Traditionally, it was believed that C4 photosynthesis required two types of chlorenchyma cells to concentrate CO2 within the leaf. However, several species have been identified that perform C4 photosynthesis using dimorphic chloroplasts within an individual cell. The goal of this research was to determine how growth under limited light affects leaf structure, biochemistry and efficiency of the single-cell CO2-concentrating mechanism in Bienertia sinuspersici.
Measurements of rates of CO2 assimilation and CO2 isotope exchange in response to light intensity and O2 were used to determine the efficiency of the CO2-concentrating mechanism in plants grown under moderate and low light. In addition, enzyme assays, chlorophyll content and light microscopy of leaves were used to characterize acclimation to light-limited growth conditions.
There was acclimation to growth under low light with a decrease in capacity for photosynthesis when exposed to high light. This was associated with a decreased investment in biochemistry for carbon assimilation with only subtle changes in leaf structure and anatomy. The capture and assimilation of CO2 delivered by the C4 cycle was lower in low-light-grown plants.
Low-light-grown plants were able to acclimate to maintain structural and functional features for the performance of efficient single-cell C4 photosynthesis.
If you can't find a tool you're looking for, please click the link at the top of the page to "Go to old article view". Alternatively, view our Knowledge Base articles for additional help. Your feedback is important to us, so please let us know if you have comments or ideas for improvement.
In C4 plants, CO2 is concentrated around the enzyme ribulose 1,5-bisphosphate carboxylase/oxygenase (Rubisco), which reduces the rate of photorespiration and increases the rate of photosynthesis (Hatch, 1987; von Caemmerer & Furbank, 1999). In Kranz-type C4 plants, this occurs by diffusive resistance of CO2 from the site of decarboxylation of C4 acids in bundle sheath (BS) to the mesophyll cells (MC). This resistance is dependent on several factors, including the site of decarboxylation in different C4 subtypes, liquid phase resistance to CO2 diffusion in the cytosol, the position of organelles and the BS cell wall (von Caemmerer & Furbank, 2003). However, some CO2 diffuses out of the BS cells, reducing photosynthetic efficiency, which increases the number of absorbed quanta per CO2 assimilated (Skillman, 2008; Ubierna et al., 2013). Therefore, at a given rate of delivery of CO2 to the BS cells via the C4 cycle, the photosynthetic efficiency depends on the fraction of the CO2 fixed by the Calvin–Benson cycle relative to the fraction lost by conductance of CO2 out of the BS cells. Thus, the efficiency of the CO2-concentrating mechanism can be estimated as leakiness (ϕ), defined as the fraction of CO2 fixed by phosphoenolpyruvate carboxylase (PEPC) in the C4 cycle that subsequently diffuses out of the BS cells following decarboxylation of C4 acids (Farquhar, 1983; von Caemmerer, 2003). Leakiness results in a decrease in photosynthetic efficiency, as the C4 cycle consumes energy; in malic enzyme-type C4 species, two ATP molecules are required per turn of the cycle for the regeneration of phosphoenolpyruvate (PEP) (Hatch & Osmond, 1976; Pengelly et al., 2010).
Unfortunately, ϕ is not directly measurable, but can be estimated by comparing measurements of photosynthetic discrimination against 13CO2 (Δobs) with the theoretical model of 13CO2 discrimination (Δmodeled; Evans et al., 1986; von Caemmerer et al., 1997; Ubierna et al., 2013). Numerous studies have shown that Δobs increases with decreasing photosynthetically active radiation (PAR), suggesting a decrease in photosynthetic efficiency of C4 plants under low PAR (Henderson et al., 1992; Cousins et al., 2006, 2008; Kromdijk et al., 2008, 2010; Tazoe et al., 2008; Pengelly et al., 2010; Ubierna et al., 2013). This increase in Δobs has been attributed to an increase in ϕ caused by a disruption in the balance between the C3 and C4 cycles, with decreased CO2 concentrations in the BS cells (Cs) and increased rates of oxygen fixation vo (Henderson et al., 1992; Cousins et al., 2006; Tazoe et al., 2006, 2008; Kromdijk et al., 2008, 2010; Pengelly et al., 2010). However, Ubierna et al. (2011, 2013) demonstrated that factors other than ϕ, such as carbon isotope fractionation associated with respiration and miscalculations of Cs, can also explain some of the reported increase in Δobs under low PAR.
Traditionally, it was thought that dual-cell Kranz-type anatomy was required for efficient C4 photosynthesis, with features of BS cells providing resistance to CO2 leakage (von Caemmerer, 2003; von Caemmerer & Furbank, 2003). However, it has been demonstrated in four terrestrial plants within the Chenopodiaceae family (Bienertia sinuspersici, B. cycloptera, B. kavirense and Suaeda aralocaspica) that Kranz-type anatomy is not required for C4 photosynthesis (Voznesenskaya et al., 2001; Sage, 2002; Edwards et al., 2004; Akhani et al., 2005, 2012). Instead, these halophytic species from the Arabian Peninsula utilize dimorphic chloroplasts within a single cell to perform NAD-malic enzyme-type C4 photosynthesis (Voznesenskaya et al., 2001; Akhani et al., 2005, 2009; Offermann et al., 2011). Bienertia sinuspersici has bienertioid anatomy, in which peripheral chloroplasts, oppressed to the plasma membrane, support the C4 CO2-concentrating mechanism, and a group of centrally located chloroplasts and mitochondria is the site for C4 acid decarboxylation and Rubisco fixation (Freitag & Stichler, 2002; Voznesenskaya et al., 2002; Edwards et al., 2004; Offermann et al., 2011; Sharpe & Offermann, 2013). It has been suggested that single-cell C4 plants maintain the resistance to CO2 diffusion by manipulating cell size and the distance between the chloroplasts involved in the C3 and C4 cycles (Leisner et al., 2010; King et al., 2012). As CO2 diffuses 10 000 times more slowly through water than air, single-cell C4 plants may gain increased resistance to CO2 diffusion by increasing cell size (von Caemmerer, 2003).
