Involvement of respiratory processes in the transient knockout of net CO2 uptake in Mimosa pudica upon heat stimulation


  • The first two authors contributed equally to this work.


Leaf photosynthesis of the sensitive plant Mimosa pudica displays a transient knockout in response to electrical signals induced by heat stimulation. This study aims at clarifying the underlying mechanisms, in particular, the involvement of respiration. To this end, leaf gas exchange and light reactions of photosynthesis were assessed under atmospheric conditions largely eliminating photorespiration by either elevated atmospheric CO2 or lowered O2 concentration (i.e. 2000 μmol mol−1 or 1%, respectively). In addition, leaf gas exchange was studied in the absence of light. Under darkness, heat stimulation caused a transient increase of respiratory CO2 release simultaneously with stomatal opening, hence reflecting direct involvement of respiratory stimulation in the drop of the net CO2 uptake rate. However, persistence of the transient decline in net CO2 uptake rate under illumination and elevated CO2 or 1% O2 makes it unlikely that photorespiration is the metabolic origin of the respiratory CO2 release.

In conclusion, the transient knockout of net CO2 uptake is at least partially attributed to an increased CO2 release through mitochondrial respiration as stimulated by electrical signals. Putative CO2 limitation of Rubisco due to decreased activity of carbonic anhydrase was ruled out as the photosynthesis effect was not prevented by elevated CO2.


Electrical excitability and signalling, frequently associated with rapid responses to environmental stimuli, are ubiquitous features of higher plants (Wildon et al. 1992; Stankovic & Davies 1997; Davies 2004). A plethora of physiological effects have been discovered recently to follow electrical signalling (Trebacz, Dziubinska & Krol 2006; Fromm & Lautner 2007), such as mechanically induced action potentials (AP) or wound-induced variation potentials (VP) (Sibaoka 1969; Pickard 1973; Stahlberg & Cosgrove 1992, 1994).

The present study is motivated by previous work on the sensitive plant Mimosa pudica (Koziolek et al. 2004), where heat-induced wounding caused a strong, transient decrease in net CO2 assimilation and operating efficiency of photosystem II (ΦPSII). While dark and light reactions of photosynthesis dropped in M. pudica, leaf transpiration rate initially increased rapidly before reaching a maximum after 150–200 s, and subsequently declined to a lowered level (Koziolek et al. 2004). The transitional and rapid transpiration response upon heat stimulation was interpreted as a hydro-passive stomatal opening movement caused by sudden loss of epidermal turgor (Kaiser & Grams 2006). Similarly, Lautner et al. (2005) analysed the effects of electrical signals on photosynthesis of Populus trichocarpa as a woody-plant species. They confirmed the electrical nature of the heat-induced wound signal, showing a signal with variation-potential character to spread via the phloem systemically within the plant. Interference of electrical signals with plant photosynthesis has now been demonstrated for a wide range of species, for example, from the green algae genus Chara, tobacco, maize and Venus flytrap or soybean (Hlavackova et al. 2006; Grams et al. 2007; Krupenina & Bulychev 2007; Pavlovic, Demko & Hudak 2010; Pavlovic 2012; Gallé et al. 2013).

Despite the recent progress that has been achieved in understanding heat-induced electrical and hydraulic signal transduction and their effects on photosynthesis and stomatal aperture, mechanisms underlying the phenomenon of the transient knockout of net CO2 uptake still require further clarification. In general, heat-induced electrical signalling was shown to increase respiratory CO2 loss (Dziubinska, Trebacz & Zawadzki 1989; Fromm, Hajirezaei & Wilke 1995; Filek & Koscielniak 1997). However, it remains obscure if a putative increase of respiratory CO2 loss is involved in the transient drop of the net CO2 uptake rate in M. pudica. Thus, corresponding assessments on respiratory metabolisms are still lacking up to date and are addressed in the present work. Moreover, it has been suggested that photosynthetic dark reactions are the primary target of interference of electrical signals with photosynthesis (Pavlovic et al. 2011), probably mediated through, for example, CO2 limitations of the Rubisco reaction upon cytosolic pH changes. Such may inhibit the activity of carbonic anhydrase or other reactions of the Calvin–Benson cycle (Grams et al. 2009; Pavlovic et al. 2011). Resulting from CO2 limitation of Rubisco, photorespiratory CO2 release may be stimulated in Mimosa and partially explain the transient reduction in net CO2 uptake. To identify a putative involvement of dark respiration and/or photorespiration, respiratory CO2 release of leaves upon heat-induced electrical signals were assessed under darkness. In addition, quenching of chlorophyll fluorescence and leaf gas exchange were quantified under experimentally reduced O2 or elevated CO2 concentration to diminish photorespiratory O2 fixation by Rubisco. Moreover, putative CO2 limitations of Rubisco were addressed under high CO2 supply.

