Abstract

The consequences of CO2‐concentrating in leaf air‐spaces of CAM plants during daytime organic acid decarboxylation in Phase III of CAM (crassulacean acid metabolism) are explored. There are mechanistic consequences of internal CO2 partial pressures, piCO2. These are (i) effects on stomata, i.e. high piCO2 eliciting stomatal closure in Phase III, (ii) regulation of malic acid remobilization from the vacuole, malate decarboxylation and refixation of CO2 via Rubisco (ribulose bisphosphate carboxylase/oxygenase), and (iii) internal signalling functions during the transitions between Phases II and III and III and IV, respectively, in the natural day/night cycle and in synchronizing the circadian clocks of individual leaf cells or leaf patches in the free‐running endogenous rhythmicity of CAM. There are ecophysiological consequences. Obvious beneficial ecophysiological consequences are (i) CO2‐acquisition, (ii) increased water‐use‐ efficiency, (iii) suppressed photorespiration, and (iv) reduced oxidative stress by over‐energization of the photosynthetic apparatus. However, the general potency of these beneficial effects may be questioned. There are also adverse ecophysiological consequences. These are (i) energetics, (ii) pH effects and (iii) Phase III oxidative stress. A major consequence of CO2‐concentrating in Phase III is O2‐concentrating, increased piCO2 is accompanied by increased piO2. Do reversible shifts of C3/CAM‐intermediate plants between the C3–CAM–C3 modes of photosynthesis indicate that C3‐photosynthesis provides better protection from irradiance stress? There are many open questions and CAM remains a curiosity.

Received 11 April 2002; Accepted 1 July 2002

The overt phenomenon of internal CO2‐concentrating: Phase III of CAM

Crassulacean acid metabolism (CAM) is a well‐known modification of photosynthesis that is covered in all the basic textbooks of plant physiology and biochemistry and which has been extensively reviewed (Osmond, 1978; Griffiths, 1989; Winter and Smith, 1996a; Cushman and Bohnert, 1997; Dodd et al., 2002). The essential mechanism of CAM is the acquisition of inorganic carbon (Ci) by dark‐fixation of bicarbonate (HCO3) via phosphoenolpyruvate carboxylase (PEPC). This leads to an organic acid (mainly malic acid)‐concentrating effect in the dark period when organic acid is stored in the central cell sap vacuole. In the subsequent light period organic acid is released from the vacuole again, decarboxylated and the resulting CO2 assimilated in the Calvin cycle. These parts of the CAM cycle are called Phase I (nocturnal CO2‐fixation via PEPC and acid accumulation) and Phase III (daytime CO2‐recovery and assimilation), respectively. These phases are separated by transition phases, Phase II in the morning and Phase IV in the afternoon, respectively (Osmond, 1978; see Fig. 1 in Dodd et al., 2002).

Nocturnally accumulated vacuolar organic acid is a CO2‐store in the form of carboxyl groups and not in the form of free Ci. Raven and Spicer (1996) have explained in detail why the storage of Ci equivalent to the large amounts of organic acid actually stored in CAM nocturnally (∼500 mol m–3) would be a less operable and favourable option. Thus, strictly speaking, Phase I is not considered as CO2‐concentrating, although it is an essential part of the whole mechanism.

Conversely, the release and decarboxylation of organic acid during the major part of the light period (Phase III) leads to a rapid build‐up of high internal CO2‐concentrations effecting stomatal closure in the light. The internal CO2‐concentrations behind closed stomata reported in the literature are compiled in Table 1. They range from a 2‐fold to an over 60‐fold increase of internal related to external CO2‐concentration.

Thus, when talking about CO2‐concentrating in CAM this refers to the Phase III phenomenon. This will be the topic of this review when the mechanistic consequences of CO2‐concentrating for leaf functions and the ecophysiological consequences for plant performance are assessed.

Historical recollections: CO2‐concentrating and O2‐concentrating

In the February of 1800 Alexander von Humboldt performed ecophysiological gas‐exchange measurements on Clusia rosea at Lake Valencia in Venezuela. It may be debated if he thus became the discoverer of CAM in Clusia. He certainly made the correct observations. Once again, in 1937, W Hartenburg also made the right observations, but like Alexander von Humboldt he did not present the appropriate explanation (Hartenburg, 1937; see Lüttge, 1995). Thus, the discovery of CAM in Clusia appears to be rightly acknowledged to Tinoco Ojanguren and Vazquez‐Janez (1983).

What, however, Alexander von Humboldt was certainly the first to discover is the high internal O2‐concentration in leaves of C. rosea, now known as a CAM plant (Ball et al., 1991a, b), during the light period. He writes

In keiner Pflanze zirkuliert vielleicht eine so ungeheure Menge an Luft als in der Clusia rosea. Wenn man die Blätter dem Sonnenlicht aussetzt, so ergeben sie ... nicht eine einzige Luftblase.’

‘In no other plant perhaps as much air is circulating than in the Clusia rosea. If one exposes the leaves to the sun light, they produce ... not a single air bubble.’

That means there is no obvious gas exchange in the light via the leaf surface. We now know, of course, that the stomata are closed in Phase III of CAM, and hence there is no gas exchange.

