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Vincent Chochois John P. Vogel Gregory J. Rebetzke Michelle Watt 《Plant physiology》2015,168(3):953-967
Seedling roots enable plant establishment. Their small phenotypes are measured routinely. Adult root systems are relevant to yield and efficiency, but phenotyping is challenging. Root length exceeds the volume of most pots. Field studies measure partial adult root systems through coring or use seedling roots as adult surrogates. Here, we phenotyped 79 diverse lines of the small grass model Brachypodium distachyon to adults in 50-cm-long tubes of soil with irrigation; a subset of 16 lines was droughted. Variation was large (total biomass, ×8; total root length [TRL], ×10; and root mass ratio, ×6), repeatable, and attributable to genetic factors (heritabilities ranged from approximately 50% for root growth to 82% for partitioning phenotypes). Lines were dissected into seed-borne tissues (stem and primary seminal axile roots) and stem-borne tissues (tillers and coleoptile and leaf node axile roots) plus branch roots. All lines developed one seminal root that varied, with branch roots, from 31% to 90% of TRL in the well-watered condition. With drought, 100% of TRL was seminal, regardless of line because nodal roots were almost always inhibited in drying topsoil. Irrigation stimulated nodal roots depending on genotype. Shoot size and tillers correlated positively with roots with irrigation, but partitioning depended on genotype and was plastic with drought. Adult root systems of B. distachyon have genetic variation to exploit to increase cereal yields through genes associated with partitioning among roots and their responsiveness to irrigation. Whole-plant phenotypes could enhance gain for droughted environments because root and shoot traits are coselected.Adult plant root systems are relevant to the size and efficiency of seed yield. They supply water and nutrients for the plant to acquire biomass, which is positively correlated to the harvest index (allocation to seed grain), and the stages of flowering and grain development. Modeling in wheat (Triticum aestivum) suggested that an extra 10 mm of water absorbed by such adult root systems during grain filling resulted in an increase of approximately 500 kg grain ha−1 (Manschadi et al., 2006). This was 25% above the average annual yield of wheat in rain-fed environments of Australia. This number was remarkably close to experimental data obtained in the field in Australia (Kirkegaard et al., 2007). Together, these modeling and field experiments have shown that adult root systems are critical for water absorption and grain yield in cereals, such as wheat, emphasizing the importance of characterizing adult root systems to identify phenotypes for productivity improvements.Most root phenotypes, however, have been described for seedling roots. Seedling roots are essential for plant establishment, and hence, the plant’s potential to set seed. For technical reasons, seedlings are more often screened than adult plants because of the ease of handling smaller plants and the high throughput. Seedling-stage phenotyping may also improve overall reproducibility of results because often, growth media are soil free. Seedling soil-free root phenotyping conditions are well suited to dissecting fine and sensitive mechanisms, such as lateral root initiation (Casimiro et al., 2003; Péret et al., 2009a, 2009b). A number of genes underlying root processes have been identified or characterized using seedlings, notably with the dicotyledonous models Arabidopsis (Arabidopsis thaliana; Mouchel et al., 2004; Fitz Gerald et al., 2006; Yokawa et al., 2013) and Medicago truncatula (Laffont et al., 2010) and the cereals maize (Zea mays; Hochholdinger et al., 2001) and rice (Oryza sativa; Inukai et al., 2005; Kitomi et al., 2008).Extrapolation from seedling to adult root systems presents major questions (Hochholdinger and Zimmermann, 2008; Chochois et al., 2012; Rich and Watt, 2013). Are phenotypes in seedling roots present in adult roots given developmental events associated with aging? Is expression of phenotypes correlated in seedling and adult roots if time compounds effects of growth rates and growth conditions on roots? Watt et al. (2013) showed in wheat seedlings that root traits in the laboratory and field correlated positively but that neither correlated with adult root traits in the field. Factors between seedling and adult roots seemed to be differences in developmental stage and the time that growing roots experience the environment.Seedling and adult root differences may be larger in grasses than dicotyledons. Grass root systems have two developmental components: seed-borne (seminal) roots, of which a number emerge at germination and continue to grow and branch throughout the plant life, and stem-borne (nodal or adventitious) roots, which emerge from around the three-leaf stage and continue to emerge, grow, and branch throughout the plant life. Phenotypes and traits of adult root systems of grasses, which include the major cereal crops wheat, rice, and maize, are difficult to predict in seedling screens and ideally identified from adult root systems first (Gamuyao et al., 2012).Phenotyping of adult roots is possible in the field using trenches (Maeght et al., 2013) or coring (Wasson et al., 2014). A portion of the root system is captured with these methods. Alternatively, entire adult root systems can be contained within pots dug into the ground before sowing. These need to be large; field wheat roots, for example, can reach depths greater than 1.5 m depending on genotype and environment. This method prevents root-root interactions that occur under normal field sowing of a plant canopy and is also a compromise.A solution to the problem of phenotyping adult cereal root systems is a model for monocotyledon grasses: Brachypodium distachyon. B. distachyon is a small-stature grass with a small genome that is fully sequenced (Vogel et al., 2010). It has molecular tools equivalent to those available in Arabidopsis (Draper et al., 2001; Brkljacic et al., 2011; Mur et al., 2011). The root system of B. distachyon reference line Bd21 is more similar to wheat than other model and crop grasses (Watt et al., 2009). It has a seed-borne primary seminal root (PSR) that emerges from the embryo at seed germination and multiple stem-borne coleoptile node axile roots (CNRs) and leaf node axile roots (LNRs), also known as crown roots or adventitious roots, that emerge at about three leaves through to grain development. Branch roots emerge from all root types. There are no known anatomical differences between root types of wheat and B. distachyon (Watt et al., 2009). In a recent study, we report postflowering root growth in B. distachyon line Bd21-3, showing that this model can be used to answer questions relevant to the adult root systems of grasses (Chochois et al., 2012).In this study, we used B. distachyon to identify adult plant phenotypes related to the partitioning among seed-borne and stem-borne shoots and roots for the genetic improvement of well-watered and droughted cereals (Fig. 1; Krassovsky, 1926; Navara et al., 1994), nitrogen, phosphorus (Tennant, 1976; Brady et al., 1995), oxygen (Wiengweera and Greenway, 2004), soil hardness (Acuna et al., 2007), and microorganisms (Sivasithamparam et al., 1978). Of note is the study by Krassovsky (1926), which was the first, to our knowledge, to show differences in function related to water. Krassovsky (1926) showed that seminal roots of wheat absorbed almost 2 times the water as nodal roots per unit dry weight but that nodal roots absorbed a more diluted nutrient solution than seminal roots. Krassovsky (1926) also showed by removing seminal or nodal roots as they emerged that “seminal roots serve the main stem, while nodal roots serve the tillers” (Krassovsky, 1926). Volkmar (1997) showed, more recently, in wheat that nodal and seminal roots may sense and respond to drought differently. In millet (Pennisetum glaucum) and sorghum (Sorghum bicolor), Rostamza et al. (2013) found that millet was able to grow nodal roots in a dryer soil than sorghum, possibly because of shoot and root vigor.Open in a separate windowFigure 1.B. distachyon plant scanned at the fourth leaf stage, with the root and shoot phenotypes studied indicated. Supplemental Table S1.
