UNIVERSITY OF PISA
Department of Agricultural, Food and
Environment
MSc in Plant & Microbial Biotechnology
Effect of C/N and N source on photochemical PSII
efficiency, total soluble sugar and nitrate reductase
in Arabidopsis thaliana.
CANDIDATE: Andrea Ciurli SUPERVISOR: Lorenzo Guglielminetti
ASSISTANT SUPERVISOR: Lucia Guidi
Summary
Abstract………..4
1. Introduction 1.1 Overview of carbon to nitrogen ratio in plants………..…6
1.2 Signaling in C/N………9
1.3 Photosynthesis and C/N………...12
1.4 N assimilation and relations with C metabolism……….……16
2. Aim of the thesis………..……….21
3. Materials and methods 3.1 Plant material and growth conditions………..22
3.2 Soluble carbohydrate quantification………22
3.3 Chlorophyll fluorescence……….23
3.4 Nitrate reductase activity……….23
3.5 Nitrate reductase protein level……….24
3.6 Statistical analysis………24
4. Results 4.1 Total soluble sugars……….25
4.2 Photochemical efficiency of PSII………26
4.3 Nitrate reductase actual activity ……….27
5. Discussion and conclusion………..…….31
Aknowledgements………..………33
Abstract
Carbon (C) and nitrogen (N), among many others, are essential elements for plant metabolism and growth. The environment conditions such as CO2, light availability,
diurnal cycles, seasonal effect and biotic or abiotic stresses and many others, modulate the crosstalk named as ‘C/N. From previous experiments it is known that stressful C/N condition affects a large amount of aspects of plant metabolism. In the work from Huarancca Reyes et al. (2016) photochemical efficiency of PSII, phosphorylation status and localization of many enzymes, and total soluble sugar level resulted affected by unbalanced C/N ratio.
Since differences in C/N affect these parameters, in this project we checked if different sources of N had different effects when stressful high C/N ratio was imposed. In the previous work NH4NO3 was used as N source, and measurement of Chl a fluorescence,
total soluble sugar and enzymes activity were conducted on seedlings after 0,5; 2 and 24 hours after transfer in C/N medium. Here, NO3- and NH4+ were separately provided in
C/N medium. In our work, we investigated on photochemical efficiency of PSII, total soluble sugars level and nitrate reductase (NR) activity under stressful C/N condition (200 mM glucose/0,3 mM NO3- or NH4+) compared with control condition (100 mM
glucose/30 mM NO3- or NH4+). For total soluble sugars and nitrate reductase activity the
same time course of previous work was analysed, while Chl a fluorescence was conducted only 24 h after the treatment.
We found that treated plants accumulated more total soluble sugars (TSS) if compared with control and, in the long period (after 24 hours), TSS was higher with NH4+ in the
medium. Photochemical efficiency of PSII did not show significant differences between the two sources of nitrogen after 24 hours. Nitrate reductase actual activity was the result of comparison between activity, activation state and protein level. This activity constantly decreased starting from time zero in control condition; by contrast, NR total activity showed a peak at 2 hours after treatment with NO3- , and at 30 minutes with NH4+. This,
according with the total soluble sugar results, can be explained with the existence of a crosstalk between the sugar in excess and low nitrate in the medium, that blocks the activity of NR in stressful sugar condition until the plant is adapted to the stress. Further research should focus on the understanding the origin of this hypothetical crosstalk between excess of sugar and N-nutrient metabolism.
1. Introduction
1.1 Overview of Carbon to Nitrogen ratio in plants
Plants are autotrophic, photosynthetic, sessile, organisms that, during their evolution, colonized almost all the environments on the Earth. This has been possible thanks to the high level of adaptability of these organisms. Since plants cannot move, thus searching for better environments or conditions when they are under abiotic or biotic stresses, they adapted themselves to survive in a harsh environment by chemical compounds.
Plants need light, water, carbon (CO2) and other nutrients present in the soil to grow,
and their metabolism is finely regulated, to allow them to rapidly change their physiology and metabolic functions, according to environmental changes. For this aim, plants have developed a complex sensing and signalling mechanism to robustly monitor and appropriately respond to the dynamic changes of their surrounding environments.
Carbon (C) and nitrogen (N), among many others, are essential elements for plant metabolism and growth. The environment conditions such as CO2, light availability,
diurnal cycles, seasonal effect and biotic or abiotic stresses and many others, modulate the availability of carbon and nitrogen (Gibon et al., 2004; Klotke et al., 2004; Roitsch and Gonzàlez, 2004; Miller et al., 2007; Smith and Stitt, 2007). Carbon compounds include various carbohydrates, in particular mono- and disaccharides. These compounds are produced through photosynthesis that reduces CO2 into carbohydrates, and these
products provide both the energy and the C-skeletons for ammonium assimilation during amino acid biosynthesis. N nutrients include inorganic compounds, such as nitrate, and organic compounds, such as ammonium cation. Thus, for plants the relative concentrations of C and N need to be constantly monitored and regulated to have the right balance between energy and protein synthesis.
C and N metabolites are tightly coordinated and their ratio, through a crosstalk named as ‘C/N balance’ (Coruzzi and Zhou, 2001; Martin et al., 2002), is central for the regulation of plant growth and development. Coordination between C and N metabolites, therefore C/N ratio regulation, occurs at different levels (Zheng, 2009) (Fig. 1.1). CO2 is
assimilated through photosynthesis,
6CO2 + 6H2O C6H12O6 + 6O2 Sunlight energy
and the resulting carbohydrates, sucrose and starch, enter in the glycolysis and in the TCA, while nitrate (NO3-) is reduced to nitrite (NO2-) and then to ammonium (NH4+).