Aus dem verwundeten Teile des Stengels fährt aber mit ungeheurer Geschwindigkeit ein Strom von perlartigen Luftbläschen ... aus jedem dieser Gefäßbündel aus – ein herrliches Schauspiel’.

‘However, at the cut end of the petiole with an immense velocity a stream of pearl‐like gas bubbles ... is released from each of these vascular bundles, a magnificent spectacle.’

We now know that behind the closed stomata there is a high internal gas pressure in Phase III.

Setzte ich den Apparat in den Schatten, so hörte der Luftstrom auf...’

‘Did I place the apparatus in the shade, the gas stream ceased ...’

That means that Alexander von Humboldt performed the appropriate control.

Luft aus dem Innern der Clusia rosea ... besteht aus 0,35 Oxygen und 0,65 Azote’.

‘The air from the interior of Clusia rosea ... consists of 0,35 oxygen and 0,65 “azote”.’

That means that the gas in the leaf air spaces had 35% O2 and 65% N2.

As measurements in the field are always difficult, later the chemical O2‐determinations were checked in the laboratory of L‐J Gay‐Lussac in Paris and were found to be too high by a systematic error of 5% (Quotations after Krätz, 2001).

Using a gas chromatograph Spalding et al. (1979) rediscovered the high internal O2‐concentrations during the light period, i.e. in Phase III of CAM sensuOsmond (1978). They found values between 21.4% and 30.5% in various CAM species at midday and 41.5% in Kalanchoë gastonis‐bonnieri. In contrast to the high internal CO2‐concentrations in Phase III, reported by Spalding et al. (1979) and elsewhere (see below), the high internal O2‐concentrations have largely been forgotten until very recently, when the ecophysiological implications of this consequence of CO2‐concentrating began to be discussed in more depth.

Questions I

CO2‐concentrating mechanisms and internal CO2‐concentrations in the leaf air spaces of CAM plants were reviewed by Griffiths (1989) and in Winter and Smith (1996a; 19 entries for ‘carbon dioxide–intracellular space’ in the subject index). The major questions covered are the mechanism of CO2‐concentrating itself, and among the consequences the effects on photorespiration, light‐stress and photoinhibition, internal CO2‐recycling, water‐use‐efficiency, energy costs, and phylogenetic implications. These aspects are well, perhaps even overly, covered in the CAM literature. The present review will only allude to them insofar as it is necessary to fathom (i) some new mechanistic aspects of CAM‐functions, i.e. non‐linear dynamics of CAM‐leaves, including CO2‐signalling versus the established functions of CAM‐leaf metabolism, and (ii) some ideas on adverse (eco‐)physiological consequences versus the established beneficial (eco‐)physiological consequences of CAM.

Mechanistic consequences: spatiotemporal non‐linear dynamics of leaf functions

CO2‐remobilization from malic acid/malate in Phase III

Gas exchange measurements during most of Phase III when the stomata are firmly closed are not possible. The traditional method of recording malic acid remobilization is to measure malate levels or titratable acidity in the cell sap during the day. Rarely are samples taken at defined locations over the leaf. Thus, this destructive and tedious approach provides time series with a low temporal resolution and mostly no spatial resolution over the leaf. With the new technique of chlorophyll fluorescence imaging (Daley et al., 1989; Siebke and Weis, 1995) it is now possible (i) to obtain in addition a much better temporal resolution, (ii) to obtain spatial resolution over a leaf, and (iii) to obtain some more insight into the underlying processes, including the functions of CO2‐evolution behind closed stomata, rather than just recording the disappearance of malic acid.

Chlorophyll fluorescence imaging allows spatiotemporal monitoring of the relative quantum efficiency of photosystem II, ϕPSII. When internal CO2‐concentration, piCO2, is high it may saturate photosynthesis. This leads to high ϕPSII, and when ϕPSII is integrated over the whole leaf, ∫ϕPSII, high overall rates are obtained. The energy consumption of gluconeogenesis recovering the C3‐carbon skeleton, pyruvate or PEP, originating from malate decarboxylation, tends to lower ϕPSII. This effect is overridden by high piCO2 saturating photosynthesis, but it may be discernible when piCO2 is lower. Thus, chlorophyll fluorescence images show some of the consequences of internal CO2‐generation from malate decarboxylation and CO2‐concentrating.

These events are not homogeneous in space over a CAM leaf of K. daigremontiana during the light period (Rascher, 2001; Rascher et al., 2001). In the morning, some cells or patches of cells may start earlier than others to release malic acid from the vacuoles, to decarboxylate malate and to raise piCO2. Since ϕPSII reflects the local availability of CO2 this can be seen as a patchiness of ϕPSII. This inhomogeneity is particularly expressed during the transition between Phase II and Phase III. In phase II, stomata are still open and CO2 is both taken up from the atmosphere and produced in the leaf interior from malate decarboxylation (Borland and Griffiths, 1996).