Open in a separate windowThe third reason for dissecting the different root types in this study was that they seem to have independent genetic regulation through major genes. Genes affecting specifically nodal root growth have been identified in maize (Hetz et al., 1996; Hochholdinger and Feix, 1998) and rice (Inukai et al., 2001, 2005; Liu et al., 2005, 2009; Zhao et al., 2009; Coudert et al., 2010; Gamuyao et al., 2012). Here, we also dissect branch (lateral) development on the seminal or nodal roots. Genes specific to branch roots have been identified in Arabidopsis (Casimiro et al., 2003; Péret et al., 2009a), rice (Hao and Ichii, 1999; Wang et al., 2006; Zheng et al., 2013), and maize (Hochholdinger and Feix, 1998; Hochholdinger et al., 2001; Woll et al., 2005).This study explored the hypothesis that adult root systems of B. distachyon contain genotypic variation that can be exploited through phenotyping and genotyping to increase cereal yields. A selection of 79 wild lines of B. distachyon from various parts of the Middle East (Fig. 2 shows the geographic origins of the lines) was phenotyped. They were selected for maximum genotypic diversity from 187 diploid lines analyzed with 43 simple sequence repeat markers (Vogel et al., 2009). We phenotyped shoots and mature root systems concurrently because B. distachyon is small enough to complete its life cycle in relatively small pots of soil with minimal influence of pot size compared with crops, such as wheat. We further phenotyped a subset of this population under irrigation (well watered) and drought to assess genotype response to water supply. By conducting whole-plant studies, we aimed to identify phenotypes that described partitioning among shoot and root components and within seed-borne and stem-borne roots. Phenotypes that have the potential to be beneficial to shoot and root components may speed up genetic gain in future.Open in a separate windowFigure 2.B. distachyon lines phenotyped in this study and their geographical origin. Capital letters in parentheses indicate the country of origin: Turkey (T), Spain (S), and Iraq (I; Vogel et al., 2009). a, Adi3, Adi7, Adi10, Adi12, Adi13, and Adi15; b, Bd21 and Bd21-3 are the reference lines of this study. Bd21 was the first sequenced line (Vogel et al., 2010) and root system (described in detail in Watt et al., 2009), and Bd21-3 is the most easily transformed line (Vogel and Hill, 2008) and parent of a T-DNA mutant population (Bragg et al., 2012); c, Gaz1, Gaz4, and Gaz7; d, Kah1, Kah2, and Kah3. e, Koz1, Koz3, and Koz5; f, Tek1 and Tek6; g, exact GPS coordinates are unknown for lines Men2 (S), Mur2 (S), Bd2.3 (I), Bd3-1 (I), and Abr1 (T). 相似文献
Phenotype | Abbreviation | Unit | Range of Variation | |
---|---|---|---|---|
All Experiments (79 Lines and 582 Plants) | Experiment 6 (36 Lines) | |||
Whole plant | ||||
TDW | TDW | Milligrams | 88.6–773.8 (×8.7) | 285.6–438 (×1.5) |
Shoot | ||||
SDW | SDW | Milligrams | 56.4–442.5 (×7.8) | 78.2–442.5 (×5.7) |
No. of tillers | TillerN | Count | 2.8–20.3 (×7.4) | 10–20.3 (×2) |
Total root system | ||||
TRL | TRL | Centimeters | 1,050–10,770 (×10.3) | 2,090–5,140 (×2.5) |
RDW | RDW | Milligrams | 28.9–312.17 (×10.8) | 62.2–179.1 (×2.9) |
Rootpc | Rootpc | Percentage (of TDW) | 20.5–60.6 (×3) | 20.5–44.3 (×2.2) |
R/S | R/S | Unitless ratio | 0.26–1.54 (×6) | 0.26–0.80 (×3.1) |
PSRs | ||||
Length (including branch roots) | PSRL | Centimeters | 549.1–4,024.6 (×7.3) | 716–2,984 (×4.2) |
PSRpc | PSRpc | Percentage (of TRL) | 14.9–94.1 (×6.3) | 31.3–72.3 (×2.3) |
No. of axile roots | PSRcount | Count | 1 | 1 |
Length of axile root | PSRsum | Centimeters | 17.45–52 (×3) | 17.45–30.3 (×1.7) |
Branch roots | PSRbranch | Centimeters · (centimeters of axile root)−1 | 19.9–109.3 (×5.5) | 29.3–104.3 (×3.6) |
CNRs | ||||
Length (including branch roots) | CNRL | Centimeters | 0–3,856.7 | 0–2,266.5 |
CNRpc | CNRpc | Percentage (of TRL) | 0–57.1 | 0–49.8 |
No. of axile roots | CNRcount | Count | 0–2 | 0–2 |
Cumulated length of axile roots | CNRsum | Centimeters | 0–113.9 | 0–47.87 |
Branch roots | CNRbranch | Centimeters · (centimeters of axile root)−1 | 0–77.8 | 0–77.8 |
LNRs | ||||
Length (including branch roots) | LNRL | Centimeters | 99.5–5,806.5 (×58.5) | 216.1–2,532.4 (×11.7) |
LNRpc | LNRpc | Percentage (of TRL) | 4.2–72.7 (×17.5) | 6–64.8 (×10.9) |
LNRcount | LNRcount | Count | 2–22.2 (×11.1) | 3.3–15.3 (×4.6) |
LNRsum | LNRsum | Centimeters | 25.9–485.5 | 48–232 (×4.8) |
Branch roots | LNRbranch | Centimeters · (centimeters of axile root)−1 | 2.1–25.4 (×12.1) | 3.2–15.9 (×5) |
3.
The C4 photosynthesis carbon-concentrating mechanism in maize (Zea mays) has two CO2 delivery pathways to the bundle sheath (BS; via malate or aspartate), and rates of phosphoglyceric acid reduction, starch synthesis, and phosphoenolpyruvate regeneration also vary between BS and mesophyll (M) cells. The theoretical partitioning of ATP supply between M and BS cells was derived for these metabolic activities from simulated profiles of light penetration across a leaf, with a potential 3-fold difference in the fraction of ATP produced in the BS relative to M (from 0.29 to 0.96). A steady-state metabolic model was tested using varying light quality to differentially stimulate M or BS photosystems. CO2 uptake, ATP production rate (JATP; derived with a low oxygen/chlorophyll fluorescence method), and carbon isotope discrimination were measured on plants under a low light intensity, which is considered to affect C4 operating efficiency. The light quality treatments did not change the empirical ATP cost of gross CO2 assimilation (JATP/GA). Using the metabolic model, measured JATP/GA was compared with the predicted ATP demand as metabolic functions were varied between M and BS. Transamination and the two decarboxylase systems (NADP-malic enzyme and phosphoenolpyruvate carboxykinase) were critical for matching ATP and reduced NADP demand in BS and M when light capture was varied under contrasting light qualities.Interest in the C4 pathway has been increased by the potential for enhancing crop productivity and maintaining yield stability in the face of global warming and population pressure (Friso et al., 2010; Zhu et al., 2010; Covshoff and Hibberd, 2012). Maize (Zea mays), a C4 plant of the NADP-malic enzyme (ME) subtype, is a leading grain production cereal (www.fao.org). C4 photosynthesis is a shared activity between mesophyll (M; abbreviations are listed in BS) cells, coupled to allow the operation of a biochemical carbon-concentrating mechanism (CCM). The CCM effectively minimizes photorespiration by increasing the CO2 concentration in the bundle sheath (CBS), where Rubisco is exclusively expressed. Since BS and M are connected by plasmodesmata, some CO2 retrodiffuses. The refixation of that escaping CO2 by the CCM increases the activity of the CCM and the total ATP demand (ATPBS + ATPM) for gross CO2 assimilation (GA; [ATPBS + ATPM]/GA), from a theoretical minimum of five ATPs (Furbank et al., 1990). Leakiness (Φ), the amount of CO2 retrodiffusing relative to phosphoenolpyruvate (PEP) carboxylation rate, is therefore a proxy for the coordination between the CCM and assimilatory activity (Henderson et al., 1992; Tazoe et al., 2008; Kromdijk et al., 2010; Ubierna et al., 2011; Bellasio and Griffiths, 2013).