Material and Methods

Plant material

Mimosa pudica plants were grown from seeds under standard greenhouse conditions at 25 °C and 75% relative humidity. Experiments were performed on a neighbouring pair of pinnae on fully developed leaves.

Leaf gas exchange

Net CO2 exchange rate (JCO2) and stomatal conductance for water vapour (gH2O) of attached leaves were assessed using a Li-Cor-6400 porometer (Li-Cor Inc., Lincoln, NE, USA) at ambient CO2 concentration (about 400 μmol mol−1), relative humidity of about 75%, cuvette temperature of about 25 °C and a photosynthetic photon flux density (PPFD) of about 170 μmol m−2 s−1. Pinna 2 was enclosed into the gas exchange chamber while the tip of pinna 1 was heat-stimulated for 1–2 s (Fig. 1). In the following experiments, the conditions were consecutively modified: CO2 concentration was raised to 2000 μmol mol−1 CO2, light was turned off (i.e. PPFD of 0 μmol m−2 s−1) and O2 concentration was reduced to 1%. The reduction of the O2 concentration down to 1% largely inhibits the oxygenase reaction of Rubisco, whereas mitochondrial respiration remains unaffected as the cytochrome c oxidase has a much lower affinity for O2 than Rubisco. Effects on net CO2 exchange and stomatal conductance were analysed.

Figure 1.

Experimental arrangement. The plant was heat-stimulated at the tip of leaf pinna 1. Electrical signals were recorded in pinna 1 (electrode A) and the neighbouring pinna 2 (electrode B). Gas exchange and chlorophyll fluorescence were recorded in the neighbouring pinna 2.

Chlorophyll fluorescence imaging

In parallel to leaf gas exchange measurements, chlorophyll fluorescence imaging was used to assess spatio-temporal variations of energy conversion at photosystem (PS) II by means of an imaging-PAM chlorophyll fluorometer (Heinz Walz GmbH, Effeltrich, Germany). The system allowed for non-invasive determination of PSII fluorescence parameters by the saturation pulse method (Schreiber & Bilger 1993). Blue light (470 nm) was applied for pulse-modulated measuring light, actinic illumination and saturation pulses. The imaging fluorometer was mounted directly on top of the porometer chamber. After insertion of a leaf into the chamber, the imaged area of pinna 2 was adapted for at least 30 min to the chamber conditions in darkness before minimal and maximal fluorescence (Fo and Fm, respectively) were determined by a single saturation light pulse (PPFD > 3500 μmol m−2 s−1). Subsequently, actinic light was turned on (PPFD of about 167 μmol m−2 s−1) and saturation pulses were employed by 30 s intervals (see Koziolek et al. 2004; Lautner et al. 2005). The neighbouring pinna was heat-stimulated when leaf gas exchange and chlorophyll fluorescence stabilized under illumination, typically 15–20 min after the actinic light was turned on. PSII operating efficiency (ΦPSII), fraction of ‘open’ PSII centres (qL), and non-photochemical quenching (NPQ) were calculated according to Baker (2008). As the imaging-PAM fluorometer was not equipped with a far-red light source, Fo′ was calculated according to Oxborough & Baker (1997).

Electrical measurements

For intracellular measurements of the membrane potential, two microelectrodes were filled with 100 mm KCl and inserted into mesophyll cells of a leaflet of pinna 1 (electrode A) and of pinna 2, respectively (electrode B, Fig. 1), using a micromanipulator and a long-distance working objective. The reference electrode was immersed together with the cut end of a neighbouring leaf pinna into artificial pond water (APW, composed of 1.0 mm NaCl, 0.1 mm KCl, 0.1 mm CaCl2 and 1.0 mm MES, adjusted with Tris to pH 5.5). The electrodes were connected to a differential amplifier (World Precision Instruments, Model Duo 773, Sarasota, FL, USA). Recordings were made between microelectrodes and the reference electrode. After inserting the microelectrodes into mesophyll cells of mature leaflets, the tip of pinna 1 was heat-stimulated through the flame of a lighter (c. 1000 °C) for 1–2 s. Plant response was monitored by a chart recorder.