Figure 1.1. A simplified whole plant view of tightly coordinated C and N metabolism. C assimilation and N uptake occur in the leaf and the root systems, respectively. 2-oxoglutarate (2OG), an important intermediate product of C metabolism, serves as the C-skeleton for the synthesis of glutamate (which uses photorespiratory ammonium; not drawn here). Ammonium (NH4+) resulted from primary N assimilation
from nitrate (NO3-) is then incorporated to glutamate, and glutamine is synthesized. Other amino acids are
then synthesized by using NH4+ donated from glutamate and glutamine, and therefore proteins can be
synthesized. Proteins are essential for almost all cellular activities, including C and N metabolism. Source: Zheng 2009.
From 2-oxaglutarate in the TCA, are produced C skeletons for the synthesis of glutamate by incorporating NH4+. Another NH4+ from the primary N assimilation is then
incorporated to glutamate, resulting in the production of glutamine. Glutamine and glutamate donate the NH4+ for the synthesis of all the other amino acids. The amino acids
are required to build proteins, so it is clear that at this level, synthesis of amino acids, therefore proteins, is strongly related to C/N balance. Another level of this tight coordination involves the sensing and signaling of C/N balance through the long distance (Zheng, 2009). Nitrogen uptake occurs in the roots and N is assimilated as NO3- and/or
NH4+, depending on soil (or substrate) availability. Ammonium in the roots is converted
into amino acids, while nitrate can be reduced in ammonium then converted in amino acids too, or being transported into the leaves and then reduced and converted. Nitrate can be also stocked in the vacuole. Carbon assimilation and metabolism occur in the chloroplast by photosynthesis and strictly conditioned the N metabolism. Similarly, the production of photosynthetic carbohydrates is affected by N uptake. Therefore, to
modulate C and N metabolism, depending on each other availability, plants are able to sense the status of N in the root system and the surrounding root environment, and coordinate with the sensory machinery in the leaf where photosynthetic output will be determined (Zheng, 2009). In this view, it has been shown that the C/N balance, rather than carbohydrate or nitrogen status alone, plays a predominant role in regulating metabolites partitioning and various aspects of post-germination seedling growth (Martin et al., 2002).
A correct C/N balance in the tissues is important to be maintained by the plants, because it affects a large variety of processes, such as photosynthesis and, depending on this parameter, all the reactions using and producing energy must be finely regulated.
1.2 Signalling in C/N
The C/N balance sensing and signalling mechanisms have been largely unknown for years, but recently start to be clarified. In particular, the ubiquitin ligases ATL31 and ATL6 have beenreported to be involved in the C/N response during several development stages, such as post-germinative growth, developmental processes and defence response (Sato et al., 2009; Sato et al., 2011b; Maekawa et al., 2012, 2014; Aoyama et al., 2014; Huarancca Reyes et al., 2015; Maekawa et al., 2015). ATL31/6 are members of the ATL family, that is a large group of ubiquitin ligases with conserved domain structures common to both monocotyledons and dicotyledon plants (Salinas-Mondrago´n et al.,1999; Takai et al., 2001). Ubiquitin ligases are enzymes responsible to protein degradation by the 26S proteasome system. Detailed studies demonstrated that ATL31/6 target 14-3-3 proteins for ubiquitination in response to C/N-nutrient availability (Sato et al., 2011a; Yasuda et al., 2014) (Fig 1.2).
Figure 1.2. Proposed model of ATL31 and ATL6 function to regulate plant C/N response during post-germinative growth (A) ATL31/ATL6 binds and ubiquitinates to the 14-3-3 protein to promote protein
degradation via the 26S proteasome, which affects the activity of 14-3-3 client proteins and results in proper C/N response. Under high C/N stress conditions, over-accumulation of 14-3-3 proteins leads to growth arrest (red-coloured arrow). Under low C/N ratios, the 14-3-3 proteins are degraded and post-germination growth continues through the phase transition checkpoint (green-coloured arrow). On the other hand, another hypothesis was proposed (B): the14-3-3 proteins function as adaptor proteins to connect the ATL31/ATL6 and target protein X for ubiquitination to be degraded. In this model, the target protein X for ubiquitination is the client of 14-3-3 proteins, indicating that the ATL31/ATL6 directly regulates degradation of multiple proteins via the proteasome. Source: Sato et al 2011b.
The stability of 14-3-3 proteins increases as the C/N ratio changed from normal to high C/N stress in Arabidopsis seedlings, and an over-expression of 14-3-3 resulted in a hypersensitive C/N stress response. So, ATL31-mediated degradation of 14-3-3 is reduced under high-C/N stress (Sato et al., 2011; Sato et al., 2009). When an over-accumulation of 14-3-3 proteins occurs, plant growth results compromised or arrested. 14-3-3 proteins bind to phosphorylated motifs to regulate the activity of proteins involved in multiple developmental processes (Mackintosh, 2004) and allow their degradation when is needing. 14-3-3 proteomic analysis using barley, among the other analysis, revealed that several enzymes of carbohydrate metabolism such as sucrose synthase and invertases are targets of 14-3-3 proteins (Alexander and Morris, 2006). Sucrose synthase reversibly catalyses the synthesis and cleavage of sucrose, which is the main form of assimilated C, regulating sucrose flux and cellular transport and location depending on the metabolic environment to participate in cellulose, callose, and starch biosynthesis (Zheng et al., 2011; Tiessen and Padilla-Chacon, 2013). Invertase is another important enzyme that regulates the level of hexoses, which catalyses sucrose hydrolysis in different subcellular compartments regulating carbohydrate partitioning, developmental processes, hormone responses and biotic-abiotic interactions (Roitsch and Gonzalez, 2004; Tiessen and Padilla- Chacon, 2013). As it was previously known, both sucrose and glucose can initiate changes in gene regulation (Coruzzi & Zhou, 2001) that reflects on C/N sensing and signalling. Together with the ATL31/6 targeting 14-3-3 proteins mechanism, sugars are finally being recognized as important regulatory molecules with signalling functions in plants and other organisms. In general, source activities like photosynthesis, nutrient mobilization, and export are upregulated under low sugar conditions whereas sink activities like growth and storage are upregulated when carbon sources are abundantly available (Rolland et al., 2006). It is known that sugar-sensing mechanisms enable plants to turn off photosynthesis when C-skeleton is elevated due to the repression of photosynthetic gene transcription and ribulose-1,5bisphosphate carboxylase/oxygenase (Rubisco) activity (Krapp and Stitt, 1995; Cheng et al., 1998; Coruzzi and Zhou, 2001).
