Skip to main content
Plant Physiology logoLink to Plant Physiology
. 2010 Nov 12;155(1):108–116. doi: 10.1104/pp.110.165472

Leaf Functional Anatomy in Relation to Photosynthesis1

Ichiro Terashima 1,*, Yuko T Hanba 1, Danny Tholen 1, Ülo Niinemets 1
PMCID: PMC3075775  PMID: 21075960

Rubisco is a large enzyme with a molecular mass of approximately 550 kD. The maximum rate of CO2 fixation (i.e. ribulose-1,5-bisphosphate [RuBP] carboxylation) at CO2 saturation is only 15 to 30 mol CO2 mol−1 Rubisco protein s−1 at 25°C. Affinity to CO2 is also low, and the Km, Kc, at 25°C in the absence of oxygen is comparable to the CO2 concentration in water equilibrated with air containing 39 Pa CO2 (approximately 390 μL L−1), 13 μm. Moreover, RuBP carboxylation is competitively inhibited by RuBP oxygenation, which is the primary step of the energy-wasting process, photorespiration. If the CO2 concentration in the chloroplast stroma is low, the carboxylation rate will decrease while the oxygenation rate will increase. Under such conditions, light energy and other resources, including nitrogen and water, are all wasted, eventually leading to a decrement of fitness of the plants. From these data, we may consider that structural features of the leaf contributing to the maintenance of the high CO2 concentration in the chloroplast stroma may have been selected during evolution.

In this Update, we focus on the key structural features that affect CO2 concentration in the chloroplast stroma. First, we analyze the conductance for CO2 diffusion from the substomatal cavity to the chloroplast stroma (mesophyll conductance [gm], also called internal conductance). Because the low gm limits photosynthesis, the mesophyll surface area exposed to the intercellular spaces (Smes, mesophyll surface area exposed to intercellular spaces per unit leaf area) should be maximized to increase the area for CO2 dissolution and the effective pathway for CO2 diffusion, and thereby photosynthesis. Second, we analyze the light environment within a leaf, because, for maximizing photosynthesis, light should be delivered to all the chloroplasts in the leaf, distributing along the cell walls. In relation to the light environment within a leaf, we also point out some technical problems in measuring photosynthetic parameters. Third, the movement and positioning of cellular organelles are discussed. Finally, we discuss the urgent need for ecologically relevant developmental and cell biological studies that clarify the mechanisms that are responsible for structural and cell biological features in nature.

THE NATURE OF MESOPHYLL DIFFUSION CONDUCTANCE

During photosynthesis, CO2 diffuses from ambient air through stomata to the intercellular spaces. Then, the CO2 dissolves in the cell wall water and diffuses across the cell wall, plasma membrane, cytosol, chloroplast envelope, and stroma to Rubisco (Fig. 1). CO2 concentration in the chloroplast can be estimated in several ways (Pons et al., 2009). One of the most reliable methods is the simultaneous measurement of gas exchange and stable carbon (C) isotope discrimination. This method relies on the discrimination against 13CO2 by Rubisco. In an open system, with unlimited supply of 12CO2 and 13CO2, Rubisco will preferentially fix the lighter molecule, 12CO2. In a closed system, Rubisco will eventually fix both 12CO2 and 13CO2 until no CO2 is left. The leaf is intermediate between these two extremes. Thus, the ratio 13CO2/12CO2 of CO2 fixed by a leaf is considerably greater than the ratio in the ambient air. Technically, the ratios of 13CO2 to 12CO2 in the air incoming to and outgoing from an assimilation chamber are measured with a mass spectrometer, a tunable laser diode absorption spectroscope, or a cavity ring-down spectroscope (Pons et al., 2009).

Figure 1.

Figure 1.

CO2 diffusion from air outside the leaf to the chloroplast stroma. C denotes CO2 concentration as follows: Ca, ambient air; Cs, substomatal cavity; Ci, bulk intercellular space; Cc, chloroplast stroma. The left panel shows a cross section of a leaf; the right panel shows a closeup of part of a chloroplast appressed against the plasma membrane; and the bottom panel shows mesophyll conductance per unit chloroplast surface area (gm/Sc). CO2 flux via route B in the right panel is negligible compared with that via route A because diffusion of inorganic C in the medium at pH 7.4 is very slow. Thus, virtually all the inorganic C is transported across the cell wall and cytosol facing chloroplasts. gm/Sc is the inverse of mesophyll resistance/Sc, which is expressed as an array of resistances. These resistances are divided into wall resistance (Rw) and cellular resistance (Rc). Rcytosol, Cytosol resistance; Renv, chloroplast envelope resistance; Rpm, plasma membrane resistance; Rstroma, stromal resistance. Resistances across the membranes are expressed as the parallel pathways through aquaporin (rpm,aq and renv,aq) and bulk membrane (rpm,bm and renv,bm). Redrawn from Terashima et al. (2006) with some modifications.

It has been established that the photosynthetic limitation by gm expressed on a leaf area basis is in the same order as that by stomatal conductance (conductance for CO2 diffusion from the air outside the leaf to the substomatal cavity; Terashima et al., 2006; Flexas et al., 2008; Evans et al., 2009; Niinemets et al., 2009a). The representative values for CO2 concentrations in the substomatal cavity (Cs), bulk intercellular space (Ci), and chloroplast stroma (Cc) in actively photosynthesizing leaves in ambient air containing CO2 at 39 Pa, expressed as ratios, are Cs/Ca = 0.60 to 0.85 (even as low as 0.12 with tightly closed stomata), Ci/Cs = 0.90 to 0.99, and Cc/Ci = 0.50 to 0.80. In this Update, we do not examine stomatal conductance (for review, see Evans and Loreto, 2000). The CO2 drawdown from the substomatal cavity to the bulk intercellular spaces is small, in particular in thin leaves with large fractions of intercellular air space and/or in amphistomatous leaves (for review, see Terashima et al., 2006).