Open in a separate windowRecently, the maize C4 subgroup has been shown to be complicated by the presence of two BS decarboxylation enzyme systems (NADP-ME and phosphoenolpyruvate carboxykinase [PEPCK]), presumably both acting as CO2 delivery pathways (via malate [MAL] and Asp, respectively; Furumoto et al., 1999, 2000; Wingler et al., 1999; Eprintsev et al., 2011; Furbank, 2011; Pick et al., 2011). There is also an extensive overlap between BS and M functions, since both cell types can synthesize starch (Spilatro and Preiss, 1987; Kanai and Edwards, 1999) and reduce phosphoglyceric acid (PGA; Majeran and van Wijk, 2009; see the overall scheme in Fig. 1). Additionally, energetic partitioning can also vary between cell types, since the total ATP produced (JATP) per CO2 fixed in GA (JATP/GA) may be produced in BS (mainly through cyclic electron flow [CEF] around PSI) or in M (mainly through linear electron flow [LEF]), depending on the light locally available in BS or M (Kramer and Evans, 2011; Yin and Struik, 2012). Furthermore, although all NADPH is produced in M, the only compartment operating linear electron transport and oxidizing water, some NADPH is exported to BS through MAL diffusion, to meet the reducing power demand therein (NADPHBS). To capture the complex C4 physiology, several models of C4 photosynthesis have been developed (Berry and Farquhar, 1978; Laisk and Edwards, 2000, 2009; von Caemmerer, 2000). The earlier approaches were developed into the von Caemmerer (2000) C4 model. In particular, the associated light-limited equations (referred to subsequently as the “C4 model”) are used to estimate the parameters needed to resolve the isotopic discrimination (Δ) model, widely employed to study Φ under low-light conditions (for review, see Ubierna et al., 2011). The C4 model partitions JATP into two fractions: (1) the ATP consumed by PEP carboxylase, and (2) the ATP consumed by the C3 activity (glyoxylate recycling, PGA reduction [PR], and ribulose 1,5-bisphosphate [RuBP] regeneration). These activities are located in M, BS, or both compartments (see the overall scheme in Fig. 1). However, the C4 model simplifies the spatial compartmentalization between BS and M, and in this paper, we now develop the energetic implications of the differential contribution of M and BS to C4 photosynthesis under different light regimes.Open in a separate windowFigure 1.Metabolic model of C4 assimilation, rates of reaction, and net fluxes between BS and M. The overall scheme reports the reactions of the CCM (Furbank, 2011), Rubisco carboxylation, the reactions of the RPP pathway, the synthesis of starch, respiration, and glyoxylate recycling reactions. The tables, with the corresponding enzyme names, show the actual reaction rates, expressed relative to GA (5.13 μmol m−2 s−1), per unit of substrate transformed. Rates were estimated by parameterizing the model equations (PAR = 125 μE m−2 s−1 (A = 3.96 μmol m−2 s−1; RLIGHT = 1.17 μmol m−2 s−1; JATP = 28.6 μmol m−2 s−1), the output of the C4 model (VC = 5.35 μmol m−2 s−1; VP = 5.89 μmol m−2 s−1; VO = 0.44 μmol m−2 s−1), and the output of the Δ model (Φ = 0.23) under three characteristic ratios of ATP partitionings. These were numbered 1, 2, and 3. Condition 1 corresponds to the lowest ATP available in BS (ATP partitioning similar to that under blue light; Fig. 4B), condition 2 corresponds to an intermediate ATP availability in BS (ATP partitioning equal to that under red light; Fig. 4B), and condition 3 corresponds to the highest ATP available in BS (ATP partitioning equal to that under green light; Fig. 4B). The inset shows net metabolite fluxes between M and BS in multiples of GA. The ATP demand in BS (ATPBS) and M (ATPM), the total NADPH demand (NADPHTOT), and the NADPHBS were also calculated in the same three relevant conditions. PYR, Pyruvic acid.Because of these anatomical, metabolic, and energetic complexities, C4 metabolism is highly sensitive to limiting light intensity (Bellasio and Griffiths, 2013) and, potentially, light quality (Evans et al., 2007). Light quality has a greater influence on C4 photosynthesis than on C3. Leaf pigments preferentially absorb the blue and red region of the spectra, and some wavelengths penetrate deeper into leaves. It was shown in C3 leaves that exposure to different wavelengths results in characteristic light penetration profiles, which, translated into different gradients in PSII yield, rates of ATP production, and assimilation (A) within the leaf (Terashima et al., 2009). In C4 leaves, because of the concentric anatomy, light reaches M cells before the deeper BS (Evans et al., 2007) and could alter the balance between light harvesting and energetic partitioning between BS and M.In this paper, we model the likely profiles of light penetration for specific wavelengths associated with red, green, and blue light within a maize M and BS leaf cross section and calculate the impact on potential ATP production for each cell type. We calculate the proportion of absorbed light (AB) for each wavelength, expressed as AB BS/M, the fraction of photons absorbed in BS relative to the photons absorbed in M, from which we derive JATPBS/JATPM, the fraction of ATP produced in BS relative to the ATP produced in M. Second, we developed a steady-state metabolic model (Fig. 1; von Caemmerer 2000), to capture the spatial separation between BS and M and partitions the ATP demand between BS and M cells in terms of PR, starch synthesis (SS), and PEP regeneration, so as to meet the ATP availability in each cell type (Evans et al., 2007). Third, photosynthetic characteristics (leaf-level ATP production rate, CO2 assimilation, stomatal conductance, and Φ derived from online carbon isotope discrimination [Δ]) were measured under red, green, and blue light, and red, green, and blue light in combination (RGB), using a decreasing photon flux density (from 500 to 50 μE m−2 s−1) to investigate the importance of metabolic plasticity under limiting light intensities.
Open in a separate windowFor instance, AB BS/M and JATPBS/JATPM were both lower under the blue light (wavelength 460 nm), which is rapidly extinguished within the M leaf profile, than under white light, confirming that light quality perturbs C4 energetics. In spite of this shift, when maize plants were exposed to different light qualities, there was no change in Φ, indicating that, at steady state, the coordination between CCM activity and Rubisco assimilation was retained (Ubierna et al., 2011; Sun et al., 2012). The modeled metabolic plasticity projected a window for ATP demand partitioning (ATPBS/ATPM), which matched the values for JATPBS/JATPM supply estimated under red, green, and blue light. We show that the plasticity of C4 metabolism, and in particular the possibility of shifting between MAL and Asp as a primary carboxylase product, was of pivotal importance in allowing the plasticity of ATP and NADPH demand. In conclusion, our study explains the extensive overlap between BS and M functions and the requirement for at least two decarboxylase systems in NADP-ME subtype plants such as maize, providing an explanation for empirical observations on the diversity of decarboxylase activities and PEP regeneration pathways (Rathnam, 1978; Chapman and Hatch, 1981; Wingler et al., 1999; Eprintsev et al., 2011; Furbank, 2011; Pick et al., 2011). 相似文献
Table I.
Variables and acronyms described in the textAbbreviation | Definition | Unit |
---|---|---|
A | Net assimilation | μmol m−2 s−1 |
AB | Absorbed light | |
AB BS/M | Partitioning of absorbed light | Dimensionless |
ATPBS | ATP demand in BS | μmol m−2 s−1 |
ATPM | ATP demand in M | μmol m−2 s−1 |
BS | Bundle sheath | |
CBS | CO2 concentration in BS | μmol mol−1 |
CCM | Carbon-concentrating mechanism | |
CEF | Cyclic electron flow | |
DHAP | Dihydroxyacetone phosphate | |
ETR | Electron transport rate | μmol m−2 s−1 |
GA | Gross assimilation (A + RLIGHT) | μmol m−2 s−1 |
gBS | Bundle sheath conductance to CO2, calculated by fitting JMOD to JATP | mol m2 s−1 |
IRGA | Infrared gas analyzer | |
JATP | Total ATP production rate | μmol m−2 s−1 |
JATPBS | ATP production rate in BS | μmol m−2 s−1 |
JATPM | ATP production rate in M | μmol m−2 s−1 |
JMOD | Modeled ATP production rate | μmol m−2 s−1 |
LEF | Linear electron flow | |
LCP | Light compensation point | |
M | Mesophyll | |
MAL | Malate | |
MDH | Malate dehydrogenase | |
MDHBS | Malate dehydrogenase reaction rate in BS | μmol m−2 s−1 |
MDHM | Malate dehydrogenase reaction rate in M | μmol m−2 s−1 |
ME | Malic enzyme | |
ME | Malic enzyme reaction rate | μmol m−2 s−1 |
NADPHBS | NADPH demand in BS | μmol m−2 s−1 |
NADPHTOT | Total NADPH demand | μmol m−2 s−1 |
OAA | Oxaloacetic acid | |
PAR | Photosynthetically active radiation | μE m−2 s−1 |
PEP | Phosphoenolpyruvate | |
PEPCK | Phosphoenolpyruvate carboxykinase | |
PEPCK | PEPCK reaction rate | μmol m−2 s−1 |
PGA | 3-Phosphoglyceric acid | |
PPDK | Pyruvate phosphate dikinase | |
PPDK | PPDK reaction rate | μmol m−2 s−1 |
PR | PGA reduction | |
PRBS | PR rate in BS | μmol m−2 s−1 |
PRM | PR rate in M | μmol m−2 s−1 |
RBS | Respiration in the light in BS | μmol m−2 s−1 |
RLIGHT | Respiration in the light | μmol m−2 s−1 |
RPP | Reductive pentose phosphate | |
RuBP | Ribulose-1,5-bisphosphate | |
RuP | Ribulose-5-phosphate | |
SS | Starch synthesis | |
SSBS | Starch synthesis rate in BS | μmol m−2 s−1 |
SSM | Starch synthesis rate in M | μmol m−2 s−1 |
SSTOT | Total starch synthesis rate | μmol m−2 s−1 |
T | Transamination rate | μmol m−2 s−1 |
VC | Rubisco carboxylation rate | μmol m−2 s−1 |
VO | Rubisco oxygenation rate | μmol m−2 s−1 |
VP | PEP carboxylation rate | μmol m−2 s−1 |
Y(II) | Yield of PSII | |
Δ | 13C isotopic discrimination | ‰ |
δ13C | 13C isotopic composition relative to Pee Dee Belemnite | ‰ |
Φ | Leakiness | Dimensionless |
Table II.