On this way, the connections between carbon metabolism, nitrogen assimilation and photosynthesis start to show a huge network made by signals and regulatory pathways. These connections allow plants to tune every nutrient that is needed, and they must be finely regulated one on each other, to keep on working.
1.3 Photosynthesis and C/N
Photosynthesis is regulated by feedback mechanisms, involving numerous signals that arise from carbon and nitrogen metabolism downstream of photosynthesis itself.C and N metabolisms are strictly depending one from each other, so C/N balance plays a key role on determining a feedback control of photosynthesis (Paul and Pellny, 2003).
Light energy absorbed by chlorophyll molecules can (i) drive photosynthesis (photochemistry); (ii) be re-emitted as heat; or (iii) be re-emitted as light (fluorescence) (Fig. 1.3.1). These three processes do not exist in isolation but rather in competition with each other. Thus, the yield of chlorophyll fluorescence emission gives us valuable information about the quantum efficiency of photochemistry and heat dissipation (Murchie and Lawson, 2013). Photochemistry is the absorption of photons by pigments (chlorophylls and others) and the conversion of radiative energy into chemical energy. Light harvesting complexes (LHCI and LHCII) have the aim to trap the light and transfer the excitation energy to the reaction centres of the photosystem II (PSII) and photosystem I (PSI), so they function as antenna and they are made by several chlorophyll molecules. Photosystems are located in the thylakoid membranes of the chloroplasts and they have many components that lead to the production of ATP and NADPH by splitting a molecule of water and transferring the two electrons derived.
Figure 1.3.1. A schematic figure showing electron transport within the PSII reaction centre complex. Energy absorbed by chlorophyll within the light-harvesting complex can be dissipated via photochemistry, by heat (non-photochemical quenching), or as fluorescence. The competition between these processes allows us to resolve the efficiency of PSII. Source: Murchie and Lawson, 2013.
When light is low, in a way, extremely efficient antenna systems absorb light and tunnel it through the reaction centres of photosystems, but when light is in excess (Figure 1.3.3), a large extent of this energy is dissipated, overall as heat, to prevent photo-damage (Guidi et al., 2017).
Figure 1.3.3 Absorbed and utilized energy in response to increasing light intensities. When light absorbed exceed photosystems requirement, the ‘excess energy’ can potentially cause photo-oxidative damage if it is not efficiently dissipated. Source: Guidi et al., 2017.
When green tissue is irradiated with photosynthetically active radiation (PAR) of approximately 400–700 nm, it emits radiation of longer wavelength in range from the red to far-red light, and this phenomenon is defined as ‘Chl a fluorescence’. Although Chl a fluorescence represents only a small fraction of the absorbed energy (from 0.5 to 10%) its intensity is inversely proportional to the fraction of energy used for photosynthesis (Kalaji et al., 2017). For this reason, Chl a fluorescence signal can be used as a probe for photosynthetic activity. Nowadays, Chl a fluorescence can be used to study several other aspects of photosynthesis, such as non-photochemical quenching (NPQ), depending on the experimental design (Kalaji et al. ,2017).
Quantum yield (Φ) for CO2 assimilation is an important parameter that expresses
Quantum yield is the result of photochemistry (or photochemical quenching), NPQ and fluorescence, that occur in competition, so that any increase in the quantum yield of one will result in a decrease in the quantum yield of the other two, according to the law of energy conservation:
ΦPSII + ΦF + ΦD = 1
where ΦPSII is the excitation energy that goes toward the photochemistry, ΦF is the energy
dissipated as fluorescence and ΦD is the amount of energy dissipated by heat. At low light
exposure and absence of stress conditions, ΦD andΦF should be closer to zero, since there
is not excess of light energy, thus photochemistry ΦPSII reaches the maximum efficiency.
When the light energy starts to saturate the photosynthetic apparatus, light energy in excess is safely dissipated through the mechanisms mentioned above and ΦPSII tends to
decrease. If the excess of light energy cannot be completely dissipated through NPQ and the other mechanisms, photo-oxidative damage may occur.
Using a Pulse Amplitude Modulation (PAM) fluorometer in vivo is possible to monitor the amount of fluorescence yield of a dark- or light adapted plant samples, thus the efficiency of photochemical and non-photochemical energy dissipation pathways (Fig. 1.3.4).
Figure 1.3.4. A stylized fluorescence trace of a typical experiment using dark-adapted leaf material to measure photochemical and non-photochemical parameters. Source: Murchie and Lawson, 2013.
Fluorescence measurement starts collecting the values of F0 (minimum yield of
fluorescence) in dark-adapted state; in this condition, all the PSII reaction centres are completely oxidised. The other parameter measured is Fm (maximum yield of
fluorescence in dark-adapted samples), obtained exposing dark-adapted samples to a saturating pulse (SP) for a short time, causing the reduction of the PSII reaction centres. Plants are then acclimated to actinic light to stabilize the photochemical reaction pathways and the parameter Ft, the fluorescence emitted in constant light condition, is
measured. Then, the parameter Fm’ (maximum yield of fluorescence in actinic light-adapted samples) is measured with an exposure to saturating pulse (SP). Generally, this parameter is lower than Fm, because of the stabilization of the photosynthesis. Finally, it is possible to calculate the ΦPSII, with the formula:
ΦPSII =
Fm’−FtFm’
The potential efficiency of PSII photochemistry can be calculated on dark-adapted leaves as Fv/Fm:
It has been shown theoretically and empirically that Fv/Fm gives a robust indicator of the maximum quantum yield of PSII chemistry (Butler, 1978; Genty et al., 1992). For unstressed leaves, the value of Fv/Fm is highly consistent, with values of ~0.83 (Demmig and Björkman, 1987).