King et al. (2012) found that single-cell C4 plants had similar photosynthetic efficiencies to their Kranz-type counterparts under high PAR, but ϕ increased more under low PAR relative to ϕ of Kranz-type C4 plants, even when taking into account changes in Cs, photorespiration via ribose-1,5-bisphosphate (RuBP) oxygenation and the contribution of mitochondrial day respiration to 13CO2 exchange. However, measurements of ϕ presented in King et al. (2012) were made under low O2 levels (4.6 kPa), reducing rates of photorespiration and complicating estimates of Cs under low PAR (Ubierna et al., 2013). In addition, King et al. (2012) assumed that the substrate for day respiration was the same as the substrate for photosynthesis (i.e. the isotopic signature of CO2 was the same for photosynthesis and respiration, Wingate et al., 2007). However, this assumption might not be valid when the isotopic signature of the growth CO2 is different from that of the measurement CO2. Indeed, the manipulation of differences in the isotopic composition of growth and measurement CO2 has been elegantly used to demonstrate the contribution of photorespiration and day respiration to leaf CO2 exchange (Gillon & Griffiths, 1997; Gillon et al., 1998).
In B. sinuspersici, the carbon isotope composition (δ13C) of leaf dry matter, which is largely determined by fractionation during photosynthesis, varies considerably. For example, plants grown in chambers ranged from −19.3‰ in King et al. (2012) to a characteristic C4 signature of −14.1‰ in Smith et al. (2009). However, analyses of Bienertia species in natural habitats consistently show C4-type isotope signatures (Akhani et al., 2005, 2009, 2012). Some of the variation in chamber-grown plants is attributed to developmental differences in young and mature leaf tissue (Voznesenskaya et al., 2002; King et al., 2012), but it has also been suggested that differences in growth conditions affect δ13C (Leisner et al., 2010; King et al., 2012). However, it has not been demonstrated how dry matter δ13C and cellular leaf structure relate to ϕ in response to growth under different light conditions.
Therefore, the goal of this study was to test the relationship among leaf structure, capacity of carboxylases and photosynthetic efficiency in the single-cell C4 plant B. sinuspersici grown under moderate and low light (ML and LL, respectively). The objectives were as follows: to determine how the photosynthetic efficiency of the CO2-concentrating mechanism responds to light-limited growth conditions; to see how leaf cell structure and biochemistry are affected by growth light conditions; and to use differences in growth and measurement CO2 isotope signatures to determine the influence of day respiration on measurements of Δobs and ϕ in response to changes in PAR. This was accomplished by measuring leaf carbon isotope discrimination with a tunable diode laser absorption spectroscope (TDLAS), short-term labeling of recent photoassimilates with −7 and −58‰ CO2, spectrometer assays of Rubisco and PEPC activities, and light microscopy to compare leaf cell structure in plants grown under ML vs LL.
Materials and Methods
Growth conditions and plant propagation
Bienertia sinuspersici Akhani was grown in an environmental chamber (Econair Ecological Chambers Inc., Winnipeg, MB, Canada) under ambient CO2 (isotopic signature −10.7‰) at 32°C : 18°C, 20% : 40% relative humidity and 16 h : 8 h day : night. PAR was 500–600 μmol quanta m−2 s−1 for ML and 150–200 μmol quanta m−2 s−1 for LL at pot level. As reported previously (King et al., 2012), all plants were top watered with tap water daily and once a week with 50 mM NaCl and Peters 21-5-20 fertilizer (Scotts Miracle-Gro, Marysville, OH, USA).
Plants were grown side by side with low light provided by a shade structure made of 0.25-in PVC pipe covered with a black shade cloth (Polysack Plastic Industries, Nir Yitzhak, Negev, Israel), which reduced light quantity by 60%, but had no effect on light quality. Plants grown under LL were rotated once a week to ensure uniform treatment, and ML plants were randomly positioned in the center of the chamber.
Plants were propagated from both cuttings and seeds (for details on plant propagation, see King et al., 2012). Once the propagated plants showed substantial root and vegetative growth (c. 6–8 wk), they were transferred to 7.6-l pots, one plant per pot, in a mixture of c. one-half potting soil (Sun Gro Horticulture, Seba Beach, AB, Canada) and one-half Turface (Profile Products LLC, Buffalo Grove, IL, USA). The ML plants were placed in the center of the chamber (PAR = 500–600 μmol quanta m−2 s−1) and the LL plants were initially placed under a slanted shade screen (PAR = 300 μmol quanta m−2 s−1) for 1.5–2 wk before being placed under the shade structure (PAR = 150–200 μmolquanta m−2 s−1). Plants grown in ML were analyzed 3–4 wk after transplanting; to compensate for their slower growth, LL plants were analyzed 5–6 wk after being placed under the shade structure. The LL plants were measured when the above-ground crown was of a similar size to that of the ML plants at the time of measurements. As a result of the differences in growth rate, there were two separate cohorts of ML plants, whereas a single cohort of LL plants was used during the measurements to ensure similar-sized plants.