Increasing piCO2 is known to be the major internal control parameter eliciting stomatal closure in Phase III. In K. daigremontiana stomata respond with a lag of about 15 min with closing movements as piCO2 builds up (Bohn et al., 2001). Local gradients of piCO2 established due to the desynchronization of leaf patches are maintained for a while, because there is a severe constraint on CO2‐diffusion inside the leaves of Kalanchoë daigremontiana. The uniformly spherical cells are densely packed, the internal air space is only about 4–9% of the leaf volume and diffusion resistance is large (Maxwell et al., 1997; Rascher et al., 2001). Patchiness of ϕPSII is almost entirely lost in Phase III when high internal piCO2 builds up (Table 1) overriding the limitations of CO2 diffusion. In Phase III there is no CO2 limitation anywhere in the leaf. However, inhomogeneity begins to peak again at the end of Phase III, as some patches have finished malate decarboxylation earlier than others. At this time not only patches but also waves of ϕPSII moving over the leaf are seen. Such waves may be explained by a complex interaction of malate depletion, CO2 diffusion from neighbouring leaf zones into the depleted zones and the energy budget determined by decreasing needs of gluconeogenesis (Rascher, 2001).

Hence, this non‐linear spatiotemporal performance of the leaves during Phases II to IV from the beginning to the end of the day reveals several CO2‐concentrating consequences. These observations elicit reflections on the regulation of CO2‐remobilization from nocturnally stored malic acid/malate.

Regulation of CO2‐remobilization from malic acid/malate in Phase III

Is anything known about how and where CO2‐remobilization from malic acid/malate is regulated in Phase III (Fig. 1)? Are there (and if so where are they) control points of metabolite fluxes in membrane‐transport and enzyme‐reaction steps? Is there any mass‐action type feedback?

(i) CO2‐consumption by Rubisco: Is it the rate of CO2‐consumption by Rubisco and assimilation in the Calvin cycle which regulates via a CO2‐demand? In favour of this assumption is the observation that malate consumption in Phase III is accelerated by increased light intensity enhancing photosynthesis (Kluge, 1968; Thomas et al., 1987). Against this is the sheer CO2‐concentration building up when it oversaturates C3‐photosynthesis (Rubisco and Calvin cycle). Substrate saturation of C3‐photosynthesis is between 0.1% and 0.4% (Berry and Downton, 1982). CO2‐assimilation during Phase III of CAM in Kalanchoë daigremontiana saturates at 0.2% (Maxwell et al., 1998).

Thus, according to the data of Table 1, there was strong substrate oversaturation of Rubisco in two Opuntia species, but not in a third species studied, and some oversaturation was observed in Agave desertii, Aloë vera, and perhaps Ananas comosus (although only in one of the two measurements) and K. daigremontiana. Such oversaturation would rule out a regulation of remobilization via the CO2 demand of Rubisco. On the other hand, this seems to be the exception rather than the rule. In the majority of cases in Table 1 Rubisco would operate at or below substrate saturation, and this allows the assumption that the regulation of remobilization is by Rubisco and that carbon dioxide is functioning as a controlling factor in decarboxy lation (Cockburn and Patel, 2002).

There appear to be no data on the critical level of piCO2 needed to close stomata in Phase III. The CO2‐response sensitivity of guard cells, in general, is very variable between species and also for a given species due to acclimation (Frechilla et al., 2002). Hence, it is hard to say if this effect is just saturated at the piCO2‐levels given (Table 1), and thus, a critical control point, or if it is oversaturated. The latter is assumed. Cockburn and Patel (2002) have performed a set of interesting experiments. When they exposed leaves of various CAM plants with the epidermis removed, leaf discs or leaf strips, with stomatal control eliminated, they found the cross‐over point where net CO2‐exchange became zero, i.e. uptake balanced release, at 0.06%. This is much lower than most piCO2 values in Table 1. In addition, patchiness of ϕPSII independent of stomatal reactions (Rascher, 2001) also suggests that CO2‐mediated regulation may be an internal effect related to Rubisco activity.

(ii) Consumption of cytoplasmic malate by decarboxy lation: Is the regulation of remobilization at the point of malate decarboxylation, where the demand for cytoplasmic malate would determine cytoplasmic malate supply from the vacuole? A mass action effect at this point is quite possible because all of the three malate decarboxylating enzymes occurring in different CAM plants, namely NAD‐ and NADP‐dependent malic enzymes and phosphoenolpyruvate‐carboxy‐kinase, catalyse readily reversible reactions. Driving the process towards decarboxylation requires reasonably high cytoplasmic malate levels. In fact, measurements of cytoplasmic pH during the diurnal CAM cycle in K. daigremontiana have shown a drop by 0.3 pH units at midday (Hafke et al., 2001). This may be due to both high levels of malic acid and CO2 in the cytoplasm (Fig. 1). With a buffer capacity of about 65 mM H+ per pH unit for a pHcyt of about 7.5, this corresponds to a cytoplasmic acid load equivalent to 10 mM malic acid.

(iii) Malic acid efflux from the vacuole: A third possibility is that it is malic acid efflux from the vacuole which determines the whole process. The mechanism by which this occurs is still unconfirmed, i.e. diffusion of malic acid (Lüttge and Smith, 1984) or a carrier‐ or channel‐mediated process (see Lüttge et al., 2000). What is known is that it is a passive process (Lüttge and Smith, 1984). It can be regulated, however, by changing tonoplast properties. During nocturnal malic acid accumulation, which has osmotic consequences, turgor pressure increases. This may have effects on the tonoplast which are important for the switching between nocturnal net malic acid accumulation and daytime net malic remobilization (Lüttge et al., 1975, 1977; Lüttge and Ball, 1977; Steudle et al., 1980; Smith et al., 1984). In fact, this is also part of a model suggesting the function of the tonoplast as the hysteresis switch in the oscillator of circadian rhythmicity of CAM (see below). As in option (ii), this would lead to rather high cytoplasmic malate concentrations in Phase III.