Diffusion of Inorganic C in the Liquid Phase

CO2 in the intercellular spaces dissolves in the apoplastic water of the cell wall and diffuses across the cell membrane, cytosol, chloroplast membranes, and stroma to Rubisco. According to Henry’s law, CO2 concentration in the liquid phase, [CO2], is proportional to the partial pressure of CO2 in the adjacent gas phase. At 25°C, [CO2] in the water equilibrated with the air containing 39 Pa CO2 is 13 μm. [CO2] is not affected by pH, whereas [HCO3] increases with the increase in pH. Assuming that [HCO3] is in complete equilibrium with [CO2], the ratio of the concentrations can be calculated using the Henderson-Hasselbalch equation:

graphic file with name pp165472E01_LW.jpg

With pKa of 6.2, [HCO3]/[CO2] at pH 5 (cell wall) is 0.063, at pH 7.4 (cytosol) is 15.8, and at pH 8.0 (chloroplast stroma in the light) is 63.1. pKa for HCO3/CO32– is 10.25, so that [CO32–] is negligible under physiological conditions. pKa is affected by various factors including ionic strength and solute species (Stumm and Morgan, 1996). Here, we adopted the value for an ionic strength of 200 mm (Nobel, 2004). CO2 diffusion in the liquid phase is slower than that in the gas phase by 4 orders of magnitude: diffusion coefficient of CO2 in air and water (Dw) at 25°C are 1.56 × 10–5 and 1.7 × 10–9 m2 s–1, respectively. The diffusion coefficient of HCO3 in water is 56% of that for CO2 (Evans et al., 2009). Thus, we may express the diffusion coefficient for the total inorganic C as Dw × (1 + 0.56 κ), where κ is [HCO3]/[CO2]. If the activity of carbonic anhydrase is high enough, κ may approach the fully equilibrated value calculated by Equation 1.

Cell Wall Conductance

In the cytosol at pH 7.4, diffusion of inorganic C can be 9.85 (=1 + 0.56 × 15.8) times faster than that of CO2. Even when diffusion of both dissolved CO2 and HCO3 is considered, diffusion of inorganic C in the liquid phase is still very slow compared with that in the gas phase. Then, the most effective diffusion pathway through the cytosol is the shortest one, across the chloroplast envelopes and cell walls adjacent to the intercellular air space (compare routes A and B in the right panel in Fig. 1). Thus, the chloroplast surface area would be important. Actually, gm for the plants of the same functional group (annuals, broad-leaved deciduous trees, broad-leaved evergreen trees, etc) is roughly proportional to the cumulated surface area of chloroplasts facing intercellular spaces divided by leaf surface area (one side), Sc (Evans and Loreto, 2000;Terashima et al., 2006). The slope, ΔgmSc, however, differs among the functional types and is highest in annual herbs, intermediate in deciduous broad-leaved trees, and lowest in evergreen broad-leaved trees.

Given that gm is proportional to Sc, we can analyze gm/Sc (mol m−2 s−1) as the inverse of a series of resistances, analogous to an electrical circuit (Fig. 1). Let us divide these resistances into the wall resistance (Rw; in m2 s−1 mol−1) and cellular resistance (Rc; in m2 s−1 mol−1) and assume that Rw is proportional to cell wall thickness (δw; in m). In Figure 2, gm/Sc is plotted against the thickness of the mesophyll cell wall. The curve with the highest determination coefficient (r2 = 0.74) is:

graphic file with name pp165472E02_LW.jpg

Figure 2.

Figure 2.

Mesophyll conductance per unit chloroplast surface area (gm/Sc) plotted against the thickness of mesophyll cell wall (δw). Data are from Terashima et al. (2006) and Evans et al. (2009). An equation fitted to the data was Inline graphic, where Inline graphic is the intrinsic wall resistance (m s−1 mol−1) and only δw was variable. The curve with the highest determination coefficient (r2 = 0.74) is gm/Sc = 1/(3.36 × 108 × δw + 27.9).

When we assume that Rc is little different among the species and the cell wall thickness is 0.1 μm (typical for annual herbs), Rw and Rc are 33.6 and 27.9 m2 s−1 mol−1. Thus, the cell wall resistance is responsible for about half the total mesophyll resistance. The equation also indicates that, with the increase in cell wall thickness, gm/Sc decreases and the contribution of cell wall resistance to the total resistance increases. This probably explains why the slopes of ΔgmSc are lower for trees.

Based on Niinemets and Reichstein (2003), Nobel (2004), and Evans et al. (2009), wall conductance may be expressed as:

graphic file with name pp165472E03_LW.jpg

where p is the porosity (m3 m–3), Dc is the diffusion coefficient of CO2 in water (m2 s–1), α is the ratio of diffusion coefficient of HCO3 in water to Dc (=0.56), H is the Henry’s law constant in mol m–3 Pa–1, P is the atmospheric pressure in Pa, τ is tortuosity (m m–1), and δw is cell wall thickness in m. p expresses how much space in the cell wall is used for CO2 diffusion, while τ expresses how much the path is lengthened or tortured by the cell wall. H is defined as:

graphic file with name pp165472E04_LW.jpg

where Cl is the CO2 concentration in the liquid phase in mol m–3 and Cg is the CO2 concentration in the gas phase in mol CO2 mol–1. Thus, P × Cg denotes partial pressure of CO2 in Pa.