Measurements of gas exchange and leaf carbon isotope exchange
A LI-6400XT (LI-COR Biosciences, Lincoln, NE, USA) was coupled to a TDLAS (model TGA100; Campbell Scientific, Inc., Logan, UT, USA) to measure online carbon isotope discrimination for both light- and O2-response curves (Sun et al., 2012; Ubierna et al., 2013). Isotope calibration consisted of a zero CO2 tank, three mixing tanks from 3.3 to 10 Pa CO2 with the same isotopic signature, a calibration tank (Liquid Technology Corporation, Apopka, FL, USA), followed by the LI-COR reference and sample (Ubierna et al., 2013). Boundary layer conductance depending on leaf area and flow rate was calculated according to Ubierna et al. (2013). Average values ranged from 5.1 ± 0.4 to 6.7 ± 0.3 mol m−2 s−1, which had an insignificant impact on the calculated parameters.
Photosynthetic discrimination Δ13C (Δobs) was calculated according to Evans et al. (1986):
where ξ is:
where δo and δe are the δ13C values of air entering (e) and leaving (o) the chamber, respectively, and Pe and Po are the CO2 partial pressures entering and leaving the chamber, respectively. Measured values are reported in Supporting Information Figs S1 and S2.
Light- and O2-response curves
Medium-aged branches, having few minor branches, were selected for analysis. The branch was placed in an opaque conifer chamber (LI-COR Biosciences) with an RGB LED light source (LI-COR Biosciences) attached to a LI-6400XT. Before sealing the chamber, the branch was photographed and the projected leaf area was estimated using ImageJ (US National Institutes of Health, Bethesda, MD, USA). The branch was acclimated for at least 1 h in the LI-COR chamber at 25°C (leaf temperature), 2000 μmol quanta m−2 s−1, 3.8 Pa CO2 and 18.4 kPa O2. Energy balance calculations of leaf temperature and boundary layer conductance were determined from the LI-COR software. The relative humidity within the chamber was between 50% and 70% with a vapor pressure deficit of 1–2 kPa. Ambient CO2 was scrubbed from the air entering the LI-6400 and added back from a −7 ± 0.11‰ CO2 cartridge (iSi GmbH, Vienna, Austria). To test the contribution of day respiration to leaf CO2 isotope exchange, light-response curves were additionally measured with a −58 ± 1.4‰ CO2 tank. CompoundQ (Apiezon Products, M&I Materials Ltd, Manchester, UK) was placed around the conifer chamber gaskets and the plant stem to minimize leaks. The light-response curves were measured in the following order: 2000, 1500, 1000, 800, 600, 500, 400 and 2000 μmol quanta m−2 s−1. Eight measurements of photosynthesis over c. 30 min were taken before PAR was changed and after the LI-COR infrared gas analyzers (IRGAs) were matched. The respiration rate was measured at 3.8 Pa CO2 and 18.4 kPa O2 after 30 min of dark adaptation following each light- or O2-response curve. After the light-response curves, six mature leaves towards the base of the branch, nine medium leaves and nine young leaves towards the apex were harvested for dry matter δ13C and total leaf nitrogen. Mature leaves were defined as those closest to developing axillary branches with few salt glands, whereas medium leaves were fully expanded with many salt glands, and young leaves were not fully expanded with numerous salt glands (Edwards et al., 2004).
The O2-response curves of photosynthesis and isotopic exchange were measured at 4.6, 13.8, 18.4, 27.6 and 36.9 kPa O2 (5%, 15%, 20%, 30% and 40% O2) at 2000 μmol quanta m−2 s−1 and 3.8 Pa CO2. Plants were acclimated for at least 1 h under the first O2 concentration with the order of the oxygen concentrations randomly selected. Oxygen concentration was controlled using mass flow controls (Aalborg Instruments & Controls Inc., Orangeburg, NY, USA) connected to compressed nitrogen and oxygen tanks and a 2-l mixing flask.
Photosynthetic efficiency (leakiness)
Leakiness (ϕ) for LL plants was estimated under high PAR (2000 and 1500 μmol quanta m−2 s−1) using the enzyme-limited model of C4 photosynthesis (von Caemmerer, 2000; Ubierna et al., 2013), and under low PAR (1000, 800, 600, 500 and 400 μmol quanta m−2 s−1) using the light-limited model of C4 photosynthesis (von Caemmerer, 2000; Ubierna et al., 2013). The light-limited model of C4 photosynthesis was used under all PAR levels for ML plants, because ML plants did not reach saturation under measurement PAR. The enzyme-limited model of C4 photosynthesis was used to estimate leakiness for all O2 partial pressures for LL plants. However, the light-limited model of C4 photosynthesis was used to estimate leakiness for all O2 partial pressures for ML plants, because these plants did not reach saturation under 2000 μmol quanta m−2 s−1. Leakiness, including the ternary effect (Farquhar & Cernusak, 2012; Ubierna et al., 2013), under high PAR was estimated as (see Table 1):
Table 1. Definitions and units for symbols in the text
where t is the ternary effect (see Farquhar & Cernusak, 2012; Ubierna et al., 2013), Pi is the CO2 partial pressure in the intercellular air space, Pa is the atmospheric partial pressure of CO2, is the weighted fractionation across the boundary layer and stomata in series (4.4‰), (−5.7‰) is the effect of CO2 dissolution and PEPC activity at 25°C (Farquhar, 1983), Rd is an estimate of leaf mitochondrial respiration occurring during the day (based on the rate measured in the dark), Rm is the rate of mitochondrial day respiration in the MCs, calculated as Rm = 0.5Rd, A is the net photosynthetic rate, is Rubisco fractionation (30‰) and s (1.8‰) is the fractionation of CO2 leaving the BS cells (Roeske & O'Leary, 1984; Henderson et al., 1992). The value was calculated as in Wingate et al. (2007):
where is the 13C fractionation during decarboxylation, including measurement artifacts, e is the respiratory fractionation during decarboxylation, −6‰ (Wingate et al., 2007), and e* is the difference between the use of recent photoassimilate and the use of other substrates for day respiration as in Wingate et al. (2007). To model ϕ, assuming all respiratory consumption of substrates occurs using recent photoassimilates, the following equation for e* was used:
where δ13Csample and Δobs are the isotopic signature of the LI-COR sample line and the photosynthetic discrimination during the measurement, respectively. However, the influence of day respiration on fractionation with the use of old photoassimilates e* was modeled as:
where δ13Cgrowth is the isotopic signature of the air in which the plants were grown (−10.7‰ in this study) and δ13Cdry is the isotopic signature of the plant dry matter (−21.3 and −17.9‰ for LL and ML plants, respectively).