Chl a fluorescence gives all the parameters needed to evaluate the physiologic state of PSII, therefore the fluctuation of photosynthesis rate. Therefore, by means of fluorescence measurement, in this experiment it will be shown how the energy photochemical efficiency of PSII is affected by changes in C/N balance and by the nitrogen sources.
1.4 N assimilation and relations with C metabolism
Plants take nutrients from the soil through their roots, and N is the fourth element in abundance in living organisms. Its availability in the soil is crucial for plant to grow, therefore for all the organisms in the ecosystem that create the food web. Nitrogen accounts for less than 0.1% in the soil, but in atmosphere it reaches about the 80%, in N2
form, and healthy plants typically contain up to 3%–4% N by dry weight. Biogeochemical cycle of nitrogen (Fig. 1.4.1) is very complex, due to the various oxidation states that the element can have.
Figure 1.4.1 Overview of nitrogen biogeochemical cycle. Source: Freeman et al., 2012; Environmental sciences.
Inorganic N is stocked mostly in igneous rocks, but this source is largely not usable for plants, while the major part of N absorbed by organisms is recycled from a pool of organic N-compounds, previously used by other organisms. Newly N is added to this pool by strong chemical reactions such as fire or lightings, or human activities. However, the highest input to this inorganic N-compounds pool derives from atmosphere, through a process called N fixation that converts N2 in NH3. Fixation is operated by prokaryotes
thanks to nitrogenase, an enzyme that catalyses the reduction of N2 to NH3. It can occur
thanks to symbiotic association of bacteria with plants, for instance the case of Fabaceae with Rhyzobium is perhaps the well-known, or free-living bacteria, such as Azotobacter among many others. After fixation process, atmospheric N is reduced to NH3 that in the
cytosol of nitrogen-fixing bacteria is present in the form of NH4+ (ammonium).
Ammonium is eventually released in the soil solution, where it can be absorbed by plants, bacteria or fungi, and be used to synthesize amino acids, or it can be oxidised by nitrificant bacteria to NO2- (nitrite) and then to NO3- (nitrate). NO3- can enter in metabolism of plants
by reduction to NH4+, and used to synthesize amino acids. Losses of nitrogen from this
cycle depend on the washout of nitrate in lower sites of the soil and on the denitrification process, caused by denitrificant bacteria (such as Pseudomonas spp.) that oxidate NO3- to
N2O and then to N2, that returns in atmosphere.
Plant cells can actively transport ammonium that is present in high concentration in acid soils or anoxic soils, where nitrification speed is low, as well as nitrate availability. However, ammonium is an essential ion for plants and agronomic fertilizations usually provide both nitrate and ammonium forms. There are two mechanisms by which ammonium can be taken up: a saturable high affinity transport and a non-saturable low affinity transport. The non-saturable low affinity transport is carried out by the AMT/Rh family genes that accounts for six genes in Arabidopsis thaliana. These genes can be found on root cells and root hair and they are site-specific, for instance on root hair there is AMT 1.5, on cortex cells AMT 1.2 and AMT 1.3. It is still unclear which transporters are involved in saturable high affinity transport. In this way, ammonium is taken up from the soil solution and transported toward xylem, but too much ammonium accumulated can be toxic for plants, therefore it is stocked in the vacuole to avoid the toxicity. Ammonium (both absorbed and synthesized) is then incorporated into amino acids via the glutamine synthetase-glutamate synthase (GS-GOGAT) pathway.
Nitrate is the major source of nitrogen for plants, and they spend a huge amount of their carbon and energy to N uptake. Usually, nitrate is assimilated through the epidermal and cortex cells of the root. Once in the cell, it can be stocked in the vacuole at high concentrations. Roots are the major sites of storage, but nitrate can be also loaded in the xylem to be transported to the shoots. There are two systems of transport, as well as in ammonium transport: high-affinity transport systems (HATS) operate at low external NO3- concentrations and display Michaelis–Menten kinetics saturating at »200 mM
(Glass & Siddiqi 1995) and they are largely responsible for root nitrate uptake; by contrast, low-affinity transport systems (LATS) make significant contribution to total nitrate uptake only at elevated concentrations (Siddiqi et al. 1990). In these mechanisms are involved several transporters, most of that belong to the NRT family. In this family, two subfamilies are contained: NRT1, that includes transporters with low and double
affinity, and NRT2, including high affinity transporters. These transporters are finely regulated by N availability, for instance in A. thaliana the transporter AtNRT1 is expressed in external cellular layer of the root, and its mRNA concentration is increased by nitrate treatment or sub-acid pH solutions, while AtNRT1:2 is constitutively expressed. Nevertheless, the whole mechanism is still largely unknown, and investigations about nitrate transporter are still in progress.
Nitrate must be reduced before that it can be incorporated in organic compounds, through a two phases process, and this reduction can occur either in the leaves or in the roots
1. NO3- is reduced to NO2- by nitrate reductase (NR)
2. NO2- is reduced to NH4+ by nitrite reductase (NiR)
Nitrate reductase is the key enzyme for reduction of nitrate to ammonium, since it catalyses the first step of nitrate assimilation. NR is a metal-enzyme complex, forming dimers and tetramers, where every subunit has specific sites for nitrate and for NAD(P)H and three domains binding three cofactors that provide redox centers: FAD, Fe-eme and cofactor MoCo (Fig. 1.4.2) Every part of the final dimer or tetramer has a partial activity. Two hinges connect every region of the monomer, and every hinge is a regulatory site, binding 14-3-3 protein.