A cytoplasmic malate concentration of 10 mM corresponds to 10 mol m–3 CO2 if considered decarboxylated to pyruvate. A 2.5% CO2 partial pressure in the aerenchyma (Opuntia basilaris) would correspond to only about one‐tenth, i.e. 1.1 mol m–3 CO2. In leaves of K. daigremontiana the volumes of cytoplasm and air space are about 0.5–1% and about 3.6–8.8%, respectively (Rona et al., 1980; Smith and Heuer, 1981; and Maxwell et al., 1997; Rascher, 2001, respectively). Thus, the air‐space volume is about 10 times the cytoplasmic volume, and the above rough estimates would not reject the possibility that even the highest piCO2 measured (Table 1) is a consequence of equilibria established via strong vacuolar malic acid efflux and build‐up of cytoplasmic malate levels. However, it is still necessary to know much more about the rates of the enzyme reactions involved in establishing these equilibria.

A very important enzyme in these equilibria is carbonic anhydrase (CA). CO2 and HCO3 may diffuse in parallel in the cytosol and in the chloroplast stroma. In terrestrial C3 plants CA enhances the diffusion of Ci from the site at which atmospheric CO2 dissolves in cell wall or apoplast water to the site of CO2 consumption by Rubisco (Raven and Spicer, 1996). In the same way it could mediate CO2 release into the leaf air spaces of CAM plants in Phase III (Fig. 1). In C3‐plants CA is mainly expressed in the chloroplast stroma, although there is also activity in the cytosol (Reed, 1979). Tsuzuki et al. (1982) did not find cytosolic CA in some of the CAM plants they tested, while Holtum et al. (1984) found in all CAM plants tested that cytosolic CA activity exceeded the requirement of dark CO2‐fixation (Phase I), where HCO3 is the substrate of PEPC. By contrast, obviously nothing is known about the role of CA in Phase III (Raven and Spicer, 1996).

High piCO2 may be a downstream consequence of the operation of the regulation network (Fig. 1) including CA‐functions. piCO2‐levels saturating or oversaturating photosynthesis in Phase III do not appear to be an upstream requirement of CAM‐function inasmuch as often levels saturating Rubisco are not attained (see above and Table 1). In conclusion then, while the precise position and function of control points is not certain, clearly piCO2 may exert a signalling function between the rate of CO2‐consumption by Rubisco and malic acid remobilization from the vacuole.

Circadian rhythmicity of CAM a complication: oscillatory CO2‐concentrating?

The circadian rhythmicity of CAM is a complication. In the normal day/night cycle, switching between nocturnal net‐accumulation and daytime net‐remobilization and back again to net‐accumulation of malic acid can be readily explained by the changing external conditions shifting the equilibria. Equilibrium thermodynamics, therefore, provide sufficient explanation for the filling and emptying, respectively, of the vacuole. However, with the endogenous circadian rhythmicity under constant environmental conditions this does not apply.

The study of circadian rhythmicity needs continuous recording of long time‐series. Therefore, for technical reasons, it is mainly gas exchange, CO2 (JCO2) and water vapour (JH2O), that is monitored. Occasionally malate analyses are available and enzyme activities, PEPC and Rubisco were followed both directly by enzyme analyses and indirectly by the online collection of samples from the air flow through gas‐exchange systems to measure the different carbon‐isotope discrimination by the two enzymes (Grams et al., 1997). However, these techniques are tedious, destructive (malate analyses, enzyme extracts) or expensive (carbon‐isotope analyses) and long, densely sampled, time‐series are not available. Little is known about the internal partial pressures of CO2 and O2, piCO2 and piO2, during the circadian oscillations. From the point of view of this review, however, the question must be asked whether there is a circadian rhythmicity of CO2‐concentrating.

Bohn et al. (2001) have calculated the ratio of internal to external pCO2 (ci/ca) in Kalanchoë daigremontiana from measurements of JH2O and the derived leaf‐conductance for water vapour, gH2O. They have studied entrainment of the CAM‐cycle in K. daigremontiana under imposed external rhythms of light intensity of 140 µmol m–2 s–1 (lower) and 250 µmol m–2 s–1 (higher) with a period of 24 h and 16 h, respectively. They found oscillations of ci/ca between a minimum of 0.64 and a maximum of 1.03 at the low and high light intensities, respectively. ci/ca‐oscillations of similar magnitude are obtained during endogenous rhythmicity under constant external conditions (Tomasz Wyka, personal communication). Thus, there was some feeble CO2‐concentrating keeping ci/ca not too much below unity or even raising piCO2 slightly above the atmospheric partial pressure during the peaks of photosynthesis. This is an order of magnitude less than the CO2‐concentrating in Phase III of the normal CAM cycle, where ci/ca for K. daigremontiana was 12.5 (Table 1).