Steady-state equations for the metabolic model of C4 assimilationProcesses described by Equations 4 to 10 can be calculated directly from the measured data for A, RLIGHT, and the output of the von Caemmerer C4 model (VO, VP, and VC), while Equations 11 to 21 require prior allocation of SS, PR, and PEPCK. For simplicity, enzyme names in italics represent the enzyme reaction rate. For stoichiometric consistency, reaction rates are calculated as rates of substrate transformation.Process | Symbol | Reaction Rate | Equation | Localization | Notes |
---|---|---|---|---|---|
Gross assimilation | GA | (4) | GA and RLIGHT rates are expressed per CO2. | ||
RuP phosphorylation | – | (5) | BS | RuP phosphorylation supplies Rubisco carboxylating activity (VC) together with oxygenating activity (VO). | |
Total PR | PRTOT | (6) | BS and M | This equation calculates the total rate of PR on the basis of the PGA produced by Rubisco carboxylation (2VC), Rubisco oxygenation (VO), and glyoxylate recycling (0.5VO) and considers the PGA consumed by respiration; 1/3 is the stoichiometric conversion between respiration (expressed per CO2) and PR (expressed per triose). | |
Total NADPH demand | NADPHTOT | (7) | BS and M | PR consumes one NADPH per PGA; the total rate of PR is PRTOT (see note to Eq. 6); in glyoxylate regeneration (per glyoxylate), 0.5 NADH is produced by Gly decarboxylase, 0.5 NADH is consumed by hydroxypyruvate reductase, and one ferredoxin (equivalent to 0.5 NADPH) is consumed by Gln synthetase; in total, 0.5 NADPH is consumed per glyoxylate (equivalent to VO rate; Supplemental Table S1; Yoshimura et al., 2004). | |
DHAP entering RPP | – | (8) | BS | The DHAP entering the RPP pathway corresponds to the total PR rate minus the DHAP used for starch synthesis, which in this work is expressed per triose. | |
Total SS | SSTOT | (9) | BS and M | In this model, assimilation is entirely converted to starch; this assumption does not influence energetics, as starch synthesis has the same ATP demand as phloem-loaded Suc; in Equation 9, 1/3 converts the stoichiometry of A (expressed per CO2) to the stoichiometry of SS (expressed per triose). | |
Total PEP regeneration | – | (10) | BS and M | PEP regeneration rate equals PEP consumption rate VP at steady state; PEP can be regenerated either by PPDK (mainly in M but active also in BS) or by PEPCK in BS; in this study, PPDK activity was assumed to be zero in BS. | |
Total ATP demand | ATPBS + ATPM | (11) | BS and M | Equation 11 calculates the total ATP demand as the sum of ATP demand for PR (one ATP per PGA, corresponding to PR), RuBP regeneration (one ATP per RuP, corresponding to VC + VO), glyoxylate recycling (one ATP per glyoxylate, corresponding to VO), starch synthesis (0.5 ATP per triose, corresponding to SS), and PEP regeneration (one ATP per PEPCK catalytic event or two ATP per PPDK catalytic event); compared with the original formulation of the C4 model, Equation 11 separates the ATP demand for PEPCK and PPDK, includes the ATP demand for SS, and considers the PGA utilized by respiration, which does not need to be reduced (see Eq. 6). | |
ATP demand in BS | ATPBS | (12) | BS | The ATP demand in BS is brought about by PR (at the rate of PRBS), RuBP regeneration (at the rate of VC + VO), glyoxylate recycling (at the rate of VO), starch synthesis (0.5 ATP per triose), and PEPCK activity (one ATP per OAA; see note to Eq. 11). | |
ATP demand in M | ATPM | (13) | M | The ATP demand in M is brought about by PR (at the rate of PRM), SS, and PPDK (two ATPs per pyruvic acid; see note to Eq. 11). | |
NADPH demand in BS | NADPHBS | (14) | BS | The NADPH demand in BS is brought about by PR (one NADPH per PGA) and glyoxylate recycling, which consumes 0.5 NADPH per glyoxylate (corresponding to VO; see Supplemental Table S2). | |
NADPH supply to BS | – | MDHM | (15) | BS | All NADPH available in BS is produced in M and exported through the MAL shuttle because we have assumed that no linear electron transport (i.e. water oxidation) occurred in BS; for this reason, the NADPH supply to BS corresponds to the NADPH consumed to reduce OAA to MAL in M, the process responsible for NADPH export, and not to the rate of MAL decarboxylation in BS, which depends on T, PEPCK, and MDHBS (Eq. 19). |
MDH activity in M | MDHM | (16) | M | MDH activity supplies the NADPH demand in BS; Equation 16 was derived from Equations 14 and 15. | |
Transamination | T | (17) | BS and M | Equation 17 expresses that, at steady state, all OAA is either transaminated or reduced; since T bypasses the MDHM reaction, which is the reaction responsible for NADPH export to BS (see note to Eq. 15), T has the function of balancing NADPH supply and demand, which becomes apparent when Equations 15 and 17 are combined. | |
MDH | MDHBS | T − PEPCK | (18) | BS | MDH is assumed to operate a fast conversion at equilibrium; therefore, it is passively regulated by the substrate availability: the OAA that is not used by PEPCK is reduced to MAL by MDH; MDH may use NADH, since no NADPH-dependent reduction of OAA has been observed in maize (Kanai and Edwards, 1999) and it is likely mitochondrial (Rathnam, 1978; Chapman and Hatch, 1981); the NADH regeneration may be carried out by chloroplastic ME, which is reported to react both with NADP and NAD (Chapman and Hatch, 1981); however, the process may be more complicated (Eprintsev et al., 2011, and refs. therein); note that in this study, we assumed that cells are decompartmentalized while PEPCK rate was manipulated to increase between zero and a maximum rate in response to ATP availability (see “Minimum and Maximum BS Allocation” for details). |
ME | ME | MDHM + MDHBS | 19 | BS | Equation 19 expresses that the rate of MAL oxidation by ME corresponds to the rate of MAL produced by MDH activity in M plus the rate of MAL produced by MDH activity in BS. |
PPDK | PPDK | VP − PEPCK | 20 | M | The PEP regenerated by PEPCK in BS diffuses to M and reduces the requirement of PEP regenerated by PPDK in M. |
PR in M | PRM | PRTOT − PRBS | 21 | M | PR is a shared process between BS and M. |
4.
5.
6.
Banumathi Sankaran Shilah A. Bonnett Kinjal Shah Scott Gabriel Robert Reddy Paul Schimmel Dmitry A. Rodionov Valérie de Crécy-Lagard John D. Helmann Dirk Iwata-Reuyl Manal A. Swairjo 《Journal of bacteriology》2009,191(22):6936-6949
GTP cyclohydrolase I (GCYH-I) is an essential Zn2+-dependent enzyme that catalyzes the first step of the de novo folate biosynthetic pathway in bacteria and plants, the 7-deazapurine biosynthetic pathway in Bacteria and Archaea, and the biopterin pathway in mammals. We recently reported the discovery of a new prokaryotic-specific GCYH-I (GCYH-IB) that displays no sequence identity to the canonical enzyme and is present in ∼25% of bacteria, the majority of which lack the canonical GCYH-I (renamed GCYH-IA). Genomic and genetic analyses indicate that in those organisms possessing both enzymes, e.g., Bacillus subtilis, GCYH-IA and -IB are functionally redundant, but differentially expressed. Whereas GCYH-IA is constitutively expressed, GCYH-IB is expressed only under Zn2+-limiting conditions. These observations are consistent with the hypothesis that GCYH-IB functions to allow folate biosynthesis during Zn2+ starvation. Here, we present biochemical and structural data showing that bacterial GCYH-IB, like GCYH-IA, belongs to the tunneling-fold (T-fold) superfamily. However, the GCYH-IA and -IB enzymes exhibit significant differences in global structure and active-site architecture. While GCYH-IA is a unimodular, homodecameric, Zn2+-dependent enzyme, GCYH-IB is a bimodular, homotetrameric enzyme activated by a variety of divalent cations. The structure of GCYH-IB and the broad metal dependence exhibited by this enzyme further underscore the mechanistic plasticity that is emerging for the T-fold superfamily. Notably, while humans possess the canonical GCYH-IA enzyme, many clinically important human pathogens possess only the GCYH-IB enzyme, suggesting that this enzyme is a potential new molecular target for antibacterial development.The Zn2+-dependent enzyme GTP cyclohydrolase I (GCYH-I; EC 3.5.4.16) is the first enzyme of the de novo tetrahydrofolate (THF) biosynthesis pathway (Fig. (Fig.1)1) (38). THF is an essential cofactor in one-carbon transfer reactions in the synthesis of purines, thymidylate, pantothenate, glycine, serine, and methionine in all kingdoms of life (38), and formylmethionyl-tRNA in bacteria (7). Recently, it has also been shown that GCYH-I is required for the biosynthesis of the 7-deazaguanosine-modified tRNA nucleosides queuosine and archaeosine produced in Bacteria and Archaea (44), respectively, as well as the 7-deazaadenosine metabolites produced in some Streptomyces species (33). GCYH-I is encoded in Escherichia coli by the folE gene (28) and catalyzes the conversion of GTP to 7,8-dihydroneopterin triphosphate (55), a complex reaction that begins with hydrolytic opening of the purine ring at C-8 of GTP to generate an N-formyl intermediate, followed by deformylation and subsequent rearrangement and cyclization of the ribosyl moiety to generate the pterin ring in THF (Fig. (Fig.1).1). Notably, the enzyme is dependent on an essential active-site Zn2+ that serves to activate a water molecule for nucleophilic attack at C-8 in the first step of the reaction (2).Open in a separate windowFIG. 1.Reaction catalyzed by GCYH-I, and metabolic fate of 7,8-dihydroneopterin triphosphate.A homologous GCYH-I is found in mammals and other higher eukaryotes, where it catalyzes the first step of the biopterin (BH4) pathway (Fig. (Fig.1),1), an essential cofactor in the biosynthesis of tyrosine and neurotransmitters, such as serotonin and l-3,4-dihydroxyphenylalanine (3, 52). Recently, a distinct class of GCYH-I enzymes, GCYH-IB (encoded by the folE2 gene), was discovered in microbes (26% of sequenced Bacteria and most Archaea) (12), including several clinically important human pathogens, e.g., Neisseria and Staphylococcus species. Notably, GCYH-IB is absent in eukaryotes.The distribution of folE (gene product renamed GCYH-IA) and folE2 (GCYH-IB) in bacteria is diverse (12). The majority of organisms possess either a folE (65%; e.g., Escherichia coli) or a folE2 (14%; e.g., Neisseria gonorrhoeae) gene. A significant number (12%; e.g., B. subtilis) possess both genes (a subset of 50 bacterial species is shown in Table Table1),1), and 9% lack both genes, although members of the latter group are mainly intracellular or symbiotic bacteria that rely on external sources of folate. The majority of Archaea possess only a folE2 gene, and the encoded GCYH-IB appears to be necessary only for the biosynthesis of the modified tRNA nucleoside archaeosine (44) except in the few halophilic Archaea that are known to synthesize folates, such as Haloferax volcanii, where GCYH-IB is involved in both archaeosine and folate formation (13, 44).