We may estimate an effective p/τ from the best-fit curve in Figure 2. Because pH in the apoplastic water is moderately acidic, let us assume that inorganic C in the cell wall is exclusively CO2. When Dc, H × P, and κ, are substituted with 1.7 × 10–9 m2 s–1, 33.9 mol m–3, and 0, respectively, p/τ is calculated to be 0.052 (i.e. much less than the values that have been assumed, such as 0.3; Nobel, 2004). When nitrate is abundant in the xylem sap and/or the soil is dry, the pH of the apoplast water could increase up to neutral levels (Jia and Davies, 2007). Then, diffusion of HCO3 should be considered. Because porosity and tortuosity would also change with pH due to the protonation or ionization of chemical groups in polysaccharides and proteins present in the cell wall, effects of pH on wall conductance should be studied.

Cytosol Conductance and Stromal Conductance

Cytosol conductance and stromal conductance can be also analyzed using Equation 3. The cytosol between the plasma membrane and the chloroplast envelope is usually very thin, typically 0.1 μm or less (Evans et al., 2009). Under steady-state conditions in the light, chloroplasts are anchored to the plasma membrane by actin filaments, which effectively minimize the cytosol thickness (for review, see Takagi et al., 2009). In the cytosol, p/τ approaches 1.0 and κ is around 9.85. Consequently, for the layer of cytosol with the same thickness as the cell wall, the cytosol conductance would be more than 100-fold greater than the wall conductance.

For the chloroplast stroma, typically 2 μm thick, p/τ may be much smaller than 1.0 because of abundant Rubisco, other proteins, and thylakoids. Effects of viscosity can also be included in the p/τ value. If the activity of carbonic anhydrase is high, 1 + 0.56κ will approach 36 at pH 8.0. Assuming a p/τ value of 0.5, then the conductance of 2-μm-thick stroma is 18 times that of a 0.1-μm-thick cell wall. The actual conductance is likely to be greater than this estimation because the effective thickness of the stroma could be roughly half if Rubisco molecules are distributed evenly throughout the stroma. On the other hand, if the p/τ value is much lower because of higher viscosity in the stroma, pH is more neutral in low light, and/or if the activity of carbonic anhydrase is not sufficiently high, then the stromal conductance can be smaller. Otherwise, these rough estimations indicate that both the cytosol conductance and the stromal conductance are much greater than the cell wall conductance.

Plasma Membrane Conductance and Chloroplast Envelope Conductance

Because the membrane is hydrophobic, the inorganic C that is transported across membranes (the plasma membrane and the inner membrane of the chloroplast envelope) is mainly CO2. For the plasma membrane, the majority of CO2 transported from the cell wall across the membrane is subject to conversion to HCO3 because pH in the cytoplasm is around 7.4. This conversion is catalyzed by carbonic anhydrases located in the plasma membrane and/or the cytosol. In the case of the chloroplast inner envelope membrane, conversions of HCO3 to CO2 and of CO2 to HCO3respectively, occur on the outer and inner sides of the membrane. Involvement of the carbonic anhydrase isozymes in different cellular components (Fabre et al., 2007) is crucial for an optimal transfer efficiency of CO2.

Another important topic related to the transport of CO2 across the membranes is the involvement of aquaporins. Changes in the aquaporin content per unit leaf area by genetic transformation techniques confirmed that gm changes in accordance with the abundance of aquaporin protein expressed (Hanba et al., 2004; Flexas et al., 2006; for review, see Katsuhara et al. 2008). Aquaporins may be involved in CO2 transfer in the plasma membrane as well as in the chloroplast envelope (Uehlein et al., 2008). It is necessary to further analyze mesophyll conductance in relation to the functions of aquaporins. Because aquaporin is not an appropriate term for proteins that transfer CO2, and because aquaporins that transfer CO2 would cooperate with carbonic anhydrases at the membrane-liquid phase interfaces, the term “cooporin” has been proposed (Terashima et al., 2006).

In summary, the magnitude of limitation of photosynthesis by gm is much greater than had been thought. CO2 diffusion across cell walls and membranes is identified to be the major limiting step. Because gm, in particular wall conductance, plays an important role in differentiating plant functional types according to leaf photosynthetic capacity (Terashima et al., 2006; Evans et al., 2009; Niinemets et al., 2009a), we suggest that this important topic requires high priority in future studies. Although we express the conductance for CO2 diffusion by a one-dimensional model, Cc in the leaf would differ considerably. We discuss this problem later.

Given that gm is an important limiting factor of photosynthesis, it is very problematic to analyze gas-exchange data assuming that Ci is equal to Cc (this assumes the infinite gm!). The spurious effects of this assumption have been repeatedly pointed out. In particular, estimation of kinetic parameters of Rubisco without taking gm into account should be avoided (for review, see Niinemets et al., 2009b).

WHY DOES LEAF THICKNESS INCREASE WITH LIGHT AVAILABILITY?

Leaves need a considerable amount of Rubisco per leaf area for photosynthesis, because Rubisco has a low catalytic rate. Given that gm is finite and not large, as analyzed above, Cc decreases with the increase in the concentration of Rubisco per unit Sc (Fig. 3). Because the affinity of Rubisco to CO2 is poor (Kc is large), with the decrease in Cc, the carboxylation rate decreases and the rate of energy-wasting photorespiration increases. Moreover, as a result of a lower total energy use by the carboxylation and oxygenation at the low Cc, chloroplasts are more liable to be overreduced and photoinhibited. Thus, Rubisco/Sc should be kept small to keep Cc at high levels. Figure 3 also shows that, for the similar Rubisco/Sc, Cc would be lower in tree species that have lower gm/Sc than in annual herbs (Evans et al., 2009; Niinemets et al., 2009a). Then, Rubisco/Sc may be lower in tree species to suppress the decrease in Cc. However, probably due to scarcity of the data, we cannot see this trend in Figure 3. Alternatively, the very low CO2 may have imposed selection pressure to Rubisco in tree species. Actually, the average specificity factor of Rubisco, Vc × Ko/Vo × Kc (where Vc and Vo are the maximum activities of carboxylation and oxygenation and Ko is the Km for oxygen of Rubisco), is highest in evergreen trees and lowest in annual herbs (Galmés et al., 2005; A. Makino, personal communication).