Measurements under PPFD of about 167 μmol m−2 s−1, CO2 concentration of 400 μmol mol−1 and O2 concentration of 21% corresponded to growth chamber conditions (Fig. 2a, Table 1). The initial level of the net CO2 exchange rate (JCO2) was 4.1 ± 0.3 μmol m−2 s−1. 110 s after heat stimulation of the neighbouring pinna, JCO2 of the observed pinna started to decline transitionally towards nil and recovered to a new level of about 3.0 ± 0.4 μmol m−2 s−1 after about 600 s. Contrasting with the drop of JCO2, the stomatal conductance for water vapour (gs) increased during the first 220 s after heat stimulation from 70 ± 14 up to 174 ± 41 mmol m−2 s−1. A new level of 59 ± 11 mmol m−2 s−1 was reached after about 400 s. Under elevated CO2 supply (i.e. 2000 μmol mol−1), JCO2 was initially increased to 6.4 ± 0.2 μmol m−2 s−1 (Fig. 2b, Table 1). In general, similar dynamics were observed upon heat stimulation of the neighbouring pinna, with JCO2 reaching a minimum of 2.2 ± 0.5 μmol m−2 s−1 after about 200 s and recovering to 4.6 ± 0.6 μmol m−2 s−1 within subsequent 600 s. Under elevated CO2, the initial level of gs (45 ± 4 mmol m−2 s−1) was distinctly lower than under ambient CO2 concentration. Upon heat stimulation, gs increased intermittently to a maximum of 146 ± 22 mmol m−2 s−1 after about 200 s, eventually stabilizing at about 27 ± 5 mmol m−2 s−1 600 s later. Measurements under low (i.e. 1%) O2 concentration yielded results similar to those under elevated CO2 concentration (Fig. 2c, Table 1): The initial JCO2 of 6.7 ± 0.2 μmol m−2 s−1 rapidly declined after heat stimulation of the neighbouring pinna to a minimum of 1.3 ± 0.3 μmol m−2 s−1. After 800 s, steady state was not reached at a level of 3.6 ± 0.4 μmol m−2 s−1. Stomatal conductance (gs) was rather high with initially 198 ± 26 mmol m−2 s−1, increased intermittently to a maximum of 634 ± 47 mmol m−2 s−1 at 130 s after heat stimulation before recovering to a level of about 58 ± 6 mmol m−2 s−1 after 800 s (Fig. 2c, Table 1).

Figure 2.

Dynamics of leaf gas exchange (a–c) and chlorophyll fluorescence (d–f) upon heat stimulation under various CO2 and O2 concentrations: a/d = 400 μmol mol-1 CO2 and 21% O2, b/e = 2000 μmol mol-1 CO2 and 21% O2, c/f = 400 μmol mol-1 CO2 and 1% O2. Time zero indicates the time of heat stimulation of a neighbouring leaf pinna. a–c present net CO2 exchange (JCO2) and stomatal conductance for water vapour (gs). d–f present PSII operating efficiency (ΦPSII), fraction of ‘open’ PSII (qL) and non-photochemical quenching (NPQ). All parameters were assessed under a PPFD of about 167 μmol m2 s-1. Data are representative for n = 8.

Table 1. Levels of net CO2 exchange (JCO2), stomatal conductance for water vapour (gs), PSII operating efficiency (ΦPSII), fraction of ‘open’ PSII centres (qL) and non-photochemical quenching (NPQ) before (initial level) and after (final level) the heat stimulation of a neighbouring pinna. Parameters were assessed under a PPFD of about 167 μmol m−2s1 and three different combinations of CO2 and O2 concentrations: (1) 400 ppm CO2 and 21% O2 (2) 2000 ppm CO2 and 21% O2 and (3) 400 ppm CO2 and 1% O2. In the case of JCO2, ΦPSII, qL and NPQ a transient decrease was observed and minimum values are presented, whereas in the case of gs a transient increase was observed and maximum values are listed. Data represent means± one SE, n = 8
  JCO2 (μmol m2 s−1)gs (mmol m2 s−1)ΦPSIIqLNPQ