TABLE 1.
Distribution and candidate Zur-dependent regulation of alternative GCYH-I genes in bacteriaaOpen in a separate windowaGenes that are preceded by candidate Zur binding sites.bZur-regulated cluster is on the virulence plasmid pLVPK.cExamples of organisms with no folE genes are in boldface type.dZn-dependent regulation of B. subtilis folE2 by Zur was experimentally verified (17).Expression of the Bacillus subtilis folE2 gene, yciA, is controlled by the Zn2+-dependent Zur repressor and is upregulated under Zn2+-limiting conditions (17). This led us to propose that the GCYH-IB family utilizes a metal other than Zn2+ to allow growth in Zn2+-limiting environments, a hypothesis strengthened by the observation that an archaeal ortholog from Methanocaldococcus jannaschii has recently been shown to be Fe2+ dependent (22). To test this hypothesis, we investigated the physiological role of GCYH-IB in B. subtilis, an organism that contains both isozymes, as well as the metal dependence of B. subtilis GCYH-IB in vitro. To gain a structural understanding of the metal dependence of GCYH-IB, we determined high-resolution crystal structures of Zn2+- and Mn2+-bound forms of the N. gonorrhoeae ortholog. Notably, although the GCYH-IA and -IB enzymes belong to the tunneling-fold (T-fold) superfamily, there are significant differences in their global and active-site architecture. These studies shed light on the physiological significance of the alternative folate biosynthesis isozymes in bacteria exposed to various metal environments, and offer a structural understanding of the differential metal dependence of GCYH-IA and -IB. 相似文献7.
8.
Jochen K. Lennerz Jonathan B. Hurov Lynn S. White Katherine T. Lewandowski Julie L. Prior G. James Planer Robert W. Gereau IV David Piwnica-Worms Robert E. Schmidt Helen Piwnica-Worms 《Molecular and cellular biology》2010,30(21):5043-5056
Par-1 is an evolutionarily conserved protein kinase required for polarity in worms, flies, frogs, and mammals. The mammalian Par-1 family consists of four members. Knockout studies of mice implicate Par-1b/MARK2/EMK in regulating fertility, immune homeostasis, learning, and memory as well as adiposity, insulin hypersensitivity, and glucose metabolism. Here, we report phenotypes of mice null for a second family member (Par-1a/MARK3/C-TAK1) that exhibit increased energy expenditure, reduced adiposity with unaltered glucose handling, and normal insulin sensitivity. Knockout mice were protected against high-fat diet-induced obesity and displayed attenuated weight gain, complete resistance to hepatic steatosis, and improved glucose handling with decreased insulin secretion. Overnight starvation led to complete hepatic glycogen depletion, associated hypoketotic hypoglycemia, increased hepatocellular autophagy, and increased glycogen synthase levels in Par-1a−/− but not in control or Par-1b−/− mice. The intercrossing of Par-1a−/− with Par-1b−/− mice revealed that at least one of the four alleles is necessary for embryonic survival. The severity of phenotypes followed a rank order, whereby the loss of one Par-1b allele in Par-1a−/− mice conveyed milder phenotypes than the loss of one Par-1a allele in Par-1b−/− mice. Thus, although Par-1a and Par-1b can compensate for one another during embryogenesis, their individual disruption gives rise to distinct metabolic phenotypes in adult mice.Cellular polarity is a fundamental principle in biology (6, 36, 62). The prototypical protein kinase originally identified as a regulator of polarity was termed partitioning defective (Par-1) due to early embryonic defects in Caenorhabditis elegans (52). Subsequent studies revealed that Par-1 is required for cellular polarity in worms, flies, frogs, and mammals (4, 17, 58, 63, 65, 71, 89). An integral role for Par-1 kinases in multiple signaling pathways has also been established, and although not formally addressed, multifunctionality for individual Par-1 family members is implied in reviews of the list of recognized upstream regulators and downstream substrates (Table (Table1).1). Interestingly, for many Par-1 substrates the phosphorylated residues generate 14-3-3 binding sites (25, 28, 37, 50, 59, 61, 68, 69, 78, 95, 101, 103). 14-3-3 binding in turn modulates both nuclear/cytoplasmic as well as cytoplasmic/membrane shuttling of target proteins, thus allowing Par-1 activity to establish intracellular spatial organization (15, 101). The phosphorylation of Par-1 itself promotes 14-3-3 binding, thereby regulating its subcellular localization (37, 59, 101).
Open in a separate windowaLKB1 also is known as Par-4; MARKK also is known as Ste20-like; (−), inhibitory/negative regulation has been shown; GPCR, G protein-coupled receptors. MARKK is highly homologous to TAO-1 (thousand-and-one amino acid kinase) (46).The mammalian Par-1 family contains four members (Table (Table2).2). Physiological functions of the Par-1b kinase have been studied using targeted gene knockout approaches in mice (9, 44). Two independently derived mouse lines null for Par-1b have implicated this protein kinase in diverse physiological processes, including fertility (9), immune system homeostasis (44), learning and memory (86), the positioning of nuclei in pancreatic beta cells (35, 38), and growth and metabolism (43).
TABLE 1.
Multifunctionality of Par-1 polarity kinase pathwaysaRegulator or substrate | Function | Reference(s) |
---|---|---|
Regulators (upstream function) | ||
LKB1 | Wnt signaling, Peutz-Jeghers syndrome, insulin signal transduction, pattern formation | 2, 63, 93 |
TAO1 | MEK3/p38 stress-responsive mitogen-activated protein kinase (MAPK) pathway | 46 |
MARKK | Nerve growth factor signaling in neurite development and differentiation | 98 |
aPKC | Ca2+/DAG-independent signal transduction, cell polarity, glucose metabolism | 14, 37, 40, 45, 59, 75, 95 |
nPKC/PKD | DAG-dependent, Ca2+-independent signal transduction (GPCR) | 101 |
PAR-3/PAR-6/aPKC | (−); regulates Par-1, assembly of microtubules, axon-dendrite specification | 19 |
GSK3β | (−); tau phosphorylation, Alzheimer''s dementia, energy metabolism, body patterning | 54, 97 |
Pim-1 oncogene | (−); G2/M checkpoint, effector of cytokine signaling and Jak/STAT(3/5) | 5 |
CaMKI | (−); Ca2+-dependent signal transduction, neuronal differentiation | 99 |
Substrates (downstream function) | ||
Cdc25C | Regulation of mitotic entry by activation of the cdc2-cyclin B complex | 25, 72, 78, 103 |
Class II HDAC | Control of gene expression and master regulator of subcellular trafficking | 28, 50 |
CRTC2/TORC2 | Gluconeogenesis regulator via LKB1/AMPK/TORC2 signaling, PPARγ1a coactivator | 49 |
Dlg/PSD-95 | Synaptogenesis and neuromuscular junction, tumor suppressor (102) | 104 |
Disheveled | Wnt signaling, translocation of Dsh from cytoplasmic vesicles to cortex | 73, 94 |
KSR1 | Regulation of the Ras-MAPK pathway | 68, 69 |
MAP2/4/TAU | Dynamic instability (67, 83) of microtubules, Alzheimer''s dementia (30) | 11, 31-33, 47, 70, 96 |
Mib/Notch | Mind bomb (Mib degradation and repression of Notch signaling results in neurogenesis) | 57, 74, 81 |
Par3/OSKAR/Lgl | Cytoplasmic protein segregation, cell polarity, and asymmetric cell division | 7, 10 |
Pkp2 | Desmosome assembly and organization; nuclear shuttling | 68, 69 |
PTPH1 | Linkage between Ser/Thr and Tyr phosphorylation-dependent signaling | 103 |
Rab11-FIP | Regulation of endocytosis (23), trafficking of E-cadherin (64) | 34 |
TABLE 2.