Does chlorophyll‐fluorescence imaging allow the black box to be denuded? In Rascher’s experiments (Rascher, 2001; Rascher et al., 2001) imaging only provided relative data on ϕPSII for comparisons within a given time series and no absolute measurements of ϕPSII which would allow the values obtained during circadian rhythmicity to be compared with those of the diurnal dark/light cycle. During the circadian time series ∫ϕPSII (integrated over the whole leaf) was low when JCO2 was zero and stomata were obviously closed. This is in contrast to Phase III of the normal day/night cycle of CAM when there is CO2‐concentrating and when ∫ϕPSII was high as JCO2 was zero. Does this mean that there is no CO2‐concentrating during the circadian rhythm? Or, perhaps, CO2‐concentrating may only be weak, as suggested by the experiments of Bohn et al. (2001; and Tomasz Wyka; see above), and could be overridden by the energy requirements of gluconeogenesis which tend to lower ϕPSII.

This then might support the idea that the strong CO2‐concentrating observed in Phase III of the normal day/night CAM cycle is a consequence of equilibria established as discussed above, but does not occur during the non‐linear dynamics of circadian oscillations. On the other hand, pronounced patchiness does occur during the circadian rhythm, and it also oscillates. Its level even increases considerably during the ongoing free running oscillations. Adjacent leaf patches of 5–10 mm diameter (approximately 30–60 cells) and 15–30 mm apart may get desynchronized, particularly during phases of ϕPSII decline. They synchronize again as ϕPSII increases. Most likely, as in the entrainment experiments of Bohn et al. (2001), the sequence of events is a reduction of net CO2‐uptake, JCO2, as piCO2 increases, followed by a decrease of gH2O.

Thus, there must be at least some internal CO2‐concentrating. Internal CO2 is thought to be the synchronizing signal. However, as seen above, due to the diffusion restriction inside the leaf of K. daigremontiana this requires high internal CO2‐concentrations. They are high enough in Phase III of normal CAM where patchiness is low and much lower, even at their highest levels, in circadian oscillations where patchiness remains high and synchronization/desynchronization shows strong dynamics.

What does this say about CAM and its Phase III CO2‐concentrating phenomenon? Although it was found that the gene for PEPC‐kinase, the enzyme which regulates PEPC‐activity, is under circadian control (Nimmo et al., 1987, 2001; Carter et al., 1991, 1996; Kusumi et al., 1994; Hartwell et al., 1996, 1999), and in fact many other genes, especially of enzymes involved in CAM are clock‐controlled genes (Boxall et al., 2001), it appears that circadian CAM performance is under the control of a post‐translational machinery or oscillator. The PEPC‐kinase gene is under the control of metabolism and cytoplasmic/vacuolar malate compartmentation (Borland et al., 1999; Nimmo, 2000). The tonoplast was shown to have oscillator function. piCO2 being directly related to malate compartmentation and metabolism, namely, malate decarboxylation, may indeed be an important internal signal. It may act directly or in a signal transduction chain, where pH effects and membrane potential transients may also be involved (Stahlberg et al., 2001). Whether this signalling function of piCO2 is necessary for the normal day/night cycle of CAM is not known. Perhaps not in Phase III, when piCO2 is homogenously high. However, piCO2‐signalling may be important in the regulation of changes between Phases II and III and III and IV, respectively, as alluded to by the transients of inhomogeneity at these times as discussed above.

Established beneficial ecophysiological consequences: but are there not open questions?

There are four ecophysiological consequences of CO2‐concentrating in CAM, which are well established in the literature and widely accepted.

CO2‐acquisition

Nocturnal CO2‐fixation and daytime interior CO2‐concentrating by submerged plants in fresh water is a mechanism of CO2‐acquisition avoiding daytime competition with non‐CAM photosynthesizers (Griffiths, 1989; Keeley, 1996). CO2‐concentrating itself is the ecophysiological benefit here. This may have been the original driving force of CAM‐evolution as it is expressed in plants as phylogenetically early as the lycopodiopsid Isoëtes (Griffiths, 1989; Keeley, 1996). However, is this beneficial consequence not restricted to fresh‐water plants?

Water‐use‐efficiency (WUE)

In terrestrial CAM plants Phase III CO2‐concentrating allows CO2‐assimilation behind closed stomata. This minimizes transpirational water loss during the hottest part of the day with the largest atmospheric water vapour pressure deficit, while stomata are open during the night for CO2 acquisition. CAM plants have the highest water‐use‐efficiency (WUE).

This has led to a widely held view that CAM plants typically are inhabitants of the driest arid sites. True, there c. 1800 species of stem succulent cacti and leaf succulent agaves, the CAM plants of deserts. However, among the orchids and bromeliads there is an estimated 10 000 CAM species which are typical elements of the flora of the moister shaded, semi‐shaded and semi‐exposed rainforests (Lüttge, 2000). Does this mean that WUE needs to be challenged as the major ecophysiological trait of CAM and foremost driving force of CAM‐evolution? Or is this explained by the fact that most of the CAM‐species of tropical forests are epiphytes with their own particular problems of water supply as they do not root in a soil substrate (Zotz and Hietz, 2001)?