400 ppm CO2

21% O2

Initial level4.1 ± 0.370 ± 140.54 ± 0.020.45 ± 0.030.55 ± 0.04
Transient min/max0.1 ± 0.4174 ± 410.37 ± 0.020.22 ± 0.010.50 ± 0.02
Final level3.0 ± 0.459 ± 110.49 ± 0.020.47 ± 0.020.92 ± 0.10

2000 ppm CO2

21% O2

Initial level6.4 ± 0.245 ± 40.55 ± 0.000.50 ± 0.000.75 ± 0.02
Transient min/max2.2 ± 0.5146 ± 220.42 ± 0.010.25 ± 0.010.47 ± 0.03
Final level4.6 ± 0.627 ± 50.52 ± 0.010.51 ± 0.010.98 ± 0.05

400 ppm CO2

1% O2

Initial value6.7 ± 0.2198 ± 260.46 ± 0.010.46 ± 0.010.85 ± 0.11
Transient min/max1.3 ± 0.3634 ± 470.30 ± 0.020.21 ± 0.020.69 ± 0.10
Final level3.6 ± 0.3658 ± 60.38 ± 0.020.42 ± 0.021.32 ± 0.11

Similar to net CO2 exchange, parameters of chlorophyll fluorescence reflected transitional decreases in ΦPSII, qL and NPQ upon heat stimulation. Under ambient CO2 and O2 concentrations, qL decreased to a minimum of 0.22 ± 0.01 but recovered to levels similar to those prior to heating, that is, 0.45 ± 0.03. Similarly, ΦPSII was initially about 0.54 ± 0.02, dropped to 0.37 ± 0.02 300 s after heat stimulation and recovered eventually to 0.49 ± 0.02. NPQ decreased slightly from 0.55 ± 0.04 to 0.50 ± 0.02 before rising to enhanced 0.92 ± 0.10 (Fig. 2d, Table 1). Dynamics resembled those under elevated CO2 (i.e. 2000 μmol mol−1, Fig. 2e, Table 1), then reaching slightly higher levels as compared to those under ambient CO2. Reduction of O2 concentration to 1% increased initial levels of NPQ to 0.85 ± 0.11 (Fig. 2f, Table 1). 150 s after heat stimulation, however, NPQ intermittently decreased to 0.69 ± 0.10 before recovering to stable 1.32 ± 0.11. Dynamics of ΦPSII and qL changed in a similar way to those in previous experimental conditions, displaying about 0.46 each before dropping to 0.30 ± 0.02 and 0.21 ± 0.02 for ΦPSII and qL, respectively. The spatio-temporal dynamics of the transitional drop in ΦPSII is exemplified under 400 μmol mol−1 CO2 and 21% O2 in Fig. 3. Up to 160 s after the heat stimulation of the neighbouring leaf pinna, the assessed chlorophyll fluorescence parameters remained unaffected, including ΦPSII of about 0.6. The transitional minimum was reached at 280 s after heat stimulation when all leaflets mirrored a distinct drop in ΦPSII to about 0.2. Only small areas in the central part of the leaflets approached levels of 0.1. After 480 s, the leaf had recovered almost completely with only small areas at the margins of the leaflets displaying ΦPSII < 0.4.

Figure 3.

Spatio-temporal dynamics of PSII operating efficiency (ΦPSII) upon heat stimulation of a neighbouring leaf pinna under 400 μmol mol-1 CO2 and 21% O2. Times are given in relation to the heat stimulation of a neighbouring leaf pinna (at time zero). The imaged area (45 × 34 mm) covers the mid-part of the leaf. Changes in ΦPSII took about 200 s to become apparent. The decrease in ΦPSII is indicated by a false-colour shift from blue to yellow, equivalent to a lowering from 0.6 to about 0.2.