Terminology and localization of mammalian Par-1 family membersOpen in a separate windowaPar should not to be confused with protease-activated receptor 1 (PAR1 [29]); C-TAK1, Cdc twenty-five C-associated kinase 1; MARK, microtubule affinity regulating kinase; MARKL, MAP/microtubule affinity-regulating kinase-like 1.bBasolateral to a lesser degree than Par-1b (37).cHuman KP78 is asymmetrically localized to the apical surface of epithelial cells (76).dVariant that does not show asymmetric localization in epithelial cells when overexpressed (95).Beyond Par-1b, most information regarding the cell biological functions of the Par-1 kinases comes from studies of Par-1a. Specifically, Par-1a has been implicated in pancreatic (76) and hepatocarcinogenesis (51), as well as colorectal tumors (77), hippocampal function (100), CagA (Helicobacter pylori)-associated epithelial cell polarity disruption (82), and Peutz-Jeghers syndrome (48), although the latter association has been excluded recently (27). As a first step toward determining unique and redundant functions of Par-1 family members, mice disrupted for a second member of the family (Par-1a/MARK3/C-TAK1) were generated. We report that Par-1a−/− mice are viable and develop normally, and adult mice are hypermetabolic, have decreased white and brown adipose tissue mass, and unaltered glucose/insulin handling. However, when challenged by a high-fat diet (HFD), Par-1a−/− mice exhibit resistance to hepatic steatosis, resistance to glucose intolerance, and the delayed onset of obesity relative to that of control littermates. Strikingly, overnight starvation results in a complete depletion of glycogen and lipid stores along with an increase in autophagic vacuoles in the liver of Par-1a−/− but not Par-1b−/− mice. Correspondingly, Par-1a−/− mice develop hypoketotic hypoglycemia. These findings reveal unique metabolic functions of two Par-1 family members. 相似文献9.
C. P. A. de Haan R. Kivist? M. L. H?nninen 《Applied and environmental microbiology》2010,76(20):6942-6943
Cj0859c variants fspA1 and fspA2 from 669 human, poultry, and bovine Campylobacter jejuni strains were associated with certain hosts and multilocus sequence typing (MLST) types. Among the human and poultry strains, fspA1 was significantly (P < 0.001) more common than fspA2. FspA2 amino acid sequences were the most diverse and were often truncated.Campylobacter jejuni is the leading cause of bacterial gastroenteritis worldwide and responsible for more than 90% of Campylobacter infections (7). Case-control studies have identified consumption or handling of raw and undercooked poultry meat, drinking unpasteurized milk, and swimming in natural water sources as risk factors for acquiring domestic campylobacteriosis in Finland (7, 9). Multilocus sequence typing (MLST) has been employed to study the molecular epidemiology of Campylobacter (4) and can contribute to virulotyping when combined with known virulence factors (5). FspA proteins are small, acidic, flagellum-secreted nonflagellar proteins of C. jejuni that are encoded by Cj0859c, which is expressed by a σ28 promoter (8). Both FspA1 and FspA2 were shown to be immunogenic in mice and protected against disease after challenge with a homologous strain (1). However, FspA1 also protected against illness after challenge with a heterologous strain, whereas FspA2 failed to do the same at a significant level. Neither FspA1 nor FspA2 protected against colonization (1). On the other hand, FspA2 has been shown to induce apoptosis in INT407 cells, a feature not exhibited by FspA1 (8). Therefore, our aim was to study the distributions of fspA1 and fspA2 among MLST types of Finnish human, chicken, and bovine strains.In total, 367 human isolates, 183 chicken isolates, and 119 bovine isolates (n = 669) were included in the analyses (3). PCR primers for Cj0859c were used as described previously (8). Primer pgo6.13 (5′-TTGTTGCAGTTCCAGCATCGGT-3′) was designed to sequence fspA1. Fisher''s exact test or a chi-square test was used to assess the associations between sequence types (STs) and Cj0859c. The SignalP 3.0 server was used for prediction of signal peptides (2).The fspA1 and fspA2 variants were found in 62.6% and 37.4% of the strains, respectively. In 0.3% of the strains, neither isoform was found. Among the human and chicken strains, fspA1 was significantly more common, whereas fspA2 was significantly more frequent among the bovine isolates (Table (Table1).1). Among the MLST clonal complexes (CCs), fspA1 was associated with the ST-22, ST-45, ST-283, and ST-677 CCs and fspA2 was associated with the ST-21, ST-52, ST-61, ST-206, ST-692, and ST-1332 CCs and ST-58, ST-475, and ST-4001. Although strong CC associations of fspA1 and fspA2 were found, the ST-48 complex showed a heterogeneous distribution of fspA1 and fspA2. Most isolates carried fspA2, and ST-475 was associated with fspA2. On the contrary, ST-48 commonly carried fspA1 (Table (Table1).1). In our previous studies, ST-48 was found in human isolates only (6), while ST-475 was found in both human and bovine isolates (3, 6). The strict host associations and striking difference between fspA variants in human ST-48 isolates and human/bovine ST-475 isolates suggest that fspA could be important in host adaptation.
Open in a separate windowaIn 0.3% of the strains, neither isoform was found. NF, not found.bNA, not associated.A total of 28 isolates (representing 6 CCs and 13 STs) were sequenced for fspA1 and compared to reference strains NCTC 11168 and 81-176. All isolates in the ST-22 CC showed the same one-nucleotide (nt) difference with both NCTC 11168 and 81-176 strains, resulting in a Thr→Ala substitution in the predicted protein sequence (represented by isolate FB7437, GenBank accession number ; Fig. HQ104931Fig.1).1). Eight other isolates in different CCs showed a 2-nt difference (isolate 1970, GenBank accession number ; Fig. HQ104932Fig.1)1) compared to strains NCTC 11168 and 81-176, although this did not result in amino acid substitutions. All 28 isolates were predicted to encode a full-length FspA1 protein.Open in a separate windowFIG. 1.Comparison of FspA1 and FspA2 isoforms. FspA1 is represented by 81-176, FB7437, and 1970. FspA2 is represented by C. jejuni strains 76763 to 1960 (GenBank accession numbers ). Scale bar represents amino acid divergence.In total, 62 isolates (representing 7 CCs and 35 STs) were subjected to fspA2 sequence analysis. Although a 100% sequence similarity between different STs was found for isolates in the ST-21, ST-45, ST-48, ST-61, and ST-206 CCs, fspA2 was generally more heterogeneous than fspA1 and we found 13 predicted FspA2 amino acid sequence variants in total (Fig. HQ104933 to HQ104946(Fig.1).1). In several isolates with uncommon and often unassigned (UA) STs, the proteins were truncated (Fig. (Fig.1),1), with most mutations being ST specific. For example, all ST-58 isolates showed a 13-bp deletion (isolate 3074_2; Fig. Fig.1),1), resulting in a premature stop codon. Also, all ST-1332 CC isolates were predicted to have a premature stop codon by the addition of a nucleotide between nt 112 and nt 113 (isolate 1960; Fig. Fig.1),1), a feature shared with two isolates typed as ST-4002 (UA). A T68A substitution in ST-1960 (isolate T-73494) also resulted in a premature stop codon. Interestingly, ST-1959 and ST-4003 (represented by isolate 4129) both lacked one triplet (nt 235 to 237), resulting in a shorter FspA2 protein. SignalP analysis showed the probability of a signal peptide between nt 22 and 23 (ACA-AA [between the underlined nucleotides]). An A24C substitution in two other strains, represented by isolate 76580, of ST-693 and ST-993 could possibly result in a truncated FspA2 protein as well.In conclusion, our results showed that FspA1 and FspA2 showed host and MLST associations. The immunogenic FspA1 seems to be conserved among C. jejuni strains, in contrast to the heterogeneous apoptosis-inducing FspA2, of which many isoforms were truncated. FspA proteins could serve as virulence factors for C. jejuni, although their roles herein are not clear at this time. 相似文献
TABLE 1.