Leakiness under low PAR was modeled according to Ubierna et al. (2013) incorporating the ternary effect (Farquhar & Cernusak, 2012; Ubierna et al., 2013):
The terms b3 and b4 are defined as (Farquhar, 1983):
where f is the fraction during photorespiration, 11.6‰ (Lanigan et al., 2008), and vo and vc are the rates of oxygenation and carboxylation respectively, by Rubisco. The parameters vp, vo, vc and total electron flux (Jt) were estimated under light-limited conditions using models described previously (von Caemmerer, 2000; Ubierna et al., 2011, 2013; eqns 3–18).
Dry matter N content and δ13C
Frozen leaf tissue, consisting of two mature leaves, three medium leaves and three young leaves, was freeze–dried for 48 h and ground to a fine powder. Total N was measured by combustion in an elemental analyzer (ECS 4010; Costech Analytical, Valencia, CA, USA). Dry matter δ13C was determined by mass spectroscopy (Delta PlusXP; Thermofinnigan, Bremen, Germany) and calculated as [(Rsample − Rstandard)/Rstandard] × 1000, where Rsample is the ratio of 13C/12C in the sample and Rstandard is the ratio of 13C/12C in the standard, V-Pee Dee Belemnite.
Specific leaf area (SLA)
To estimate SLA, mature, medium and young leaves were cut from different branches and placed on wet filter paper before obtaining fresh weight and leaf area. Subsequently, leaves were dried in an oven at 60°C for 1 wk and then weighed. Six replicate plants were measured per treatment.
Enzyme assays and chlorophyll content
Enzyme activity was measured for both LL and ML plants according to Cousins et al. (2007). Three leaves (mature, medium and young) from several different branches were taken from a single plant. Leaf area was determined from photographs of leaves using ImageJ and leaves were ground together in a cold room at 4°C in 1000 μl of extraction buffer (50 mM Hepes–KOH, pH 7.8, 1% polyvinylpyrrolidone (PVPP), 1 mM EDTA, 10 mM dithiothreitol, 0.1% Triton X), 5 μl of protease inhibitor cocktail (Sigma) and fine sand. After grinding, the leaf extract was briefly centrifuged and 500 μl of supernatant was incubated with 15 mM MgCl2 and 15 mM NaHCO3 to fully activate Rubisco.
Rubisco activity was determined in an assay buffer of 100 mM EPPS–NaOH, pH 8.0, 20 mM MgCl2, 1 mM EDTA, 10 mM ATP, 50 mM creatine phosphate, 20 mM NaHCO3, 0.2 mM NADH, 12.5 U ml−1 creatine phosphokinase, 250 U ml−1 carbonic anhydrase, 22.5 U ml−1 phosphoglycerate kinase, 20 U ml−1 glyceraldehyde-3-phosphate dehydrogenase, 56 U ml−1 triose-phosphate isomerase and 20 U ml−1 glycerol-3-phosphate dehydrogenase, and the reaction was initiated with 20 μl of 20.4 mM RuBP. PEPC activity was determined in an assay buffer of 100 mM EPPS–NaOH, pH 8.0, 20 mM MgCl2, 1 mM EDTA, 0.2 mM NADH, 5 mM glucose-6 phosphate, 1 mM NaHCO3 and 12 U ml−1 malate dehydrogenase, and the reaction was initiated with 10 μl of 4 mM PEP. Enzyme activity for both assays was determined spectrophotometrically by following the decrease in NADH absorbance over time at 340 nm, correcting for the non-specific decrease in absorbance at 400 nm (Thermo Fisher Scientific Inc., Houston, TX, USA).
Chlorophyll was extracted in 95% ethanol for 48 h in the dark on a shaker at 4°C, and the absorbance of chlorophyll was measured at 649 and 665 nm (any non-specific absorbance measured at 700 nm was subtracted from these values) with a spectrometer. Chlorophyll concentration was calculated according to Ritchie (2006) with five replicates per treatment, consisting of pooled mature, medium and young leaves.
Two square millimeters of medium-aged leaves from the middle of the branch were fixed in 2% paraformaldehyde and 2% glutaraldehyde in 0.1 M phosphate buffer. Subsequently, the samples were transferred to 2.5% glutaraldehyde, 3.5% formaldehyde, 0.1 M sodium cacodylate, 0.12 M sucrose, 10 mM ethylene glycol tetra-acetic acid (EGTA) and 2 mM magnesium chloride overnight at 4°C. The samples were rinsed in a 0.1 M sodium cacodylate buffer and post-fixed in 2% OsO4 for 2 h at room temperature, dehydrated in an ethanol series and embedded in Spurr's resin. Cross-sections (800 nm thick) were made and stained with toluidine blue in 1% sodium borohydrate, and digital images were collected with a camera (Jenoptik ProgRes Camera; Optik, Systeme, Jena, Germany) attached to a compound light microscope (Olympus BH-2; Olympus Optical Co. Ltd, Tokyo, Japan).