Figure 3.

Figure 3.

Cc plotted against Rubisco content per Sc. Theoretical lines are drawn based on the kinetic parameters of rice Rubisco at 25°C (Makino et al., 1985) assuming that Ci is 28 Pa. White and black circles indicate annual herbs and broad-leaved trees (deciduous and evergreen trees are included). Data are from Hanba et al. (1999), Evans and Loreto (2000), Miyazawa and Terashima (2001), Oguchi et al. (2003), Terashima et al. (2006), and D. Tholen (unpublished data). For the same gm/Sc, Cc in the chloroplast with a greater Rubisco/Sc is lower than that in thin chloroplast with a lower Rubisco/Sc. There may be some decline in Cc within a chloroplast. ΔC in the cartoon denotes CiCc.

To realize higher photosynthetic capacities on a leaf area basis (Pmax, maximum photosynthetic rate at light saturation), sun leaves should have more Rubisco per unit leaf area than shade leaves. If leaves have more Rubisco, they should also have greater Smes that are prerequisite for larger Sc. Because neither the cell diameter nor the cell density differs greatly between sun and shade leaves of the same species, a larger Smes can be attained by increasing the cell height. This may explain why sun leaves have to be thicker than shade leaves (Terashima et al., 2001).

The importance of the increasing Sc for increasing Pmax in shaded leaves on exposure to high growth light has been shown by a series of elegant studies (Oguchi et al., 2003, 2005, 2006). In shade leaves of Chenopodium album, part of the mesophyll surface area was unoccupied by chloroplasts and Sc/Smes was around 0.78. When these shade plants were exposed to high light, the Smes was unchanged but Sc increased. Sc/Smes attained around 0.96 by exposure for 2 weeks. Pmax increased proportionally with the increase in Sc (Oguchi et al., 2003). By cutting of a tall dominant tree in a deciduous broad-leaved forest, the saplings of deciduous tree species on the forest floor were suddenly exposed to high light. In Betula ermanii, Kalopanax pictus, Magnolia obovata, and Quercus crispula, Sc/Smes and Pmax increased, while Smes were unchanged, as in C. album. Interestingly, leaves of three Acer (maple) species showed elongation of mesophyll cells, leading to increases in Smes. Sc and Pmax also increased accordingly (Oguchi et al., 2005, 2006). The leaves of species such as Fagus crenata, which have neither mesophyll surfaces unoccupied by chloroplasts nor the ability to increase Smes, failed to increase Pmax (Oguchi et al., 2005). In the study with C. album, Rubisco/Sc showed surprisingly constant values around 0.3 μmol m–2 across different light environments (Oguchi et al., 2003).

Figure 4 illustrates several strategies for increasing mesophyll surface area. For example, Sage and Sage (2009) showed that the highly lobed cells (armed cells) in rice (Oryza sativa) can achieve a very high Smes and Sc. For other strategies to increase Smes and their possible advantages and disadvantages, see Terashima et al. (2001, 2006).

Figure 4.

Figure 4.

Various strategies to increase mesophyll surface areas. A, Cell elongation. B, Cell elongation accompanied by cell division. C, Decrease in cell size. D, Armed cells of grass species having lobes. For cells having the same cell walls of equal thickness, the tissue with smaller cells is mechanically tougher (Terashima et al., 2001). The leaves with larger cells would expand faster. On the other hand, these would inevitably be thicker to have enough Smes to accommodate chloroplasts, in which case the diffusion in the intercellular spaces becomes significant. Such leaves, mostly in annual herbs, tend to have stomata on both epidermises (Terashima et al., 2006). The acclimation of photosynthetic properties of chloroplasts to the local light environment may be more precise in cells in B than in those in A. In grass leaves, armed cells have large cell surface areas. In rice, almost all the cell surfaces facing the intercellular spaces are occupied by chloroplasts. Mitochondria are located in the capsule made by chloroplasts. For more insightful discussion for grass leaves, see Sage and Sage (2009).

LIGHT ABSORPTION BY THE LEAF

For efficient use of the light energy, light needs to be delivered to all the chloroplasts distributed along the cell surfaces throughout the leaf. Leaves, therefore, have to fulfill two contrasting requirements: to absorb much light, and to deliver light to all the chloroplasts. Red and blue light is largely absorbed by chloroplasts located near the illuminated surface of the leaf, because chlorophylls preferentially absorb these wavelengths. Light penetrating deeper inside the leaf is mostly green. Chlorophylls are less efficient in absorbing green light. However, multiple scattering increases the path length of light such that leaves absorb 80% of the green light incident on the leaves (for review, see Terashima et al., 2009). The cell shape of the spongy tissue enhances light scattering. There are few comparative studies on in situ absorption coefficients between the palisade and spongy tissues. In Spinacia oleracea, the difference in the in situ absorption coefficient between these tissues was small (Vogelmann and Evans, 2002), while in other species, including Camellia japonica, Helianthus annuus, and Antirrhinum majus, considerable differences were reported (Terashima et al., 2009; Brodersen and Vogelmann, 2010). The question of whether the irregular shape of spongy tissue plays a role in the efficient absorption of green light should be addressed in more species.