Percent distributions of fspA1 and fspA2 variants among 669 human, poultry, and bovine Campylobacter jejuni strains and their associations with hosts, STs, and CCsHost or ST complex/ST (no. of isolates) | % of strains witha: | P valueb | |
---|---|---|---|
fspA1 | fspA2 | ||
Host | |||
All (669) | 64.3 | 35.4 | |
Human (367) | 69.5 | 30.0 | <0.001 |
Poultry (183) | 79.2 | 20.8 | <0.001 |
Bovine (119) | 25.2 | 74.8 | <0.0001 |
ST complex and STs | |||
ST-21 complex (151) | 2.6 | 97.4 | <0.0001 |
ST-50 (76) | NF | 100 | <0.0001 |
ST-53 (19) | NF | 100 | <0.0001 |
ST-451 (9) | NF | 100 | <0.0001 |
ST-883 (11) | NF | 100 | <0.0001 |
ST-22 complex (22) | 100 | NF | <0.0001 |
ST-22 (11) | 100 | NF | <0.01 |
ST-1947 (9) | 100 | NF | 0.03 |
ST-45 complex (268) | 99.3 | 0.7 | <0.0001 |
ST-11 (7) | 100 | NF | NA |
ST-45 (173) | 99.4 | 0.6 | <0.0001 |
ST-137 (22) | 95.5 | 4.5 | 0.001 |
ST-230 (14) | 100 | NF | <0.0001 |
ST-48 complex (18) | 44.4 | 55.6 | NA |
ST-48 (7) | 100 | NF | NA |
ST-475 (8) | NF | 100 | <0.001 |
ST-52 complex (5) | NF | 100 | <0.01 |
ST-52 (4) | NF | 100 | 0.02 |
ST-61 complex (21) | NF | 100 | <0.0001 |
ST-61 (11) | NF | 100 | <0.0001 |
ST-618 (3) | NF | 100 | 0.04 |
ST-206 complex (5) | NF | 100 | <0.01 |
ST-283 complex (24) | 100 | NF | <0.0001 |
ST-267 (23) | 100 | NF | <0.0001 |
ST-677 complex (59) | 100 | NF | <0.0001 |
ST-677 (48) | 100 | NF | <0.0001 |
ST-794 (11) | 100 | NF | <0.001 |
ST-692 complex (3) | NF | 100 | 0.04 |
ST-1034 complex (5) | NF | 80 | NA |
ST-4001 (3) | NF | 100 | 0.04 |
ST-1287 complex/ST-945 (8) | 100 | NF | NA |
ST-1332 complex/ST-1332 (4) | NF | 100 | 0.02 |
Unassigned STs | |||
ST-58 (6) | NF | 100 | <0.01 |
ST-586 (6) | 100 | NF | NA |
10.
11.
Stomata control gaseous fluxes between the internal leaf air spaces and the external atmosphere and, therefore, play a pivotal role in regulating CO2 uptake for photosynthesis as well as water loss through transpiration. Guard cells, which flank the stomata, undergo adjustments in volume, resulting in changes in pore aperture. Stomatal opening is mediated by the complex regulation of ion transport and solute biosynthesis. Ion transport is exceptionally well understood, whereas our knowledge of guard cell metabolism remains limited, despite several decades of research. In this review, we evaluate the current literature on metabolism in guard cells, particularly the roles of starch, sucrose, and malate. We explore the possible origins of sucrose, including guard cell photosynthesis, and discuss new evidence that points to multiple processes and plasticity in guard cell metabolism that enable these cells to function effectively to maintain optimal stomatal aperture. We also discuss the new tools, techniques, and approaches available for further exploring and potentially manipulating guard cell metabolism to improve plant water use and productivity.Stomata are microscopic, adjustable pores on the leaf surface. The evolution of stomata more than 400 million years ago (Edwards et al., 1986, 1992, 1998) helped facilitate the adaptation of plants to a terrestrial environment, where water is typically a limiting resource. Each stoma is composed of two kidney- or dumbbell-shaped guard cells, whose volume changes to adjust pore aperture, allowing plants to simultaneously regulate CO2 uptake and water loss. This facilitation of gas exchange by stomatal opening is one of the most essential processes in plant photosynthesis and transpiration, affecting plant water use efficiency and agricultural crop yields (Lawson and Blatt, 2014).Plant physiologists have a long history of investigating the behavior of these fascinating structures, reaching back more than a century to the pioneering work of Sir Francis Darwin (Darwin, 1916) and the American botanist Francis Ernest Lloyd (Lloyd, 1908). Major contributions to stomatal research arose from inventing and improving equipment and methods for quantitatively measuring the effects of environmental factors on stomatal pore aperture. After Darwin’s work, it became clear that the stomatal aperture actively responds to changes in the environment and regulates leaf transpiration rates (Meidner, 1987). Over the past century, much has been learned about their structure, development, and physiology.Despite the anatomical simplicity of the stomatal valve, the surrounding guard cells are highly specialized. Guard cells are morphologically distinct from general epidermal cells and possess complex signal transduction networks, elevated membrane ion transport capacity, and modified metabolic pathways. These features allow rapid modulations in guard cell turgor in response to endogenous and environmental signals, promoting the opening and closure of the stomatal pore in time scales of seconds to hours (Assmann and Wang, 2001). A variety of osmotically active solutes contribute to the buildup of stomatal turgor. Potassium (K+) and chloride (Cl−) act as the main inorganic ions, and malate2− and sucrose (Suc) function as the main organic solutes. Whereas K+ and Cl− are taken up from the apoplast, Suc and malate2− can be imported or synthesized internally using carbon skeletons deriving from starch degradation and/or CO2 fixation in the guard cell chloroplast (Roelfsema and Hedrich, 2005; Vavasseur and Raghavendra, 2005; Lawson, 2009; Kollist et al., 2014). The accumulation of these osmotica lowers the water potential, promoting the inflow of water, the swelling of guard cells, and the opening of the stomatal pore. Most of the ions taken up, or synthesized by guard cells, are sequestered into the vacuole (Barbier-Brygoo et al., 2011). As a result, the guard cell vacuoles undergo dynamic changes in volume and structure, which are crucial for achieving the full amplitude of stomatal movements (Gao et al., 2005; Tanaka et al., 2007; Andrés et al., 2014). During stomatal closure, guard cells reduce their volume through the release of ions into the cell wall and the consequent efflux of water.The transport of osmolytes across the plasma and tonoplast guard cell membranes is energized by H+-ATPase activity, which generates a proton motive force by translocating H+ ions against their concentration gradient (Blatt, 1987a, 1987b; Thiel et al., 1992; Roelfsema and Hedrich, 2005; Gaxiola et al., 2007). After the pioneering work of Fischer demonstrated the importance of K+ uptake in stomatal opening (Fischer, 1968; Fischer and Hsiao, 1968), K+ transport became of central interest and has long been considered the essence of stomatal movement regulation. The development of the voltage clamp technique, along with the relative easy acquisition of knockout mutants and transgenics in the model plant Arabidopsis (Arabidopsis thaliana), helped to uncover the precise mechanism and function of K+ fluxes in guard cells. It is well established that changes in membrane potential in response to several stimuli (e.g. light/darkness, CO2, and abscisic acid [ABA]) alter the direction of K+ transport (Thiel et al., 1992; Blatt, 2000; Roelfsema et al., 2001, 2002, 2004). During stomatal opening, the activation of the proton pump generates a sufficiently negative electric potential to cause the uptake of K+ through the inward-rectifying K+ channels (K+in; Fig. 1). During stomatal closure, K+ outflow from outward-rectifying K+ channels (K+out) results from membrane depolarization (Fig. 2; Blatt, 1988; Schroeder, 1988; Anderson et al., 1992; Sentenac et al., 1992). Besides being gated by opposing changes in voltage, the activation of (K+out) channels is dependent on the extracellular K+ concentration, while that of K+in is not (Blatt, 1988, 1992; Roelfsema and Prins, 1997; Dreyer and Blatt, 2009). There is also strong evidence for H+-coupled K+ symport in guard cells, which could account for up to 50% of total K+ uptake during stomatal opening (Blatt and Clint, 1989; Clint and Blatt, 1989; Hills et al., 2012). At least for K+in, the loss of a single-channel gene in Arabidopsis has little or no impact on stomatal movement (Szyroki et al., 2001), showing the redundancy among the different K+in isoforms and of K+ transport in general.Open in a separate windowFigure 1.Integration of guard cell carbohydrate metabolism with membrane ion transport