Suppressed photorespiration

Rubisco has two competing substrates, CO2 and O2. Phase III CO2‐concentrating elevates substrate levels for the carboxylating activity of Rubisco so that it can operate at or close to substrate saturation beginning above 0.1% CO2 (Berry and Downton, 1982; Table 1). This suppresses the oxygenase activity of Rubisco, i.e. photorespiration (Osmond et al., 1982; Cushman and Bohnert, 1997), and hence, the loss of CO2 related to photorespiration. However, CO2‐concentrating itself may lead to some loss of CO2, namely by diffusion out of the leaf at the very high CO2 gradient between the leaf air spaces and the atmosphere (Friemert et al., 1986).

The photorespiratory machinery remains active in CAM plants (Osmond et al., 1982; Whitehouse et al., 1991; Edwards et al., 1996; Heber et al., 1996). It is needed in Phase IV when plants perform C3‐photosynthesis with open stomata and atmospheric CO2. But what about Phase III? To what extent is O2‐concentrating as a consequence of CO2‐concentrating counteracting the effect of the latter? In most cases O2/CO2 ratios in the leaves in Phase III remain well below those in the ambient atmosphere by a factor of 0.1–0.5 (Spalding et al., 1979; Table 2). Nonetheless, photorespiration may make important contributions to oxygen metabolism in Phase III of CAM (Thomas et al., 1987). Catalase (CAT), the H2O2 scavenging enzyme in photorespiration, remained unchanged during C3 to CAM shifts in the C3/CAM intermediate species Sedum album under mild drought stress (Castillo, 1996). In the C3/CAM intermediate species Mesembryanthemum crystallinum CAT activity was significantly reduced in the CAM‐state compared to the C3‐state. However, during the diurnal cycle it increased in the light period and showed a peak in the late afternoon (Niewiadomska et al., 1999). Nothing is known about the dynamics of piO2 in the light period and to what extent piO2 remains high at the end of Phase III when the malic acid store is exhausted, piCO2 declined and Phase IV stomatal opening has not (yet) occurred. Under particular water stress Phase IV stomatal opening may not be expressed at all (Smith and Lüttge, 1985). Photorespiration may then become very important. Miszalski et al. (2001) used the mitochondrial enzyme fumarase as an indicator of the function of the tricarboxylic acid cycle. Its activity was much reduced in Mesembryanthemum crystallinum in the CAM‐state as compared to the C3‐state. High mitochondrial activity during the light period decreases photorespiration because it consumes O2 and generates CO2 and hence tends to increase the internal CO2:O2 ratio (Krömer, 1995). Thus, low mitochrondrial activity in the CAM‐state may indicate increased photorespiration as a protection against stress by high O2‐levels (Miszalski et al., 2001). Therefore, can suppressed photorespiration really be considered as a beneficial consequence of CO2‐concentrating?

Prevented overenergization of the photosynthetic energy transduction pathway

Another long‐accepted beneficial consequence of Phase III CO2‐concentrating and operation of Rubisco near substrate saturation was the reduction or even prevention of stress due to over‐energization of the photosynthetic apparatus and production of reactive oxygen species (ROS). This has even been considered to have been a major driving force for the evolution of CAM (Gil, 1986). Conversely, it is now known that CAM plants are subject to photoinhibition, not only in Phase IV when stomata are open and CO2 is taken up from the atmosphere, but also in Phase III with the very high piCO2 (see Lüttge, 2000, for a review).

This is not the occasion to get involved in the increasingly sophisticated discussion of protective acute photoinhibition or scaling down of photosystem II efficiency, with electrical or zeaxanthin cycle quenching of potential quantum yield of PSII, destructive but protective D1‐protein turnover and chronic destructive photoinhibition. It is just important to note that in the CO2‐concentrating Phase III, photoinhibition does occur; the xanthophyll cycle is involved (Adams et al., 1987; Adams, 1988; Keiller et al., 1994; Herzog et al., 1999; see Lüttge, 2000, for a review) and acclimation to high light stress may occur in CAM plants (Adams et al., 1987; Winter and Awender, 1989; Fetene et al., 1990; Lüttge et al., 1991; Robinson and Osmond, 1994; Adams and Demmig‐Adams, 1996).

Therefore, is reduced oxidative stress really to be considered as a beneficial consequence of CO2‐concentrating in Phase III of CAM? What are the benefits of C3‐photosynthesis contrasted with CAM? In the studies of C3/CAM‐intermediate plants the question mostly asked is what drives the shift from C3‐photosynthesis to CAM? When reversible shifts occur in both directions, which is the case in the perennial C3/CAM‐intermediate Clusia minor, what drives a shift from CAM to C3‐photosynthesis (de Mattos et al., 2001)? In fact, for C. minor it was shown that it was not so much CAM per se which offered ecological advantages but the flexibility given by CAM and C3 options (Herzog et al., 1999). Moreover, in growth chamber experiments when C. minor switched from CAM to C3‐photosynthesis, CO2‐uptake over 24 h (dark fixation plus light fixation of external CO2) more than doubled. This implies that the integrated daily photon utilization for the photochemical work of CO2‐assimilation increased. The cost paid by the plant is water because, concomitantly, WUE was reduced to a third (de Mattos et al., 2001). However, this observation supports the idea that C3‐photosynthesis may provide superior protection from irradiance stress compared to CAM. This was already suggested some time ago by the finding that an increase of photosynthetically active radiation from 360 to 1200 µmol m–2 s–1 led to a suppression of night‐time CO2‐fixation and much increased daytime CO2‐fixation in Clusia minor, but only when plants were well watered (Schmitt et al., 1988; Lüttge, 1996).