It has been argued that the spongy tissue may facilitate lateral CO2 diffusion from substomatal cavities. However, in many herbaceous species, lateral diffusion can also occur in the palisade tissue through gaps between the lateral walls of the neighboring cells. Such herbaceous species tend to have stomata with substomatal cavities on the adaxial leaf surfaces as well, and the size of the cavities is sufficient for CO2 homogenization (Parkhurst, 1977). Thus, this casts doubt on the hypothesis that the function of the spongy tissue is related to the facilitation of lateral CO2 diffusion. Instead, the effects on the lengthening of the light path may be more important.

It is well established that there is a vertical gradient in chloroplast properties within a leaf. Several theoretical models of this phenomenon have been developed (for review, see Terashima et al., 2009). According to one of the theories, the maximum photosynthetic efficiency will be realized if the photosynthetic capacity is proportional to light absorption at each point. However, the gradient of the absorption profile is generally steeper than that of the photosynthetic capacity (for review, see Terashima et al., 2009). The reason for this imperfect matching is not clear. Certainly, the abaxial leaf surfaces also receive some reflected light from the surroundings, which would affect the gradient of photosynthetic capacity. However, a more likely explanation is that the dynamic range for light acclimation of chloroplasts is limited: chloroplasts cannot fully acclimate to very high or very low light.

When the leaf is illuminated with white light from the adaxial side, the chloroplasts located near the adaxial leaf surfaces are light saturated at lower incident irradiances than the chloroplasts near the abaxial surface, because of the imperfect matching of the profiles of photosynthetic capacity and light absorption. When the upper chloroplasts are light saturated, green light that can penetrate deep is more effective in driving photosynthesis of chloroplasts in the deep part. We assessed the efficiency of green light by the differential quantum yield method (Terashima et al., 2009). Figure 5 gives an outline of this method and typical results.

Figure 5.

Figure 5.

Differential quantum yield method (A) and differential quantum yield of green and red monochromatic light in white light in a H. annuus leaf (B). The quantum yield of a given monochromatic light is usually measured at low irradiances in the absence of background light. In the differential quantum yield method, weak monochromatic light is added to the various white irradiances. When the upper chloroplasts are light saturated, green light can drive photosynthesis in chloroplasts located deep in the leaf, where additional red or blue light fails to penetrate, instead being dissipated as heat. For details, see Terashima et al. (2009).

Once absorbed, green light efficiently drives photosynthesis. This has been shown by the conventional quantum yield measurements with monochromatic lights (for review, see Terashima et al., 2009). However, the differential quantum yield measurements revealed that the average quantum yield of green monochromatic light in white light would be comparable to that of red monochromatic light, even when the data were plotted against the incident irradiance. This finding highlights the importance of green light. It is probable that land plants may have been using chlorophylls that strongly absorb red and blue light but only weakly absorb green light, because such a system allows the delivery of light not only to chloroplasts near the irradiated surface but to those deep in the leaf. Given that the dynamic range of acclimation of chloroplasts to irradiance is limited, black leaves that equally absorb light at each wavelength would use light less efficiently at high irradiances than green leaves.

In most optics/photosynthesis studies, collimated light is used. However, in nature, leaves also receive diffuse light and/or collimated light from oblique direction(s). Diffuse light is transmitted less than collimated light (Brodersen and Vogelmann, 2010). When the collimated light hits the leaf obliquely, transmittance also decreases. Therefore, when a significant part of the light is noncollimated, even less light would penetrate deeply into the leaf and green light would be an even more important driver of photosynthesis.

The discrepancy between the profiles of light absorption and photosynthetic capacity in the leaf causes several serious practical problems in measurements of photosynthetic parameters. In many leaf photosynthesis studies, it is implicitly assumed that the leaf behaves like one big chloroplast: all chloroplasts respond to light in accordance with the solution for optimum photosynthesis. In most leaves, this is not the case. Most commercially available fluorometers use red or blue light as the measuring beam that tends to be absorbed by chloroplasts near the leaf surface. Photosynthesis by these surface chloroplasts would be light saturated at lower incident irradiances than chloroplasts deeper into the leaf. The quantum yield of PSII in the light estimated by conventional fluorometry, therefore, is underestimated (Evans, 2009). By contrast, the PSI measurement at 830 nm samples the whole leaf, because this wavelength is only weakly absorbed. The estimation of cyclic electron flow around PSI could be overestimated because of the underestimation of linear electron flow rate based on the PSII quantum yield measurement. The degree of photoinhibition of PSII assessed by fluorometry may not represent the PSII of the whole leaf (for review, see Terashima et al., 2009).

It should also be stressed here that, when the degree of light saturation differs between the chloroplasts near the irradiated surface and those located deep in the leaf, drawdown of CO2 would also differ. When photosynthesis in the chloroplasts near the leaf surface is light saturated while that in the chloroplasts in the deep part of the leaf is not, it is highly probable that the drawdown of CO2 concentration is smaller in the deep chloroplasts. Obviously, this situation affects the estimation of gm. Care should be taken when gm is estimated.

ORGANELLE MOVEMENT

Chloroplast movements in response to light have been studied intensively. In weak light, chloroplasts move so as to maximize light absorption, whereas in strong light, they move to minimize it. These movements (Kadota et al., 2009) as well as anchoring (Kadota and Wada, 1992) are both mediated by actin filaments (Takagi et al., 2009). Phototropins 1 and 2 have been identified as the photoreceptors responsible for weak and strong light responses, respectively (Wada et al. 2003). It is worth pointing out that phototropins are involved in various phenomena closely related to photosynthesis, such as phototropism, leaf flattening, stomatal opening, and chloroplast movement. Nuclei also show an avoidance movement from strong light, which is mediated by phototropin 2, whereas the weak-light response is absent (Iwabuchi et al., 2007).