during stomatal opening. Sugars in guard cells can be imported from the apoplast, derive from starch breakdown, or be synthesized in the Calvin cycle. These sugars then can be stored as osmotically active solutes in the vacuole or metabolized in the cytosol to yield energy, reducing equivalents, and phosphoenolpyruvate (PEP). PEP can be further metabolized to pyruvate in the mitochondrial tricarboxylic acid (CAC) cycle or used as carbon skeletons for the biosynthesis of malate via PEP carboxylase (PEPC) and NAD-dependent malate dehydrogenase (NAD-MDH). Malate (which also can be imported from the apoplast) and the inorganic ions K+ and Cl− accumulate in the vacuole, lowering the guard cell osmotic potential, thereby promoting stomatal opening. ABCB14, ATP-binding cassette transporter B14; AcetylCoA, acetyl-CoA; ALMT, aluminum-activated malate transporter; ATP-PFK, ATP-dependent phosphofructokinase; AttDT, dicarboxylate transporter; cINV, cytosolic invertase; cwINV, cell wall invertase; Fru6P, Fru-6-P; Fru1,6P2, fructose 1,6-bisphosphate; Gl6P, Glc-6-P; G3P, glyceraldehyde 3-phosphate; iPGAM, phosphoglycerate mutase isoforms; NRGA1, negative regulator of guard cell ABA signaling1; OAA, oxaloacetate; 2-PGA, 2-phosphoglycerate; 3-PGA, 3-phosphoglycerate; PPi-PFK, PPi-dependent Fru-6-P phosphotransferase; STP, monosaccharide/H+ cotransporter; SUC, Suc/H+ cotransporter; SuSy, Suc synthase; TPT, triose phosphate/phosphate translocator. Compartments are not to scale. The dotted line indicates multiple metabolic steps.Open in a separate windowFigure 2.Proposed pathways of osmolyte dissipation during stomatal closure. While the removal of Cl− and K+ is well described in the literature, the fate of Suc and malate during stomatal closure is unclear. Suc can be cleaved by cytosolic invertase (cINV), and the resulting hexoses can be imported into the chloroplast in the form of Glc-6-P (Glc6P). Glc6P is used subsequently for starch biosynthesis. Malate can be removed from the cell via decarboxylation to pyruvate by malic enzyme (ME) and the subsequent complete oxidation in the mitochondrial tricarboxylic acid (CAC) cycle. Alternatively, malate can be converted to PEP via NAD+-dependent malate dehydrogenase (NAD-MDH) and PEP carboxykinase (PEPCK). Gluconeogenic conversion of PEP to Glc6P establishes a possible link between malate removal and starch synthesis. Compartments are not to scale. PEP, Phosphoenolpyruvate; OAA; oxaloacetate; STP, monosaccharide/H+ cotransporter; SUC, Suc/H+ cotransporter; SuSy, Suc synthase; cINV, cytosolic invertase; NRGA1, negative regulator of guard cell ABA signaling1; ALMT, aluminum-activated malate transporter; GPT, Glc-6-P/Pi translocator; cwINV, cell wall invertase; HK, hexokinase; QUAC1, quickly activating anion channel1.Despite the undisputed importance of K+ uptake in stomatal opening, the accumulation of K+ ions alone cannot account for the increase in osmotic pressure necessary to explain stomatal aperture. Studies from the 1980s by MacRobbie and Fischer demonstrated that Vicia faba guard cells take up approximately 2 pmol of K+ during stomatal opening. Assuming that K+ uptake is balanced by the accumulation of similar amounts of counter ions (Cl− and/or malate2−), the expected increase in stomatal turgor to approximately 3 MPa is less than the 4.5 MPa expected for fully open stomata (Fischer, 1972; MacRobbie and Lettau, 1980a, 1980b; Chen et al., 2012). The realization that other solutes must accumulate in addition to K+ salts was one of the major paradigm shifts in stomatal physiology research in the last decades, equal to the discovery of ion channels. Suc was put forward as the most likely candidate for the additional osmoticum to support stomatal opening (MacRobbie, 1987; Tallman and Zeiger, 1988; Talbott and Zeiger, 1993, 1998). Nonetheless, this research area subsequently failed to attract notice commensurate with its importance.In the last few years, the metabolism of starch, sugars and, organic acids in guard cells has seen a rebirth, making this the perfect time to review the developments in this field. In this review, we focus on photosynthetic carbon assimilation and respiratory metabolism in guard cells and provide a historical overview of the subject that highlights the most up-to-date and novel discoveries in guard cell research. We describe the various metabolic pathways separately, but as metabolism is an integrated network, we also discuss their reciprocal and beneficial interactions. Finally, we highlight their connection with the metabolism in the subjacent mesophyll cells and how they integrate with guard cell signal transduction networks and membrane ion transport to regulate stomatal movements. The enzymes and transporters discussed in this review are listed in Arabidopsis Genome Initiative Code Gene Protein Function Malate transport AT1G28010 ABCB14 ATP-binding cassette transporter B14 Import of apoplastic malate AT5G47560 tDT Dicarboxylate transporter Transport of carboxylates into the vacuole AT3G18440 ALMT9 Aluminum-activated malate transporter9 Transport of Cl−/malate2− into the vacuole AT2G17470 ALMT6 Aluminum-activated malate transporter6 Transport of malate2− into the vacuole AT4G17970 ALMT12/QUAC1 Aluminum-activated malate transporter12 Export of cytosolic Cl−/malate2− to the apoplast Malate metabolism – PEPC Phosphoenolpyruvate carboxylase β-Carboxylation of PEP to OAA – NAD-MDH NAD+-dependent malate dehydrogenase Reduction of OAA to malate – ME Malic enzyme Oxidative decarboxylation of malate to pyruvate AT4G37870 PEPCK1 PEP carboxykinase1 Conversion of OAA to PEP – PPDK Pyruvate, orthophosphate dikinase Conversion of pyruvate to PEP Other carboxylates – TPT Triose phosphate/phosphate translocator Export of triose phosphate from the chloroplast to the cytosol AT4G05590 NRGA1 Negative regulator of guard cell ABA signaling1 Putative mitochondrial pyruvate carrier – SDH2 Succinate dehydrogenase2 Oxidation of succinate to fumarate AT2G47510 FUM1 Fumarase1 Hydration of fumarate to malate – iPGAM Phosphoglycerate mutase Interconversion of 3-PGA to 2-PGA – PPi-PFK PPi-dependent Fru-6-P phosphotransferase Phosphorylation of Fru-6-P to Fru-1,6-bisphosphate – ATP-PFK ATP-dependent phosphofructokinase Phosphorylation of Fru-6-P to Fru-1,6-bisphosphate Calvin cycle – Rubisco Rubisco Carboxylation of ribulose 1,5-bisphosphate AT3G55800 SBPase Sedoheptulose-bisphosphatase Dephosphorylation of sedoheptulose-1,7-bisphosphate to sedoheptulose-7-phosphate Sugar metabolism AT4G29130 HK1 Hexokinase1 Phosphorylation of Glc to Glc-6-P AT4G02280 SuSy Suc synthase3 Interconversion of Suc to Fru and UDP-Glc – cINV Cytosolic invertase Hydrolysis of Suc to Fru and Glc – cwINV Cell wall invertase Hydrolysis of Suc to Fru and Glc Sugar transport AT1G11260 STP1 Monosaccharide/H+ cotransporter1 Import of apoplastic hexose sugars AT3G19930 STP4 Monosaccharide/H+ cotransporter4 Import of apoplastic hexose sugars AT1G71880 SUC1 Suc/H+ cotransporter1 Import of apoplastic Suc AT2G02860 SUC3 Suc/H+ cotransporter3 Import of apoplastic Suc Starch degradation AT3G23920 BAM1 β-Amylase1 Hydrolysis of α-1,4 external glucoside linkages in starch AT1G69830 AMY3 α-Amylase3 Hydrolysis of α-1,4 internal glucoside linkages in starch Starch synthesis – GPT Glc-6-P/Pi translocator Uptake of cytosolic Glc-6-P into the chloroplast AT4G24620 PGI Phosphoglucose isomerase Conversion of Fru-6-P to Glc-6-P AT5G51820 PGM1 Phosphoglucomutase1 Conversion of Glc-6-P to Glc-1-P AT5G48300 APS1 ADPGlc pyrophosphorylase small subunit Conversion of Glc-1-P to ADPGlc, catalytic subunit – APL ADPGlc pyrophosphorylase large subunit Conversion of Glc-1-P to ADPGlc, regulatory subunit Various AT3G45780 PHOT1 Phototropin1 Blue light photoreceptor AT5G58140 PHOT2 Phototropin2 Blue light photoreceptor AT4G14480 BLUS1 Blue light signaling1 Protein kinase, regulator of blue light-induced stomatal opening – PP1 Protein phosphatase1 Regulator of blue light-induced stomatal opening AT3G01500 CA1 Carbonic anhydrase1 Interconversion of CO2 and water into H2CO3 AT1G70410 CA4 Carbonic anhydrase4 Interconversion of CO2 and water into H2CO3 AT1G62400 HT1 High leaf temperature1 Protein kinase, regulator of CO2-induced stomatal closure