Images of five replicate plants from each treatment were analyzed with ImageJ (US National Institutes of Health) to measure the distance between the central chloroplastic compartment (CCC) and the intercellular airspace (IAS) (μm), the total length of MC wall exposed to IAS (μm) and the width of the leaf section analyzed (μm). The corresponding total MC surface area exposed to IAS per (one side) leaf surface area (Ames, μm2 μm−2) was inferred (Evans et al., 1994) as:
where 1.43 is the curvature correction factor taken from Evans et al. (1994). The cross-sectional area of MC and their CCC was measured by light microscopy on cells isolated by gentle maceration of leaves as described by Leisner (2009). Leaf thickness, length and width were measured with digital calipers on medium-aged leaves from the middle of the branch.
Statistical analysis was performed using Statistix software (Analytical Software, Tallahassee, FL, USA). Three-way repeated-measures ANOVAs were used for A, gs, Δ13C and leakiness by PAR, treatment and tank for light-response curves. For O2-response curves, two-way repeated-measure ANOVAs were used to compare ML and LL plants for A, gs, Δ13C and leakiness by O2 and treatment. The branch was the repeated measure for all analyses. Results were deemed to be significant at P < 0.05 and Tukey's test was used for post hoc comparisons. A paired two-tailed Student's t-test was used for enzyme assays, microscopy measurements, SLA, chlorophyll content, dry matter δ13C and dark respiration rate between treatments.
The net rate of CO2 assimilation was higher for ML relative to LL plants under all O2 partial pressures (Fig. 1a). In addition, the net rate of CO2 assimilation declined with increasing partial pressure of O2 in plants from both treatments; however, the O2 sensitivity was greater in LL (1.8% decrease (kPa O2)–1) relative to ML (1.0% decrease (kPa O2)–1) plants (Fig. 1a, Table S1). Stomatal conductance (gs) was higher in ML plants across all partial pressures of O2, but was constant with O2 for both treatments (Fig. 1b, Table S1). There was no difference in Pi/Pa between treatments or in response to O2 (Fig. 1c, Table S1).
Photosynthetic discrimination (Δobs) increased with O2 partial pressure in both ML and LL plants, but Δobs responded more to O2 in LL plants (Fig. 2a, Table S1). In ML plants, there was a gradual increase in ϕ with increasing O2, whereas, in LL plants, there was an immediate rise in ϕ at 13.8 kPa O2, which leveled out after this point (Fig. 2b, Table S1). Leakiness in LL plants was higher than in ML plants under all O2 conditions (Fig. 2b).
In ML plants, there was a strong response of photosynthesis up to full sunlight (PAR = 2000 μmol quanta m−2 s−1), whereas the response in LL plants was more hyperbolic, increasing gradually at higher PAR (Fig. 3a). Rates of net CO2 assimilation (A) were higher in ML relative to LL plants at PAR > 800 μmol m−2 s−1, but similar at PAR < 800 μmol m−2 s−1 (Fig. 3a, Table S2). Rates of assimilation at PAR = 2000 μmol quanta m−2 s−1 (equivalent to full sunlight) in ML plants (c. 40 μmol m−2 s−1) were c. two-fold higher than in LL plants (< 20 μmol m−2 s−1). In addition, in ML plants, A was higher when the source was −58‰ rather than −7‰ CO2 (P <0.01), but not for LL plants (Fig. 3a, Table S2). Stomatal conductance (gs) was higher in ML relative to LL plants across all PAR levels (Fig. 3b, Table S2), regardless of the isotopic signature of the measurement CO2. In ML plants, gs was significantly higher in −58‰ than in −7‰ CO2 (P <0.01) under high PAR, but, in general, gs increased in ML plants with increasing PAR (Fig. 3b, Table S2). The ratio of Pi/Pa was higher in ML relative to LL plants (P <0.01) and decreased with increasing PAR for both treatments (Fig. 3c, Table S2; P <0.01). Values of Pi/Pa were significantly different between CO2 isotopic signatures for both growth conditions and all PAR levels (P <0.01).
Measured Δobs increased more with decreasing PAR in ML relative to LL plants, regardless of the CO2 isotopic signature (Fig. 4a,b). In ML plants, Δobs increased with decreasing PAR more under −58‰ relative to −7‰ CO2 (Fig. 4b); however, there was no significant difference in Δobs between the two measurement CO2 isotopic signatures in LL plants (Fig. 4a, Table S2). Leakiness (ϕ) was calculated assuming the substrate for day respiration was either from recent photosynthate (recent) fixed during the measurements or from previous photosynthate (old) fixed in the growth chamber. Leakiness generally did not respond to changes in PAR in LL plants, except when measurements were made with the −58‰ CO2 source and it was assumed that old photosynthate was the substrate for day respiration (Fig. 4c). Under these conditions, ϕ decreased with decreasing PAR and was significantly different from the other estimates of ϕ below 1000 μmol quanta m−2 s−1. However, ϕ did not change with PAR in LL plants measured under −7‰ CO2, whether the substrate for Rd was assumed to be recent or old photoassimilate (Fig. 4c). In plants grown under ML, ϕ increased with decreasing PAR measured under −58‰ CO2 and assuming recently fixed photoassimilate was used as the substrate for day respiration (Fig. 4d). However, ϕ modeled assuming that plants used old photoassimilate did not increase under decreasing PAR, and was significantly lower at low PAR than ϕ estimated under −58‰ CO2, assuming that recent photoassimilate was the substrate for day respiration. Leakiness in ML plants measured under −7‰ CO2 did not change with declining PAR, assuming either recent or old photoassimilate.