Under darkness, JCO2 and gs each displayed similar dynamics as compared to those under illumination (Fig. 4a, Table 2). Throughout the experiments, JCO2 was negative, that is, reflecting respiratory CO2 release at an initial rate of −0.9 ± 0.1 μmol m−2 s−1 under ambient CO2 and O2 conditions. 140 s after heat stimulation, JCO2 decreased to −2.9 ± 0.1 μmol m−2 s−1 before recovering to the initial level after 800 s. Conversely, the stomatal conductance (gs) increased intermittently, becoming highest at about 180 s after heat stimulation at 278 ± 22 mmol m−2 s−1, before recovering to its initial level of 61 mmol m−2 s−1. Dynamics of JCO2 and gs remained largely unaffected by the reduction of O2 concentration to 1% (Fig. 4b, Table 2). The initial JCO2 of −0.7 ± 0.1 μmol m−2 s−1 was followed by a minimum (−2.4 ± 0.2 μmol m−2 s−1) that occurred 120 s after heat stimulation, before approaching −1.0 ± 0.1 μmol m−2 s−1, thus similar to initial levels. In consistency with all other experiments, gs transitionally increased to maximum 320 ± 55.99 mmol m−2 s−1 at about after 160 s after heat stimulation. The eventually reached steady state of 31 ± 4 mmol m−2 s−1 was somewhat lower than under initial conditions.

Figure 4.

Dynamics of net CO2 exchange (JCO2) and stomatal conductance for water vapor (gs) upon heat stimulation in darkness under 21% (a) and 1% (b) O2 concentration. Time zero indicates the time of heat stimulation of a neighbouring leaf pinna. Data are representative for n = 4–8.

Table 2. Levels of net CO2 exchange (JCO2), stomatal conductance for water vapour (gs) before (initial level) and after (final level) the heat stimulation of a neighbouring pinna. Parameters were assessed during darkness under different CO2 and O2 concentrations: (1) 400 ppm CO2 and 21% and (2) 400 ppm CO2 and 1% O2. In the case of JCO2, a transient decrease was observed and minimum values are presented, whereas in the case of gs, a transient increase was observed and maximum values are listed. Data represent means ± one SE, n = 4 to 8
  JCO2 (μmol m−2s−1)gs (mmol m−2s−1)

400 ppm CO2

21% O2

Initial level−0.9 ± 0.161 ± 5
Transient min/max–2.9 ± 0.1278 ± 22
Final level–0.9 ± 0.161 ± 10

400 ppm CO2

1% O2

Initial value–0.7 ± 0.181 ± 18
Transient min/max–2.4 ± 0.2320 ± 56
Final level–1.0 ± 0.131 ± 4

Electrical signalling

The heat stimulation performed in the dark triggered electrical signals detected with both electrodes A and B with amplitudes ranging between −50 and −70 mV and velocities of 5–6 mm s−1 (Fig. 5a). Hence, amplitudes stayed similar to those recorded upon heat stimulation under illumination (Fig. 5b). Furthermore, sudden light exposure elicited action potentials with amplitudes of almost 60 mV at 4 to 5 min after stimulation (Fig. 5c). In contrast, sudden light extinction promptly induced gradual hyperpolarization on mesophyll plasma membranes of 20 mV (Fig. 5d).

Figure 5.

Electrical recording in pinna 2 in the dark (a) and under light conditions (b) after heat stimulation of a neighbouring pinna (pinna 1). Arrows denote the instant of injury at the tip of pinna 1. (c) After switching on the light, a typical action potential is induced in the leaf mesophyll. (d) In contrast, switching off the light induces a gradual hyperpolarization.


Numerous reports refer to the prominent sensitive plant M. pudica (Weintraub 1952; Sibaoka 1962; Allen 1969; Abe & Oda 1976; Samejima & Sibaoka 1983; Fromm 1991; Volkov et al. 2010), in which nearly every part of the plant body can perceive environmental stimuli and transmit them as electrical signals to the pulvini, the places of visible movements. Recent work showed heat-induced electrical signals to affect photosynthesis of Mimosa and other species performing rapid movements (Koziolek et al. 2004; Pavlovic et al. 2010, 2011) as well as in the absence of movements in, for example, poplar, soybean and maize (Lautner et al. 2005; Grams et al. 2009; Gallé et al. 2013). In Mimosa, membrane potential measurements, combined with assessment of chlorophyll fluorescence and leaf gas exchange, revealed flaming to evoke an electrical signal that travels rapidly into the neighbouring leaf pinna and transitionally eliminates net CO2 uptake (Koziolek et al. 2004; Kaiser & Grams 2006). A few seconds later, ΦPSII becomes reduced, this change spreading acropetally via the veins throughout the leaflets (Koziolek et al. 2004). To date, it is not definitely explained whether the transitional response of photosynthesis directly resulted from changes in membrane potentials and/or concomitant ion fluxes that may evoke downstream effects on respiratory or photosynthetic processes. In fact, recent studies on soybean demonstrate ion shifts on leaf chloroplasts as well as a decline of mesophyll conductance for CO2 diffusion as a result of heat-induced electrical signalling (Gallé et al. 2013).

