine/glutamate ratio (Stitt & Krapp, 1999). Moreover, for most nutrients low supply induces strong shift in the allocation of biomass to the roots (Poorter & Nagel 2000). Low nutrient availability typically, decreases growth more than photosynthesis, resulting in starch accumulation. |
| Further reading: |
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| Background: |
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Water deficit occurs when the water flux which crosses the plant is insufficient to cope
with the evaporative demand (linked to light intensity, air VPD and leaf area). It is therefore a
transient imbalance between soil supply and air demand, which causes a partial dehydration
of leaves, and therefore a decrease in leaf water potential. The plant reduces transpiration by
stomatal closure, thereby decreasing the water flux through the plant. This protects leaves
against dehydration, but also leads to an inhibition of photosynthesis and an increase of the
leaf temperature. The variables which are involved in water deficit are, therefore, multiple
(leaf water potential or relative water content, stomatal conductance, leaf temperature...).
They can vary rapidly in fluctuating conditions (e.g. four-fold variations in some minutes
when light varies), and control each other by multiple feedbacks. Hence, an isolated
measurement of plant water status is almost meaningless without an environmental context.
One can either study time courses of each variable, which is only possible in experiments with
special designs, or fix most environmental conditions during measurements. In particular, leaf
water potential measured at the end of the night (predawn leaf water potential) gives a good
indication of the soil water status as sensed by the plant because, in the absence of
transpiration, soil and leaf water potentials are theoretically equal. In nature, soil water
potential ranges from near-zero in saturated soil to -100 MPa in a fully air-dried soil, but
plants of most species can only extract water in the range from 0 to -2 MPa. Stomatal control
prevents leaves (and roots) from drying below this value, which is therefore a biological
threshold for water extraction. This corresponds to ca 8% gravimetric humidity in a sandy
soil, 12% in a loam, 18% in a clay and up to 90% in a potting compost.
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| Controlled environments: |
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The necessity to water plants strongly depends on plant size, potting volume and
substrate. Generally, it is better to water plants frequently. Especially in the glasshouse at high
radiation, plants may experience mild water stress within a day if pots are small. The best
method for ensuring equal water status to all plants consists of weighing the pots and adding
water until a pre-defined target weight is reached corresponding to the target soil water
content. Spraying plants may not result in a very equal distribution of water. Sub-irrigation
systems, where a small layer of water is mechanically applied around the pots for a limited
amount of time ensures a much better supply, also during weekends. However, make sure that
highly mobile nutrients like nitrate are not washed away in such a system and the substrate is
suitable for capillary water uptake. Plants that receive water from the top may be supplied
with saucers to avoid leakage of too many nutrients. Plants grown in hydroponics obviously
require less attention in this respect.
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| Water stress as a treatment: |
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Water logging is a treatment where the water level is at the soil surface. Complete
submergence is achieved by placing the whole plant under water. The latter treatment will
also decrease light intensity, especially as algae may start to grow. Replacement of the water
is then necessary. Water flow may critically affect the plant’s performance.
In its simplest form, water deficit can be applied by withholding irrigation. From an
ecophysiological point, this is meaningless because the actual deficit experienced by the plant
depends on the amount of water taken up by plants during the considered days, which
depends on evaporative demand and leaf area. This point is crucial when genotypes are
compared: genotypes with large leaf area experience a more severe deficit than those with a
small leaf area. The latter are, therefore, mechanically ‘drought tolerant’, and the underlying
genes may be incorrectly interpreted as such.
Water deficit should be defined by both soil water status and evaporative demand (see above). Soil water status can be measured either via predawn leaf water potential or deduced from soil water content using special measuring devices. Soil water content per se does not give an indication because a given content (e.g. 20 g.g-1) can correspond to high water availability in a sandy soil and to a very stressing humidity in peat. The minimum indication for characterising water deficit should be (i) soil water content at sampling time, (ii) type of soil, (iii) evaporative demand at sampling time. Sampling at the end of night is usually preferred because it is reproducible, which is not the case for the light period in most cases. In the complex situation presented above, the choice of a method for imposing water deficit will depend on the objective of the experiment, as all methods have drawbacks. Organ dehydration with a warm air flux may be used for very short term experiments, provided that the corresponding organ water potential is measured. Dehydrations obtained by this method are often too severe in comparison to natural situations. Osmolytes such as PEG or mannitol can be used for short term experiments (usually not more than 1-2 d) in hydroponics. They impose a constant and known water potential to roots, but may cause toxicities to the plant. For experiments in soil, two strategies can be adopted. (i) If the measuring capacity of the research group is limited, it may be preferred to impose a constant evaporative demand in a growth chamber and to maintain soil water content constant by daily irrigation after weighing pots. (ii) Fluctuating evaporative demand and naturally declining soil water content are closer to natural situations, but need a more careful design of the experiment and frequent measurements of plant, air and soil variables. Both strategies can be used in phenotyping platforms for analysing hundreds of plants (e.g. Granier et al. 2006, Sadok et al 2007). |
| Further reading: |
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