Leaf biochemical characteristics
Rubisco activity was c. two-fold higher and PEPC activity was c.three-fold higher per unit leaf area in ML relative to LL plants (P <0.01; Table 2), although the ratio of PEPC to Rubisco was not significantly different between treatments. In addition, total chlorophyll, chlorophyll a/b and total leaf N were not significantly different between treatments (Table 2). However, the isotopic signature of leaf dry matter (δ13C) was significantly more depleted in 13C in LL (−21 ± 0.3‰) relative to ML (−17.9 ± 0.5‰; P <0.01; Table 2) plants. Rates of dark respiration were significantly higher (c. two-fold) in ML (6.1 ± 0.4 μmol CO2 m−2 s−1) relative to LL (2.7 ± 0.3 μmol CO2 m−2 s−1; P <0.01; Table 2) plants.
Table 2. Leaf biochemical properties of Bienertia sinuspersici grown under low-light (LL, PAR = 150–200 μmol m−2 s−1) and moderate-light (ML, PAR = 500–600 μmol m−2 s−1) conditions
Measurements represent averages ± SE of four to six replications from pooled mature, medium and young leaves. Similar to the measurements of photosynthesis, the rates of dark-type respiration were made on branches and leaves, and the rates are expressed on a projected leaf area. Significant differences between treatments determined with Student's t-test: **, P <0.01.
The SLA, the leaf area to dry mass ratio, was significantly higher in LL relative to ML plants (P <0.01; Table 3). In addition, the ratio of the individual MC area to the area of the CCC was significantly higher in LL relative to ML plants (P <0.05; Table 3). However, leaf length and thickness, the path length from the CCC to the IAS, and the distance of the MCs exposed to the IAS were not significantly different between treatments (Table 3). The planar area per MC was greater in LL relative to ML plants; however, this difference was not significant. The planar area of the CCC was the same between treatments (Table 3).
Table 3. Leaf properties of Bienertia sinuspersici grown under low-light (LL, PAR = 150–200 μmol m−2 s−1) and moderate-light (ML, PAR = 500–600 μmol m−2 s−1) conditions
CCC, central cytoplasmic compartment in mesophyll cells; IAS, intercellular air space.
Measurements show the means and standard errors of five to six replicates from medium-aged leaves for all parameters except SLA. The estimates of SLA were pooled from mature, medium and young leaves. Significant differences between treatments determined with Student's t-test: *, P <0.05; **, P <0.01.
Ames, mesophyll cell length exposed to IAS per leaf area (μm2 μm−2)
2274 ± 392.7
1692 ± 214.1
Our results demonstrate the capacity for photosynthesis and the CO2-concentrating mechanism in the single-cell C4 plant B. sinuspersici is influenced by the growth light condition. ML plants had c. two-fold higher rates of photosynthesis under high PAR, c. two-fold higher Rubisco and c. three-fold higher PEPC activities per unit leaf area, and an estimated c. 20% lower leakiness under current atmospheric levels of O2 (18.4 kPa O2, Fig. 4). This shows that the single-cell C4 plant has a characteristic acclimation potential: when light is limiting during growth, there is less investment in biochemistry for carbon assimilation. However, growth light conditions caused only subtle changes in leaf structure and anatomy, except that the SLA was higher in LL plants. In species with non-succulent planar leaves grown under LL, the leaves are often thinner, cells smaller and, in some cases, layers of cells in the leaf are reduced (Terashima et al., 2006; Pengelly et al., 2010). Below, we discuss how biochemical and structural acclimation under light-limited growth conditions affects photosynthesis in B. sinuspersici.
The CO2-concentrating mechanism in Kranz-type C4 plants reduces the rate of photorespiration by concentrating CO2 around Rubisco (Edwards & Walker, 1983; Hatch, 1987; Keeley & Rundel, 2003; Sage, 2004). Therefore, C4 plants are typically not as sensitive as C3 plants to high O2 concentrations. However, the efficiency of the single-cell CO2-concentrating mechanism to changes in O2 partial pressure has not been studied. King et al. (2012) estimated the photosynthetic efficiency in two single-cell C4 plants, B. sinuspersici and S. aralocaspica, under low O2 (4.6 kPa), and showed that, under these conditions, ϕ in single-cell C4 plants was similar to that of Kranz-type plants. However, the low O2 during these previously published estimates of ϕ in the single-cell C4 plants may have masked an inefficient CO2-concentrating mechanism. If the single-cell C4 plants have an inefficient CO2-concentrating mechanism, Δobs and ϕ should be more sensitive to changes in O2 than expected for C4 plants. Furthermore, differences in leaf anatomy caused by the light-limited growth condition could also affect the O2 sensitivity of the CO2-concentrating mechanism in single-cell C4 plants. In our measurements, as the O2 partial pressure increased, Δobs increased by 3‰ and 2‰ in LL and ML plants, respectively (Fig. 2a,b), suggesting a slightly greater sensitivity to increasing O2 in LL plants.
Rates of net CO2 assimilation (A) were higher in ML relative to LL plants regardless of O2 partial pressure (Fig. 1a). In addition, at ambient CO2 concentrations (3.8 Pa), A decreased with increasing O2 in plants from both growth conditions. However, it should be noted that the O2 sensitivity of A seen in B. sinuspersici has also been demonstrated in several Kranz-type C4 plants (Dai et al., 1993; Maroco et al., 1997, 2000). Some sensitivity of A to O2 can occur in C4 plants as a result of photorespiration, but it is expected to be rather low relative to C3 plants because of the CO2-concentrating mechanism.
In B. sinuspersici, the slightly higher sensitivity of A to O2 in LL plants suggests that the acclimation of the single-cell C4 system to LL altered the capacity and efficiency of the CO2-concentrating mechanism. This is further supported by measurements of leaf CO2 isotope exchange in response to O2, which showed that Δobs increased more in LL relative to ML plants from low to high O2. Furthermore, ϕ increased by 10% in LL plants from 4.6 to 13.8 kPa O2 and then remained constant; however, in ML plants, ϕ decreased by < 10% across all O2 partial pressures (Fig. 2c,d). These estimates of ϕ used the full Wingate et al. (2007) equation ((Eqn 5) and (Eqn 6) in this text), taking into account differences between the substrate for day respiration and the substrate for photosynthesis. This has important implications when comparing ϕ values reported here with those of King et al. (2012) for B. sinuspersici.