Figura 1.4.2. Proposed model for NR structure in schematic representation a) and 3D representation b). Source: Buchanan, 2006
Nitrate reduction occurs in the cytosol of all the plant’s cells. NR is located mostly in roots and shoots, but its localization can be tuned and it is related to environment condition. Into a specific organ, NR has a cell-specific localization; at low external concentration of nitrate, NR is located mostly in epidermal cells and cortex cells of the
root, while at high concentration NR can be found also in vascular system. Through NR, NO3- is reduced to NO2-. Nitrite can be toxic in the cellular environment, so plants have
higher NiR activity, to avoid accumulations of nitrite, and regulation of NiR is strictly related with NR activity. NR appears to be the rate-limiting step in acquisition of nitrogen (Campbell, 1999; Tischner, 2000). NR is positively regulated by nitrate, light and carbohydrates and this regulation occurs at two levels: at transcription level, that allows long time regulation (days) and at post-transduction level, allowing short time regulation (minutes, hours). NR’s mRNA can be rapidly accumulated in response to one of these factors, despite the protein synthesis is generally slower. NR’s mRNA is substrate inducible, therefore nitrate plays a central role as a signal for NR transcription, as well as other environmental stimuli such as light, CO2, reduced carbon (particularly sucrose),
N-metabolites (particularly glutamine) and well-functioning chloroplasts in leaves. At post-transduction level, a serine residue in the hinge 1 (Fig 1.4.3) is phosphorylated through a Ca2+ dependent kinase followed by a Mg2+ or Ca2+ dependent bound of a 14-3-3 protein.
Figure 1.4.3. Schematic representation of kinase/phosphatase regulation of Nitrate Reductase by 14-3-3 protein mediated degradation. Source: Buchanan, 2006.
This mechanism allows a rapid and reversible inhibition of NR activity at unfavourable nitrate assimilation conditions (for example limiting light or CO2).
Between all the regulative factors of nitrate reductase, light plays a crucial role. Most of the energy for nitrate assimilation in a cell derives from photosynthesis, and it is also involved in the light regulation of NR activity and NR gene expression. When plants are exposed at high and stressful irradiance level, part of the energy in excess is diverted
toward nitrate reduction, by means of transferring reducing power to NR. In addition, light entrains the plant’s circadian rhythm, which has been proposed to influence the cyclic accumulation of NR transcript in anticipation of daylight, and corresponding decrease as night approaches (Lillo and Ruoff 1989, Deng et al. 1990).
2. Aim of the thesis
C/N balance must be optimal to allow plants to grow, but it rapidly changes depending on the environment. So, it is common that crop and wild plants must fight against stress related to imbalances in this ratio. Stressful conditions occur when carbon to nitrogen ratio is high, meaning low availability of nitrate and too many C metabolites that can produce stress.
From previous experiments it is known that stressful C/N condition affects a large amount of aspects of plant metabolism (Huarancca Reyes et al., 2016; Aoyama et al., 2014; Makeawa et al., 2012; Pompeiano et al., 2013; Sato et al., 2011a; Sato et al., 2011b; Yasuda et al., 2014). In the work from Huarancca Reyes et al., 2016, several physiological aspects resulted affected by unbalanced C/N ratio (200 mM glucose/0,3 mM N). Photochemical efficiency of PSII was different under C/N stressful condition depending on the timing if compared with time zero. It was shown that change of C/N balance can also modulate the phosphorylation status of many enzymes which may modify their activity, subcellular localization, stability or even signal transduction, as well as total soluble sugar level.
The aim of this project was to check if different sources of N had different effects on photochemical efficiency of PSII, soluble sugars level and nitrate reductase activity under stressful C/N condition.
In previous work NH4NO3 was used as N source, and measurement of fluorescence,
soluble sugar and enzymes activity were conducted on seedlings after 0,5; 2 and 24 hours after transfer in C/N medium. Here, since plant can assimilate N through different sources, NO3- and NH4+ were provided in C/N medium. The time course analysed was
the same for NR protein level, potential activity, activation state, and total soluble sugars. Conversely, Chl a fluorescence analysis was conducted only after 24 h of treatment, because the previous experiment showed that the decrease in ΦPSII occurred after two
hours, so we chose the final point of experiment as indicator of the decreased photochemical efficiency of PSII effect.
3. Material and methods
3.1 Plant material and growth conditions
Wild-type Arabidopsis thaliana Columbia-0 was used in this study. Sterilized seeds were sown on modified MS medium containing 100 mM glucose and 30 mM N (10 mM NH4NO3 and 10 mM KNO3) and growth for 10 days after germination at 16/8 h light/dark
(long day) photoperiod. Plants are then transferred to C/N mediums with different concentrations of glucose and N: 100 mM glucose and 30 mM N; 200 mM glucose and 0,3 mM N. As N source was used nitrate as KNO3 or ammonium as NH4Cl for each
medium. Each sample was growth in triplicate. Plants were harvested immediately, after 30 minutes, 2 hours and 24 hours after transfer in C/N medium.