It has recently been shown that chloroplast movement causes changes in Sc and proportional changes in gm (Tholen et al., 2008). In Arabidopsis (Arabidopsis thaliana) grown at moderate light, irradiation with strong light causes decreases in Sc. These leaves were not typical sun leaves, and cell surfaces parallel to the light direction were less abundant than those perpendicular to the light direction. Avoidance of photodamage occurred at the expense of decreases in gm. In addition, movement of chloroplasts in the upper part of the leaf would allow greater light penetration to deep parts within the leaf. While reducing gm of the upper cells, this would enhance whole leaf light use efficiency.

Mitochondria and peroxisomes are involved in photorespiration together with chloroplasts. Mitochondria also dissipate excessive reducing power generated by chloroplasts in excess light (Noguchi and Yoshida, 2008). Recently, it has been shown that mitochondria (Islam et al., 2009) and peroxisomes (Mano et al., 2002) also move with chloroplasts in the light. For mitochondria, blue light receptors, possibly phototropins, may be involved in the light-induced movement with chloroplasts (Islam et al., 2009).

The arrangement of mitochondria, peroxisomes, and chloroplasts in the cell has been attracting attention. CO2 is released from mitochondria during photorespiration and respiration. Refixation of CO2 by Rubisco will be more efficient if mitochondria are sitting inside a capsule made of chloroplasts covering cell surfaces. In rice leaves, Sage and Sage (2009) found that sheet-like extensions of chloroplasts cover almost all the surfaces of plasma membranes facing intercellular spaces and that mitochondria are almost exclusively distributed within the chloroplast capsule. This arrangement would also facilitate the refixation of NH3 released in the course of photorespiration. Although reassimilation of ammonia is costly (using ATP and ferredoxin), the loss of ammonia is more costly, particularly in plants that take up nitrate or obtain ammonium via nitrogen-fixing symbiosis.

Cell morphogenesis and organelle positioning clearly require further investigations in relation to photosynthesis. Concerning NH3 trapping, it may be worth mentioning that thick mesophyll cell walls found in plants surviving in stressful environments could serve as a good gas trap.

HOW DO LEAVES REGULATE MESOPHYLL SURFACE AREA?

Thick leaves having extensive Smes are advantageous to realize high photosynthetic rates in high-light environments. An ecotype of the Japanese beech F. crenata, on the Pacific side of Japan, prepares all leaves in its winter buds. Both the number of leaves and the cell layers in the palisade tissue can be counted in overwintering buds. Whether the leaves unfold as sun leaves or shade leaves is determined in the previous year. When such sun leaves on the shoot are artificially heavily shaded, winter buds prepare shade leaves with one cell-layered palisade tissue. The structure of the current-year leaves in this late successional species is determined to a large extent by the light environment or photosynthetic productivity of the previous year (Uemura et al., 2000). On the other hand, in several temperate evergreen species, leaf development depends on the average irradiance level (Niinemets et al., 2004), suggesting greater anatomical plasticity. The capacity for adjustment to new light environments depends on the stage of leaf development. Young developing leaves can acclimate to the new light conditions, while older leaves with partly or entirely lignified cell walls can only partly acclimate to altered light conditions (Yamashita et al., 2002).

When leaves develop successively, developing young leaves are covered by older leaves and may be unable to directly sense the light environment outside the bud. Using C. album plants, Yano and Terashima (2001) found that the number of cell layers in the palisade tissue of developing young leaves was determined depending on the light environment of mature leaves, irrespective of the light conditions of the developing young leaves. When mature leaves were exposed to strong light, periclinal divisions of palisade-tissue precursor cells occurred in young leaves that produced two cell layers of palisade tissue, whereas when mature leaves were kept in weak light, such periclinal divisions did not occur.

Stomatal density of developing leaves may also be regulated by the conditions of mature leaves (e.g. CO2 or light; Lake et al., 2001). Molecular mechanisms for these systemic regulations are unknown. However, such mechanisms may play an important role in allowing a leaf to function efficiently immediately after unfolding.

CONCLUDING REMARKS

We have argued several structural features that appear to be highly important for efficient leaf photosynthesis. Nevertheless, it is still essential to assess their functional roles quantitatively. It is also necessary to assess such relationships across a variety of plant species and to determine how the structure-function relationships vary depending on the ecological strategy of the species, such as the position in succession and plant functional type (Hallik et al., 2009a, 2009b).

Further clarifications of the mechanisms that are responsible for these adaptive morphogenesis and organellar movements are strongly awaited. Such developmental and/or cell biological studies should be made together with actual measurements of photosynthetic parameters. Overall, we believe that it is highly relevant to promote ecologically meaningful developmental studies and cell biological studies through the close collaboration of molecular physiologists, cell biologists, and ecophysiologists.

Acknowledgments

We thank Drs. Shingo Takagi, Riichi Oguchi, Amane Makino, Kaoru Kitajima, and Youshi Tazoe for invaluable inputs. We also thank Drs. John Evans and Rowan Sage as well as an anonymous reviewer for constructive comments. I.T. thanks late Prof. Michihiro Kasahara (died November 4, 2010) for his enthusiastic instruction of how to tackle aquaporins.