In King et al. (2012), estimates of ϕ were determined in ML-grown plants (the isotopic signature of growth CO2 was c.−10‰) and measured under low O2 (4.6 kPa) with a CO2 signature of c. −45‰. The apparent influence of day respiration in King et al. (2012) on 13C fraction during decarboxylation (e′) was estimated using e′ = e + δ13Cmeasurement − δ13Cgrowth. This assumes that the substrate for day respiration is recent photoassimilate, which has an isotopic signature dependent on Δobs and δ13C of the CO2 used during the measurements (Ubierna et al., 2013). However, ϕ will be miscalculated if a plant is using old photoassimilate as a substrate for day respiration and the measurement CO2 has a different isotopic signature from the growth conditions. Therefore, under low PAR, when day respiration represents a greater proportion of branch net CO2 exchange, the estimate of ϕ from King et al. (2012) may be incorrect because of the simplification used to estimate e*. Below, we discuss the influence of e* and differences in growth light conditions on ϕ in response to changes in measurement PAR.
In plants grown under ML and measured under −58 and −7‰ CO2, there was a linear increase in A up to full sunlight. ML plants measured under −58‰ CO2 were a different cohort from those measured under −7‰ CO2, which probably contributed to the differences seen in A and gs (Fig. 3a,b). However, in LL plants, A became saturated at light levels above growth conditions. If plants acclimate, this typically occurs by not over-investing in components that cannot be used in light-limited conditions (e.g. high capacity of carboxylases). This has also been observed in some Kranz-type C4 plants grown under two light levels (Tazoe et al., 2008; Pengelly et al., 2010).
The increase in Δobs as measurement PAR decreased in both LL and ML plants (Fig. 4) is also in agreement with previous reports (see Tazoe et al., 2008; Pengelly et al., 2010; Ubierna et al., 2013). The increase in Δobs at low PAR was more pronounced in ML relative to LL plants, regardless of the isotopic signature of the measurement CO2 (Fig. 4a). Previous studies (Tazoe et al., 2008; Pengelly et al., 2010) have also shown a greater increase in Δobs for plants grown under high light or ML relative to plants grown under LL. It is assumed that day respiration stays relatively constant under all PARs, whereas photosynthesis decreases with decreasing PAR. Therefore, under high PAR, the contribution of respired CO2 to total leaf CO2 isotope exchange is smaller than the contribution under low PAR. In the current study, rates of dark-type respiration were 2.1 and 6.1 μmol CO2 m−2 s−1 for LL and ML plants, respectively (Table 2). Therefore, the contribution of respired CO2 to net CO2 exchange in ML plants was greater than in LL plants. If a plant was measured using CO2 depleted in 13C, the recent photoassimilates would be depleted in 13C. In addition, plants with higher day respiration rates would yield a greater contribution to leaf CO2 exchange, which would influence Δobs. The simplified equation of Wingate et al. (2007) for e′, where e′ = e + (δ13Cmeasurement − δ13Cgrowth), assumes that the isotopic signature of the substrate for day respiration is the same as the isotopic signature of the substrate for photosynthesis (Ubierna et al., 2013). For this assumption to be met, the plant must use recent photoassimilate as a substrate for day respiration; however, this may not always be the case. To test this, we used (Eqn 5) and (Eqn 6) to account for the differences in substrates used for photosynthesis and day respiration.
The estimated ϕ changed from 0.4 to 0.15 with decreasing PAR in LL plants, assuming old photoassimilate (acquired in the growth chambers with CO2 = −10.7‰) and measured with −58‰ CO2 (Fig. 4c). However, there was little response of ϕ to PAR with measurement CO2 of −58‰ and assuming that recent photoassimilate was the substrate for day respiration (Fig. 4c). In addition, when measurements were conducted under −7‰ CO2, there was little change in ϕ in response to PAR (Fig. 4c). This demonstrates that misrepresentation of the isotopic signature of the photoassimilate used for day respiration can misestimate ϕ when there is a significant difference between the isotopic composition of recent and old photoassimilates. Furthermore, in the LL plants, ϕ does not appear to change in response to PAR when correctly accounting for the impact of day respiration on leaf CO2 isotope exchange.
For ML plants measured under −58‰ CO2, values of ϕ increased from 0.4 to 1.4 with declining PAR, assuming that plants used recent photoassimilate for day respiration (Fig. 4d). However, ϕ does not increase with decreasing PAR if ϕ is modeled assuming plants are using old photoassimilate as a substrate for day respiration. In addition, when plants were measured under −7‰ CO2, estimates of ϕ were not influenced by errors in assuming recent vs old substrate for day respiration. Therefore, measurements made at −7‰ CO2 can be used to determine whether plants measured at −58‰ CO2 are using recent or old photoassimilate as the substrate for day respiration. For example, in ML plants, there was a sharp increase in Δobs when measured under −58‰ CO2 (Fig. 4d) relative to plants measured under −7‰ CO2. If plants used old photoassimilate as a substrate for day respiration, there would not have been a large increase in Δobs or ϕ with declining PAR. However, there was a large increase in Δobs under −58‰ CO2 with declining PAR, indicating that B. sinuspersici used recent photoassimilate as a substrate for day respiration for both LL and ML plants under low PAR. It should be noted that leaves probably use a mixture of both recent and old photosynthate as substrates for day respiration. Therefore, our assumption of leaves exclusively using recent or old photosynthate is probably an over-simplification. This may explain why ML plants measured under −58‰ CO2 assuming recent photosynthate over-estimated ϕ and assuming old photosynthate under-estimated ϕ relative to plants measured under −7‰ CO2 (Fig. 4). In addition, two separate cohorts of plants were used for the −58‰ and −7‰ CO2 measurements; therefore, some of the difference in ϕ could also be attributed to differences between plants.