3.2 Soluble carbohydrate quantification
Soluble carbohydrates were extracted from fresh tissues (50 mg), homogenized with 3 mL of 5.5% (v/v) HC1O4, placed for 60 minutes at 4°C then centrifuged at 12000g
for 5 minutes and the supernatant was neutralized with 3.5 M K2CO3 and placed on ice
for 60 min to allow further precipitation of the potassium perchlorate, which interfered with the coupled enzymatic assay of metabolites (Tobias et al. 1992). The neutralized extract was then centrifuged (12000g, 10 min), the supernatant is saved, the volume measured and stocked ad -20°C. Samples were assayed with coupled enzymatic assay methods (Pompeiano et al., 2013) measuring the increase in A340. Incubations of samples
and standards were carried out at 37°C for 30 min. The accuracy of the method was tested using standards with known amounts of carbohydrates. Recovery experiments were carried out to evaluate losses during extraction. Two tests were performed for each metabolite by adding known amount of authentic standards to the samples before proceeding with the extraction. The concentrations of standards added were similar to those estimated to be present in the tissues in preliminary experiments. The quantity of soluble carbohydrates was corrected on the basis of the recovery percentages for each sample, and expressed as moles hexose equivalents g-1 FW.
3.3 Chlorophyll fluorescence
Chlorophyll fluorescence measurements were conducted using a miniaturized pulse-amplitude-modulated fluorometer (Mini-PAM) (Heinz Walz GmbH, Effeltrich, Germany) on mono-layers leaf spot. The Photosynthetic Photon Flux Density (PPFD) of the saturation pulses to determine the maximal fluorescence emission in the presence (Fm’) and in the absence (Fm) of actinic light was about 8000 mol photons m-2 s-1.
Fluorescence parameters were determined at growth light intensity (100 mol photons m
-2 s-1) and at increasing PPFD (from 50 to 400 mol photons m-2 s-1) at the indicated times
after C/N treatment depending on the experiment. The potential efficiency of PSII photochemistry was calculated on dark adapted leaves as Fv/Fm. The actual photochemical PSII efficiency for photochemistry (ΦPSII) in the light was determined for
each PPFD value as ΦPSII = (Fm’-Ft)/Fm’ (Genty et al., 1989) when steady state was
achieved. Fm’ represents the maximum fluorescence yield with all PSII reaction centres in the reduced state obtained from superimposing a saturating light flash during exposition to actinic light, while F0 is the minimum yield of fluorescence in dark adapted samples. Fluorescence nomenclature is according to van Kooten and Snel (1990).
3.4 Nitrate reductase activity
Samples (0.2-0.5 g fresh weight) were extracted in 5 volume of 50 mM MOPS-NaOH, pH 7.5; 1 mM EDTA; 15 mM MgCl2; 2.5 mM DTT; 0,1% Triton X-100; 1 tab Phosphatase inhibitor PhosSTOP by Roche®. Extracts were centrifuged twice at 20.000g, 15 min, 4°C and the resulting supernatants were used for the enzymatic assays after desalting. Desalting column were equilibrated with the extraction buffer minus Triton. On resulting samples, Bio-Rad Protein Assay, based on the method of Bradford, was used to quantify protein concentration (Bradford M., 1976) and BSA were used as a standard. Reagents for enzymatic activity assay were prepared: A) 25 mM Potassium Phosphate Buffer with 10 mM Potassium Nitrate and 0.05 mM Ethylenediaminetetraacetic Acid, pH 7.3 at 30°C. Add 1 tablet PhosSTOP by Roche®. B) 2.0 mM ß-Nicotinamide Adenine Dinucleotide, Reduced Form Solution (ß-NADH). C) 3 M Hydrochloric Acid Solution (HCl). D) 58 mM Sulfanilamide Solution. E) 0.77 mM N-(1-Naphthyl) ethylenediamine Dihydrochloride Solution. F) 1.45 mM Nitrate Standard Solution. G) Samples. 5 nitrate standards were prepared, from 1:200 to 1:20 v/v in 25µL of B) and raise up with A).
Blank was prepared. Samples were mixed and equilibrated at 30°C. A volume of 1:20 v/v samples was added; Mixed and incubated at 30°C for few minutes; Added a half volume of D) to each tube. Then samples were mixed and the same volume of E) was added to stain, mixed and incubate at 25°C for 10 min. Absorbance at 540 nm was read.
3.5 Nitrate reductase protein level
After transferred in treated medium, plants were harvested, frozen in liquid nitrogen and immediately put in refrigerator at -80 °C. Plant material was then grinded in fine powder in liquid N and weighted. This time extraction was conducted without Phosphatase inhibitor. On resulting samples, Bio-Rad Protein Assay, based on the method of Bradford, was used to quantify protein concentration (Bradford M., 1976). Samples were prepared with protein concentration of 100µg in 60 µL of volume of E.B. (then 100 µg 120 µL in SDS 2X + E.B.). Concentration of protein were adjusted with extraction buffer, then the same amount of SDS 2X was added in the tube. Samples were incubated at 95°C for 5 minutes and kept in -30°C. 10 µL of samples were loaded for SDS-page. Western blot was carried out with anti-NR 1:2000 as primary antibody then polyclonal from rabbit 1:25000 as secondary antibody. Illuminance reaction with chemiluminescent HRP substrate was conducted. Coomassie blue dye was used to prepare stained gel.
3.6 Statistical analysis
Values presented are means ± standard error of three replicates. Data were subjected to analysis of variance (ANOVA), and the mean values were compared by using Tukey test. Significant differences for all statistical tests were evaluated at the level of P < 0.05.
4. Results
4.1 Total Soluble Sugars
Soluble sugars were extracted from Arabidopsis thaliana seedlings after transferring in C/N medium, and harvested at different times (30 minutes, 2 hours and 24 hours after treatment). In general, we found higher sugar concentration in treated samples if compared with control ones. In control conditions (30 mM N and 100 mM glucose) with NO3- (C NO3) total soluble sugars (TSS) maintained the same level of time zero
(T0) until the 2 h after treatment, then after 24 h the TSS level is slightly higher; with NH4+ in control condition (C NH4) TSS started increasing at 2h after treatment,
maintaining this trend in 24h, at which point, however, showed similar levels to C NO3. In treated condition (0,3 mM N and 200 mM glucose) with NO3- (T NO3) the TSS level
was clearly higher than T0 already from 30 minutes, and increasing until the 24 h; the same occurred with NH4+ (T NH4), starting with lower concentrations than T NO3, but
at 24h shows higher concentration than T NH4. (Fig. 4.1).