Adverse ecophysiological consequences

If some of the established beneficial consequences of CO2‐concentrating are being questioned, is it also necessary to even pinpoint some adverse consequences? There are at least three, of which the third is a major one:

Energetics

CAM is more costly energetically than C3‐photosynthesis. The stoichiometrics of paper‐biochemistry have been worked out in every detail (Winter and Smith, 1996b). Energy demand was also measured under laboratory conditions (Maxwell et al., 1998). Whether this is a limiting factor under actual environmental conditions with adverse ecophysiological consequences is much less clear. It might be in shaded habitats of CAM plants in tropical forests. However, as an example, among shaded, semi‐shaded and exposed plants of the CAM‐bromeliad Bromelia humilis it was the shaded individuals, which showed the lushest growth (Lee et al., 1989).

pH

Phase III CO2‐concentrating has pH‐effects. The acidification of the cytosol by 0.3 pH units (Hafke et al., 2001; see above) may have useful consequences in cellular regulation in that it inhibits PEPC and the vacuolar H+‐transporting V‐ATPase, thus avoiding futile recycling of CO2 into malate and vacuolar malic acid during Phase III.

However, the high apoplastic and leaf air‐space CO2‐concentrations must have effects on cell wall pH (Fig. 1). These effects may be quite important when piCO2 attains several per cent, although these cases may be rare (Table 1). There may also be other consequences, for example, on plasma membrane electrical potentials (Hedrich et al., 2001). This has been completely overlooked so far. There is no work on apoplastic pH in CAM cells in Phase III nor any discussion of it in the CAM literature.

Phase III oxidative stress

The O2‐concentrating consequence of CO2‐concentrating in planta has been demonstrated above. Studies of O2‐exchange by mass spectroscopy of stable oxygen isotopes (16O, 18O) and the main electron flow of chloroplasts have revealed the strong O2 production in CAM plants in the light (Thomas et al., 1987). Experiments measuring O2 evolution in the leaf disc O2‐electrode, where leaf discs of CAM plants have been artificially exposed to 5% ambient CO2, were performed (Adams et al., 1987; Adams and Osmond, 1988; Borland and Griffiths, 1989; Maxwell et al., 1998), and Osmond et al. (1996) note the action of the ‘O2‐pump’ leading ‘to ever‐increasing O2‐concentration in the experimental chamber as malate decarboxylation proceeds’. Thus Phase III CO2‐concentrating not only stays short of preventing photorespiration and photoinhibition (see above), but the corresponding O2‐concentrating even may be one of the major adverse consequences.

High piO2 supports formation of aggressive reactive oxygen species (ROS) such as hydrogen peroxide (H2O2), superoxide (O2•–) and the hydroxyl radical (OH). CAM plants are especially equipped to deal with this oxidative stress by the increased expression of an antioxidative response system (ARS). In the C3/CAM‐intermediate Sedum album this appeared to be especially important during the C3 to CAM transition when the activities of ascorbate peroxidase, superoxide dismutase (SOD), gluthatione reductase, and monodehydroascorbate reductase as ROS scavenging enzymes were increased (Castillo, 1996). In the C3/CAM‐intermediate M. crystallinum, where CAM‐induction is elicited by salt stress, it is often difficult to distinguish ARS expression in response to salinity from requirements of CAM, for example, the up‐regulation of cytosolic CuZn‐dependent SOD (Hurst and Ratajczak, 2002). It appears, however, that up‐regulation of mitochrondrial Mn‐dependent SOD is a typical reaction to the induction of CAM (Miszalski et al., 1998; Broetto et al., 2002).

By cross‐tolerance this may also explain the observation that CAM plants are less sensitive to the oxidative stress given by SO2 and O3 than C3 plants (Olszyk et al., 1987; Miszalski et al., 1997). This is not an effect of day‐time stomatal closure as shown by using protoplasts or leaf discs of Kalanchoë daigremontiana with the epidermis stripped off. The larger SO2‐tolerance is clearly a CAM‐inherent phenomenon related to the high activity of photosynthetic electron transport, and the O2‐evolution enhancing oxidation of SO2. Miszalski et al. (1997) compared ‘acidified’ protoplasts loaded with malic acid and ‘deacidified’ protoplasts, where malic acid was broken down. Bicarbonate in the medium strongly enhanced O2 evolution by the acidified protoplasts and SO2 reduction was much higher than in the deacidified protoplasts (Table 3).

These particular protective measures must be reflections of the adverse consequences to avoid damage.