Anatomical and biochemical changes
Sage & McKown (2006) suggested that Kranz-type C4 plants are less phenotypically plastic in their response to light-limited growth conditions because of their unique anatomical requirements. However, Pengelly et al. (2010) demonstrated that the Kranz-type C4 plant Flaveria bidentis showed substantial plasticity when grown under LL, including decreased leaf thickness, smaller cells and increased SLA. In the single-cell C4 plant B. sinuspersici, there were more subtle effects on leaf anatomy for plants grown under LL vs ML. The characteristic structure of the C4 chlorenchyma cells was maintained under LL, and growth light conditions did not influence leaf thickness, which is probably related in part to the thick succulent nature of the leaves. The ratio of leaf area to dry mass (SLA) was higher in plants grown under LL conditions, suggesting higher leaf density (possibly associated with cell wall density and starch accumulation).
In Kranz-type C4 plants, it has been suggested that the mesophyll surface area next to the IAS correlates with C4 photosynthetic capacity (Evans & von Caemmerer, 1996). In LL plants, the mesophyll surface area exposed to IAS was greater than in ML plants; however, this relationship was not statistically significant. In addition, the distance from the CCC to the IAS was 30% greater in ML relative to LL plants (Table 3), but this relationship was not statistically significant. However, the ratio of the individual cell area to the area of the CCC (CA : CCC) was greater in LL relative to ML plants (Table 3). This increase in CA : CCC suggests a greater distance from the CCC to the IAS in LL plants, which might increase the resistance to leakage of CO2 from the CCC during C4 photosynthesis. Greater resistance to CO2 leakage could increase the photosynthetic efficiency of the single-cell CO2-concentrating mechanism (von Caemmerer, 2003); however, ϕ in LL plants was higher than in ML plants under all O2 and PAR conditions when correctly accounting for the influence of day respiration (Figs 3, 4).
This suggests that something other than leaf and cell anatomy is driving the differences in ϕ between LL and ML plants. It has been demonstrated that the cell structure, chlorophyll content and biochemical capacity of the leaf can also influence the efficiency of C4 photosynthesis. However, total chlorophyll was not significantly different between LL and ML plants. These findings are different from other studies with C4 plants, where ML plants showed lower total chlorophyll per unit leaf area relative to LL plants (Tazoe et al., 2008; Pengelly et al., 2010). In addition, the ratio of chlorophyll a/b was not significantly different in B. sinuspersici between growth treatments, but the ratio followed the same pattern as in Tazoe et al. (2008) and Pengelly et al. (2010), where high-light or ML plants had higher chlorophyll a/b ratios than LL plants.
With respect to biochemistry, the activities of Rubisco and PEPC were two and three times higher, respectively, in ML relative to LL plants (Table 2). Pengelly et al. (2010) also observed higher rates of Rubisco and PEPC activity in ML relative to LL F. bidentis. However, the ratio of PEPC/Rubisco activity in B. sinuspersici was not significantly different between treatments (Table 2). This suggests that changes in the relative capacity of the C3 and C4 cycles were not a major factor influencing the efficiency of the single-cell CO2-concentrating mechanism. However, modeling simulations have demonstrated that ϕ is highest at low photosynthetic capacity (low Vcmax and Vpmax; see von Caemmerer, 2003). Dry matter δ13C values (Table 2) were within the range of the values reported previously for chenopod species grown in environmental chambers (Voznesenskaya et al., 2002; Akhani et al., 2009; Leisner et al., 2010; King et al., 2012). However, LL plants had more negative dry matter δ13C relative to ML plants (Table 2). This was also observed in F. bidentis reported in Pengelly et al. (2010) and Amaranthus cruentus (Tazoe et al., 2008), suggesting that LL plants had greater ϕ under growth conditions, which was also observed in the online measurements of carbon isotope discrimination in B. sinuspersici.
There have been few studies on the effect of growth light levels on structural, biochemical and physiological features associated with photosynthesis. The goal of this research was to determine how growth under limited light affects leaf structure and photosynthetic efficiency in a unique single-cell C4 system. Similar to Kranz-type C4 plants, a functional C4 system was maintained in this single-cell C4 species. Plants grown under ML were more effective in capturing and assimilating CO2 delivered by the C4 cycle than were plants grown under LL, which was linked to biochemical rather than anatomical changes. The photosynthetic efficiency of the single-cell C4 system is insensitive to changes in measurement PAR when correctly accounting for differences in day respiration and photosynthetic discrimination. The results also indicate that this single-cell C4 plant uses recent photoassimilate as a substrate for day respiration. Together, these data demonstrate that the fully developed single-cell C4 system in B. sinuspersici is robust when grown under ML. Although, under natural growth conditions, this species is exposed to high-light environments in semi-arid deserts, it can acclimate to growth under LL conditions (c. 10% full sunlight).
We thank Drs A. Gandin and N. Ubierna for helpful discussions on modeling and estimating leakiness. We are also grateful to Drs E. Voznesenskaya and N. Koteyeva for helpful discussions on microscopy, plant propagation and growth. In addition, thanks are due to C. Cody for growth chamber maintenance. This research was supported in part by instrumentation obtained through a National Science Foundation (NSF) Major Research Instrumentation grant no. 0923562 (A.B.C.) and partly by the Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the US Department of Energy through grant DE-FG02-09ER16062 (A.B.C.), and NSF grant MCB 1146928 (G.E.E.).