Figure 4.1 Effect of glucose and NH4+ or glucose and NO3- on total soluble sugars (TSS) in control condition
(30 mM N and 100 mM glucose) and treated condition (0,3 mM N and 200 mM glucose). Means ± standard error are shown. Letters indicate significant differences within treatment (p < 0.05) determined by analysis of variance (ANOVA), and the mean values were compared by using Tukey test.
treatment m ole s g FW -1 Sucrose Fructose Glucose e e e e d d c c cd dc cd b a
4.2 Chlorophyll fluorescence
The photochemical efficiency of PSII for photochemistry was measured between control and treatment conditions and the two source of N. Chl a fluorescence was measured at 0 PPFD (µmol m-2 s-1) (potential photochemical efficiency or Fv/Fm) and at increasing light 50, 100, 200 and 400 PPFD (actual photochemical efficiency of PSII or ΦPSII) after 24 hours of C/N treatment in control and treated condition (Fig. 4.2).
Figure 4.2. Light response curve of ΦPSII. Effect of high N levels and low glucose level (control), or low N
levels and high glucose level (treated) on photochemical efficiency of PSII at different light intensity after 24h of treatment. Results are compared by different N source in the same condition (T or C) (a,b) and by different condition per N source (b,c). ). Means ± standard error are shown. Asterisks indicate significant differences within treatment (p < 0.05) determined by by analysis of variance (ANOVA), followed by Tukey's test.
Between the two sources of nitrogen in the medium, nitrate or ammonium, there were not significant differences neither in control nor in treated plants (Fig 4.2 a,b). Comparing control with treated results for each N source, plants with both NO3- or NH4 in treated
C/N medium (T NO3 and T NH4), showed lower ΦPSII than the ones treated in control
C/N medium; with NH4+ in C/N medium, the difference starts at 100 PPFD, whereas with
NO3- difference start at 50 PPFD (Fig 4.2 b,c).
4.3 Nitrate reductase actual activity
Nitrate reductase (NR) is the key enzyme for nitrate reduction, therefore for nitrogen uptake. To check how stressful C/N condition affects the ability of Arabidopsis seedlings to assimilate nitrogen in presence of nitrate or ammonium, potential NR activity (assayed without phosphatase inhibitor) was investigated (Fig. 4.3.1).
Figure 4.3.1 Effect of high N levels and low glucose level (control), or low N levels and high glucose level (treated) on potential Nitrate Reductase activity (mU mg prot-1). Since the assay was conducted without
phosphatase inhibitors, these values represent the maximum possible of enzymatic activity. Means ± standard error were calculated from three independent experiment replications.
From time zero (T0), after 30 minutes of treatment in C/N medium the activity of NR remained the same for all the samples except for T NO3, that increased instead; after 2 hours of treatment all of the values decreased, except C NO3 that strongly increased; after 24 hours the activity decreased for all the samples with significant differences between the samples: both the treatment samples (T NO3 and T NH4) show the lower value, C NH4 that had a quite higher value and particularly C NO3 was strongly higher than all the other values.
We compared potential NR activity results with the NR protein level in the same treatment condition, doing western blot on the protein extracts. The results of western blot are shown in Fig. 4.3.2. At 30 minutes after treatment, while NR activity slightly increased and clearly increased in the case of C NO3 respect to T0, the actual protein
mU mg pro t-1
level decreased or remained constant. In C NO3 and T NO3 the protein level was significantly lower than T0, while it does not significantly change in C NH4 and T NH4. At 2 hours protein level returned the same of T0 for T NO3 and C NH4, but slightly decreased in C NO3 and clearly decreases in T NH4. At 24 hours, in C NO3 protein level was quite the same, while all the other decreased, particularly with ammonium in C/N medium.
Figure 4.3.2Effect of high N levels and low glucose level (control), or low N levels and high glucose level (treated) on Nitrate Reductase protein level with nitrate and ammonium in C/N medium. Results at 30 minutes (a), 2 hours (b) and 24 hours (c) after transfer in C/N medium are shown. Images of membrane from western blot were taken at 600 seconds (a), 60 seconds (b) and 250 seconds (c) of exposure after chemoilluminance reaction. Band intensity is relative to the higher average value of each graph. Means ± standard error were calculated from three independent experiment replications.
Nitrate reductase activation state can provide information about NR regulation, and compared with the above results can demonstrate as NR is regulated by post-transcriptional modifications (Fig. 4.3.3); in C NO3 the activation state significantly decreased compared to T0 form 2 hours after treatment, keeping this level at 24 hours; in T NO3 activation state at 30 minutes after treatment was strongly lower, but then increased again remaining the same of T0. C NH4 decreased if compared with T0 level, already from 30 minutes after treatment, remaining the same; the opposite occurred in T NH4, where activation state was the only one to be dramatically lower than T0.
Figure 4.3.3 Effect of high N and low glucose (C), or low N and high glucose (T) on Nitrate Reductase activation state (%) with nitrate and ammonium in C/N medium. Activation state is expressed as % of enzymatic activity with inhibitors by the one without inhibitor in each sample. Means ± standard error are shown. Letters indicate significant differences within treatment (p < 0.05) determined by analysis of variance (ANOVA), and the mean values were compared by using Tukey test.