References

  1. Brodersen CR, Vogelmann TC. (2010) Do changes in light direction affect absorption profiles in leaves? Funct Plant Biol 37: 403–412 [Google Scholar]
  2. Evans JR. (2009) Potential errors in electron transport rates calculated from chlorophyll fluorescence as revealed by a multilayer leaf model. Plant Cell Physiol 50: 698–706 [DOI] [PubMed] [Google Scholar]
  3. Evans JR, Kaldenhoff R, Genty B, Terashima I. (2009) Resistances along the CO2 diffusion pathway inside leaves. J Exp Bot 60: 2235–2248 [DOI] [PubMed] [Google Scholar]
  4. Evans JR, Loreto F. (2000) Acquisition and diffusion of CO2 in higher plant leaves. Leegood RC, Sharkey TD, von Caemmerer S, , Photosynthesis: Physiology and Metabolism. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp 321–351 [Google Scholar]
  5. Fabre N, Reiter IM, Becuwe-Linka N, Genty B, Rumeau D. (2007) Characterization and expression analysis of genes encoding α and β carbonic anhydrases in Arabidopsis. Plant Cell Environ 30: 617–629 [DOI] [PubMed] [Google Scholar]
  6. Flexas J, Ribas-Carbó M, Diaz-Espejo A, Galmés J, Medrano H. (2008) Mesophyll conductance to CO2: current knowledge and future prospects. Plant Cell Environ 31: 602–621 [DOI] [PubMed] [Google Scholar]
  7. Flexas J, Ribas-Carbó M, Hanson DT, Bota J, Otto B, Cifre J, McDowell N, Medrano H, Kaldenhoff R. (2006) Tobacco aquaporin NtAQP1 is involved in mesophyll conductance to CO2 in vivo. Plant J 48: 427–439 [DOI] [PubMed] [Google Scholar]
  8. Galmés J, Flexas J, Keys AJ, Cifre J, Mitchell RAC, Madgwick PJ, Haslam RP, Medrano H, Parry MA. (2005) Rubisco specificity factor tends to be larger in plant species from drier habitats and in species with persistent leaves. Plant Cell Environ 28: 571–579 [Google Scholar]
  9. Hallik L, Kull O, Niinemets Ü, Aan A. (2009a) Contrasting correlation networks between leaf structure, nitrogen and chlorophyll in herbaceous and woody canopies. Basic Appl Ecol 10: 309–318 [Google Scholar]
  10. Hallik L, Niinemets Ü, Wright IJ. (2009b) Are species shade and drought tolerance reflected in leaf-level structural and functional differentiation in Northern Hemisphere temperate woody flora? New Phytol 184: 257–274 [DOI] [PubMed] [Google Scholar]
  11. Hanba YT, Miyazawa SI, Terashima I. (1999) Influences of leaf thickness on internal resistance to CO2 diffusion and δ13C in leaf dry matter. Funct Ecol 13: 632–639 [Google Scholar]
  12. Hanba YT, Shibasaka M, Hayashi Y, Hayakawa T, Kasamo K, Terashima I, Katsuhara M. (2004) Overexpression of the barley aquaporin HvPIP2;1 increases internal CO(2) conductance and CO(2) assimilation in the leaves of transgenic rice plants. Plant Cell Physiol 45: 521–529 [DOI] [PubMed] [Google Scholar]
  13. Islam MS, Niwa Y, Takagi S. (2009) Light-dependent intracellular positioning of mitochondria in Arabidopsis thaliana mesophyll cells. Plant Cell Physiol 50: 1032–1040 [DOI] [PubMed] [Google Scholar]
  14. Iwabuchi K, Sakai T, Takagi S. (2007) Blue light-dependent nuclear positioning in Arabidopsis thaliana leaf cells. Plant Cell Physiol 48: 1291–1298 [DOI] [PubMed] [Google Scholar]
  15. Jia W, Davies WJ. (2007) Modification of leaf apoplastic pH in relation to stomatal sensitivity to root-sourced abscisic acid signals. Plant Physiol 143: 68–77 [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Kadota A, Wada M. (1992) Photoinduction of formation of circular structures by microfilaments on chloroplasts during intracellular orientation in protonemal cells of the fern Adiantum capillus-veneris. Protoplasma 167: 97–107 [Google Scholar]
  17. Kadota A, Yamada N, Suetsugu N, Hirose M, Saito C, Shoda K, Ichikawa S, Kagawa T, Nakano A, Wada M. (2009) Short actin-based mechanism for light-directed chloroplast movement in Arabidopsis. Proc Natl Acad Sci USA 106: 13106–13111 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Katsuhara M, Hanba YT, Maeshima M. (2008) Extended roles of plant aquaporins in plasma membrane and endomembranes: diversity of plant aquaporins in physiological function and intracellular localization. Funct Plant Biol 35: 1–14 [DOI] [PubMed] [Google Scholar]
  19. Lake JA, Quick WP, Beerling DJ, Woodward FI. (2001) Plant development: signals from mature to new leaves. Nature 411: 154. [DOI] [PubMed] [Google Scholar]
  20. Makino A, Mae T, Ohira K. (1985) Enzymic properties of ribulose-15-bisphosphate carboxylase/oxygenase purified from rice leaves. Plant Physiol 79: 57–61 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Mano S, Nakamori C, Hayashi M, Kato A, Kondo M, Nishimura M. (2002) Distribution and characterization of peroxisomes in Arabidopsis by visualization with GFP: dynamic morphology and actin-dependent movement. Plant Cell Physiol 43: 331–341 [DOI] [PubMed] [Google Scholar]
  22. Miyazawa SI, Terashima I. (2001) Slow chloroplast development in the evergreen broad-leaved tree species: relationship between leaf anatomical characteristics and photosynthetic rate during leaf development. Plant Cell Environ 24: 279–291 [Google Scholar]