The intermittent hydro-passive stomatal opening response that was previously demonstrated in Mimosa under illumination (Kaiser & Grams 2006) persisted in darkness. In parallel, CO2 release was distinctly stimulated. The enhanced release, however, did not result from the increase of stomatal aperture per se as the internal CO2 concentration was raised in parallel. Such outcome is consistent with recent findings in Venus flytrap where the dark respiration and internal CO2 concentration were enhanced in the absence of stomatal movements (Pavlovic et al. 2010, 2011). Hence, the transitional decline in the net CO2 uptake rate upon the arrival of electric signals appears, at least partially, to result from an increased CO2 release by respiratory processes. In the present work, leaf gas exchange and light reactions of photosynthesis were assessed under darkness and atmospheric conditions that largely eliminated photorespiration by either elevated CO2 or lowered O2 concentration (i.e. 2000 μmol mol−1 or 1%, respectively), based on the assumption that respiration rates varies little in the light and in the dark (Krömer 1995; Yin et al. 2011). Persistence of both the respiratory stimulation and the transitional decline in the net CO2 uptake rate under such conditions negates photorespiration as the metabolic origin of the CO2 release.

In parallel to the transitional drop in CO2 uptake, energy conversion at PSII was affected upon heat stimulation, proving involvement of declines in ΦPSII and in qL as well as of increase in non-photochemical quenching (NPQ; Niyazova & Bulychev 1990; Herde et al. 1999; Koziolek et al. 2004; Pavlovic et al. 2010). Nevertheless, it was hypothesized that suppression of CO2 availability or Calvin–Benson cycle activity may be the primary target of the photosynthetic inhibition. Subsequently, decreased ATP and NAPDH+ consumption might feedback on electron transport (Pavlovic et al. 2011). Likewise, Davies (2004) reported heat wounding in tomato to cause rapid changes in the Rubisco transcript. In maize, evidence was provided that electrical signalling affects photosynthesis via changes of cytosolic pH (Grams et al. 2009), presumably through pH-dependent enzymes such as the carbonic anhydrase. It is known that an important part of the regulation of mesophyll conductance to CO2 has metabolic origin, related to the activity of carbonic anhydrase (Flexas et al. 2008), even though its impact has been shown to be dependent on the plant species as well as on the type of carbonic anhydrase (for review, see Flexas et al. 2012). Hence, a decrease in mesophyll conductance through down-regulation of carbonic anhydrase upon arrival of electric signals may induce CO2 limitation of Rubisco with negative feedback on PSII energy conversion. Such a view was not supported, however, by our experiments: (1) an increase in photorespiration resulting from putative CO2 limitation of Rubisco was not observed; (2) enhancement of the CO2 concentration should have been conducive to overcoming CO2 limitation of Rubisco; however, experiments under high CO2 supply did not prevent the drop of the net CO2 uptake rate.

In conclusion, leaf photosynthesis of the sensitive plant M. pudica is highly responsive to electrical signals, induced for instance by heat stimulation of a neighbouring pinna. The transitional drop in leaf net CO2 uptake is unlikely to be caused by decrease in carbonic anhydrase activity and subsequent CO2 limitation of Rubisco, although concurrent effects of electric signals on Calvin–Benson cycle enzymes and PSII energy conversion cannot be ruled out at the present stage. Nevertheless, the transitional knockout of net CO2 uptake is partially, at least, attributed to increased CO2 release through mitochondrial respiration upon stimulation by electrical signals.


The authors like to thank Dr Markus Löw, University of Melbourne, for help with combining the Li-Cor-6400 and WALZ imaging-PAM instruments to assess leaf gas exchange and chlorophyll fluorescence parameters in parallel. S.L. was supported by a postdoctoral fellowship from the Center for a Sustainable University, Universität Hamburg.