Finally, we compared results of potential activity, activation state and protein level multiplying their values by each other in respect to 100 and calculated the actual activity of Nitrate Reductase in each condition. (Fig. 4.3.4). The result of this analysis showed how much nitrate reduction is occurring in each C/N condition, that is the sum of NR translation plus its real activity. These comparative results are shown as relative percentage to the higher activity found (the one found in T NH4 after 24 hours). NR total activity generally constantly decreased in respect to T0. In C NO3 this trend was
maintained until 24 hours; in T NO3 NR total activity strongly decreased at 30 minutes after treatment, but at 2 hours returned to a similar level of T0, then decreased again at 24 hours. In C NH4 the trend was similar to C NO3, but at 2 hours there was an increase; finally, in T NH4 at 30 minutes there was an increasing from T0, to rapidly decrease until 24 hours, at which point total activity had the minimum value.
Figure 4.3.4 Effect of high N and low glucose (C), or low N and high glucose (T) on Nitrate Reductase actual activity with nitrate and ammonium in C/N medium. Results are shown as relative percentage to the higher activity found (the one found in T NH4 after 24 hours) that is assumed as 100. Means ± standard error are shown. Letters indicate significant differences within treatment (p < 0.05) determined by analysis of variance (ANOVA), and the mean values were compared by using Tukey test.
0 20 40 60 80 100 120 C NO3 T NO3 C NH4 T NH4 T0 0,5 h 2 h 24 h
a
a
a
a
a
a
b
b b
b
c
c
c
c
c
d
5. Discussion and Conclusion
N nutrients trigger several metabolic pathways either depending on the availability of C and N and their relative ratio in the medium, or on the source availability in the substrate. In our results, in control conditions (30 mM N and 100 mM glucose), nitrate and ammonium did not show any strong difference in soluble sugar accumulation. By contrast, in treated condition (0.3 mM N and 200 mM glucose) it was observed that in the short period, nitrate in the medium leaded to higher accumulation of sugars, but in the long period was ammonium that leaded to higher sugar accumulation.
Photochemical efficiency of PSII was lower in high C/N condition after 24 h, independently by the nitrogen source. This indicates that under high C/N ratio plant are less tolerant to high light intensities. An excess of sugars, from the knowledge about sugar sensing, also negatively affect the photosynthetic rate. Moreover, photosynthesis was affected by nitrate availability. Nitrate reductase (NR) is a dissipation channel of energy in excess derived from photosynthesis, by means of transferring reducing power to NR to reduce nitrate (Lillo and Appenroth, 2001; Gniazdowska-Skoczek, 1998). So, when there is no nitrate in the medium NR cannot normally work, and photosynthesis is negatively affected because energy in excess cannot be dissipated through a convenient pathway. These factors agree with our results. Our data showed also that photosynthesis rate is not affected by the N source in high C/N condition.
Nitrate reductase is finely regulated in its activity at the different levels of post-transduction, through phosphorylation, and transcription level. Actual activity of NR is calculated by a comparison between potential activity, activation state and protein level, so from here we refer to NR activity meaning NR actual activity. Time zero (T0) seedlings were just transferred in the treatment medium in control and treated condition with NO3- or NH4+. At the T0 seedlings had high NR activity because most of all the N in
the sowing growth C/N medium (100 mM glucose and 30 mM NH4NO3) was consumed.
In control condition, both the N source were perceived by plants that modulate NR activity as N is assimilated. During the time, NO3- acted as a signal and constantly reduced
the NR activity; NH4+ had slower but similar effect, and this could be due to the missing
nitrate-signal, therefore a slower perception of NH4+ assimilation as feedback mechanism.
In treated condition with nitrate in the medium, in respect to T0, at 30 minutes after treatment, NR activity collapsed and this correlates with the high excess of accumulated
sugars, that seemed to inhibit nitrate assimilation. When the plant adapted to the stressful condition, NR activity had a peak probably due to the low availability of nitrate in the treated medium, then it strongly decreased; by contrast, with ammonium in the medium NR activity remained stable at 30 minutes, then decreased. This temporal shift between the peak of NR activity could be explained assuming that nitrate and sugar excess trigger a crosstalk that blocks every other cellular activity to allow plants to dispose sugar in excess. Therefore, when nitrate is missing, sugar excess could be not promptly sensed, resulting in higher accumulation and increasing inhibitory effect of sugars.
In conclusion, as a perspective of this work, further research should focus on the understanding the origin of this crosstalk between excess of sugar and N-nutrient metabolism, in order to understand the complex protein network in response to C/N-nutrient availability.
Acknowledgement
I would like to thank the Cell Structure and Function lab of Faculty of Science and Graduate School of Life Science (Hokkaido University, Japan), in particular Prof. Yamaguchi Junji and Dr. Sato Takeo to hosting me and take care of me during my work at this project, and during my daily life there. I am also very grateful to Yu Lu and Xingwen Li for helped me with my experiment and taught me so many things, Yoko Hasegawa for her friendship and support, and to all the members of the lab and wonderful people I met in Sapporo. I could never thank you enough.
I am greatly thankful to Lorenzo Guglielminetti, Lucia Guidi, Thais Huarannca Reys and Andrea Scartazza to have supervised my work and have been so patient with me.
I obviously cannot forget to thank all my professors from University of Pisa that, during the years, transmitted some of their passion and knowledge to me. I would like to mention Prof. Gianfranco Denti, that held my first university lecture and showed me what professionality and competence are.
Special thanks to all my friend and colleagues, particularly to Luca Marchetti, Daniel Seghieri, Alekos Garivalis, Lawyer Andrea Carpinelli, Jacopo Bettin, Federico Vittori, Gaia Morini, Gea Scarselli, my girlfriend Isotta Tonarelli, Marco Marino and Giovanni Macchi from my band ‘Acustica’ and Nicola Barghi. All of you are something I could never do without while reaching this goal. I am sorry I cannot mention all the people I would like, whoever spent some of their time with me in this period gave me something I could never forget. Thank you also to my parents Luigi Ciurli and Lucia Granito to give me the possibility to study and for many other things I could hardly explain here.
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