  23. Niinemets Ü, Díaz-Espejo A, Flexas J, Galmés J, Warren CR. (2009a) Importance of mesophyll diffusion conductance in estimation of plant photosynthesis in the field. J Exp Bot 60: 2271–2282 [DOI] [PubMed] [Google Scholar]
  24. Niinemets Ü, Díaz-Espejo A, Flexas J, Galmés J, Warren CR. (2009b) Role of mesophyll diffusion conductance in constraining potential photosynthetic productivity in the field. J Exp Bot 60: 2249–2270 [DOI] [PubMed] [Google Scholar]
  25. Niinemets Ü, Kull O, Tenhunen JD. (2004) Within canopy variation in the rate of development of photosynthetic capacity is proportional to integrated quantum flux density in temperate deciduous trees. Plant Cell Environ 27: 293–313 [Google Scholar]
  26. Niinemets Ü, Reichstein M. (2003) Controls on the emission of plant volatiles through stomata: a sensitivity analysis. J Geophys Res 108: 4208–4224 [Google Scholar]
  27. Nobel PS. (2004) Physicochemical and Environmental Plant Physiology, Ed 3. Elsevier Academic Press, Burlington, MA [Google Scholar]
  28. Noguchi K, Yoshida K. (2008) Interaction between photosynthesis and respiration in illuminated leaves. Mitochondrion 8: 87–99 [DOI] [PubMed] [Google Scholar]
  29. Oguchi R, Hikosaka K, Hirose T. (2003) Does the photosynthetic light-acclimation need change in leaf anatomy? Plant Cell Environ 26: 505–512 [Google Scholar]
  30. Oguchi R, Hikosaka K, Hirose T. (2005) Leaf anatomy as a constraint for photosynthetic acclimation: differential responses in leaf anatomy to increasing growth irradiance among three deciduous trees. Plant Cell Environ 28: 916–927 [Google Scholar]
  31. Oguchi R, Hikosaka K, Hiura T, Hirose T. (2006) Leaf anatomy and light acclimation in woody seedlings after gap formation in a cool-temperate deciduous forest. Oecologia 149: 571–582 [DOI] [PubMed] [Google Scholar]
  32. Parkhurst DF. (1977) A three-dimensional model for CO2 uptake by continuously distributed mesophyll in leaves. J Theor Biol 67: 471–488 [DOI] [PubMed] [Google Scholar]
  33. Pons TL, Flexas J, von Caemmerer S, Evans JR, Genty B, Ribas-Carbo M, Brugnoli E. (2009) Estimating mesophyll conductance to CO2: methodology, potential errors, and recommendations. J Exp Bot 60: 2217–2234 [DOI] [PubMed] [Google Scholar]
  34. Sage TL, Sage RF. (2009) The functional anatomy of rice leaves: implications for refixation of photorespiratory CO2 and efforts to engineer C4 photosynthesis into rice. Plant Cell Physiol 50: 756–772 [DOI] [PubMed] [Google Scholar]
  35. Stumm W, Morgan JJ. (1996) Aquatic Chemistry: Chemical Equilibria and Rates in Natural Waters, Ed 3. Wiley, New York [Google Scholar]
  36. Takagi S, Takamatsu H, Sakurai-Ozato N. (2009) Chloroplast anchoring: its implications for the regulation of intracellular chloroplast distribution. J Exp Bot 60: 3301–3310 [DOI] [PubMed] [Google Scholar]
  37. Terashima I, Fujita T, Inoue T, Chow WS, Oguchi R. (2009) Green light drives leaf photosynthesis more efficiently than red light in strong white light: revisiting the enigmatic question of why leaves are green. Plant Cell Physiol 50: 684–697 [DOI] [PubMed] [Google Scholar]
  38. Terashima I, Hanba YT, Tazoe Y, Vyas P, Yano S. (2006) Irradiance and phenotype: comparative eco-development of sun and shade leaves in relation to photosynthetic CO2 diffusion. J Exp Bot 57: 343–354 [DOI] [PubMed] [Google Scholar]
  39. Terashima I, Miyazawa SI, Hanba YT. (2001) Why are sun leaves thicker than shade leaves? Consideration based on analyses of CO2 diffusion in the leaf. J Plant Res 114: 93–105 [Google Scholar]
  40. Tholen D, Boom C, Noguchi K, Ueda S, Katase T, Terashima I. (2008) The chloroplast avoidance response decreases internal conductance to CO2 diffusion in Arabidopsis thaliana leaves. Plant Cell Environ 31: 1688–1700 [DOI] [PubMed] [Google Scholar]
  41. Uehlein N, Otto B, Hanson DT, Fischer M, McDowell N, Kaldenhoff R. (2008) Function of Nicotiana tabacum aquaporins as chloroplast gas pores challenges the concept of membrane CO2 permeability. Plant Cell 20: 648–657 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Uemura A, Ishida A, Nakano T, Terashima I, Tanabe H, Matsumoto Y. (2000) Acclimation of leaf characteristics of Fagus species to previous-year and current-year solar irradiances. Tree Physiol 20: 945–951 [DOI] [PubMed] [Google Scholar]
  43. Vogelmann TC, Evans JR. (2002) Profiles of light absorption and chlorophyll within spinach from chlorophyll fluorescence. Plant Cell Environ 25: 1313–1323 [Google Scholar]
  44. Wada M, Kagawa T, Sato Y. (2003) Chloroplast movement. Annu Rev Plant Biol 54: 455–468 [DOI] [PubMed] [Google Scholar]
  45. Yamashita N, Koike N, Ishida A. (2002) Leaf ontogenetic dependence of light acclimation in invasive and native subtropical trees of different successional status. Plant Cell Environ 25: 1341–1356 [Google Scholar]
  46. Yano S, Terashima I. (2001) Separate localization of light signal perception for sun or shade type chloroplast and palisade tissue differentiation in Chenopodium album. Plant Cell Physiol 42: 1303–1310 [DOI] [PubMed] [Google Scholar]

Articles from Plant Physiology are provided here courtesy of Oxford University Press

RESOURCES