Questions II

While the mechanistic consequences of CO2‐concentrating in CAM are fascinating, there is scepticism about some of the long‐accepted beneficial ecophysiological consequences and even some adverse ecophysiological consequences are being recognized. Is this the answer to the question: ‘Why aren’t more plants committed to CAM?’ (Griffiths, 2001). ‘Why do so few plants bother to induce CAM and when they do, why is it then merely a maintenance mechanism?’ (Griffiths, 2001). Is it then in fact only ‘the fringe benefits of CAM which makes it pervasive in many families’? (Griffiths, 2001). What are these fringe benefits in terms of ecophysiological competitiveness and ecological niche occupation? When the fringe benefits are essential in such respects, are the adverse ecophysiological consequences only unavoidable side‐effects of the fascinating mechanistic consequences of CO2‐concentrating that are to be accepted and tolerated? Alas! Is CAM really ‘not so much a curiosity’ (Griffiths et al., 2001)? Doesn’t it remain a ‘curiosity’ (Osmond, 1978)?

Acknowledgements

I thank Richard Leegood and Uwe Rascher for reading this review and for valuable suggestions prior to publication.

Fig. 1. Malic acid/malate remobilization and decarboxylation and build‐up of high internal CO2‐concentrations in Phase III of CAM and control points discussed in the text, i.e. (i) CO2‐consumption by Rubisco, (ii) consumption of cytoplasmic malate by decarboxylation, and (iii) malic acid efflux from the vacuole. CA=carbonic anhydrase.

Fig. 1. Malic acid/malate remobilization and decarboxylation and build‐up of high internal CO2‐concentrations in Phase III of CAM and control points discussed in the text, i.e. (i) CO2‐consumption by Rubisco, (ii) consumption of cytoplasmic malate by decarboxylation, and (iii) malic acid efflux from the vacuole. CA=carbonic anhydrase.

Table 1.

Maximum internal CO2‐concentrations and ci/ca‐ratios (where cawas taken as 0.04%) measured in various CAM‐plants in the light period, at midday, in Phase III of CAM by various methods Note that the determination of piCO2 from measurements of gas exchange, i.e. leaf‐conductance for water vapour, gH2O, is only possible when stomata are open, at least to some extent, and that errors become large as stomata close in Phase III. Thus the values reported by Friemert et al. (1986) are approximations, showing the order of magnitude though.

Species  Family piCO2 (%) ci/ca Method  Reference 
Opuntia ficus‐ indica Cactaceae 1.30 32.5 Thermal conductivity Cockburn et al., 1979 
Opuntia basilaris Cactaceae 2.50 62.5 Thermal conductivity Cockburn et al., 1979 
Opuntia monacantha  Cactaceae 0.12 3.0 Gas chromatography Spalding et al., 1979 
Agave desertii Agavaceae 0.80 20.0 Thermal conductivity Cockburn et al., 1979 
Yucca schidigera  Agavaceae 0.40 10.0 Thermal conductivity Cockburn et al., 1979 
Aloë vera  Liliaceae 0.60 15.0 Thermal conductivity Cockburn et al., 1979 
Ananas comosus  Bromeliaceae 0.50 12.5 Thermal conductivity Cockburn et al., 1979 
Ananas comosus  Bromeliaceae 0.13 3.3 Gas chromatography Spalding et al., 1979 
Cattleya sp.  Orchidaceae 0.15 3.8 Thermal conductivity Cockburn et al., 1979 
Phalaenopsis sp.  Orchidaceae 0.23 5.8 Thermal conductivity Cockburn et al., 1979 
Hoya carnosa  Asclepiadaceae 0.08 2.0 Gas chromatography Spalding et al., 1979 
Huernia sp.  Asclepiadaceae 0.14 3.5 Gas chromatography Spalding et al., 1979 
Kalanchoë gastonis‐bonnieri Crassulaceae 0.27 6.8 Gas chromatography Spalding et al., 1979 
Kalanchoë tomentosa  Crassulaceae 0.35 8.8 Gas chromatography Spalding et al., 1979 
Kalanchoë daigremontiana Crassulaceae 0.50 12.5 Gas chromatography Kluge et al., 1981 
Kalanchoë tubiflora  Crassulaceae 0.09 2.3 Gas exchange Friemert et al., 1986 
Sedum praealtum  Crassulaceae 0.29 7.3 Gas chromatography Spalding et al., 1979 
Sedum morganianum  Crassulaceae 0.26 6.5 Gas exchange Friemert et al., 1986 
Sempervivum tectorum  Crassulaceae 0.12 3.0 Gas exchange Friemert et al., 1986 
Species  Family piCO2 (%) ci/ca Method  Reference 
Opuntia ficus‐ indica Cactaceae 1.30 32.5 Thermal conductivity Cockburn et al., 1979 
Opuntia basilaris Cactaceae 2.50 62.5 Thermal conductivity Cockburn et al., 1979 
Opuntia monacantha  Cactaceae 0.12 3.0 Gas chromatography Spalding et al., 1979 
Agave desertii Agavaceae 0.80 20.0 Thermal conductivity Cockburn et al., 1979 
Yucca schidigera  Agavaceae 0.40 10.0 Thermal conductivity Cockburn et al., 1979 
Aloë vera  Liliaceae 0.60 15.0 Thermal conductivity Cockburn et al., 1979 
Ananas comosus  Bromeliaceae 0.50 12.5 Thermal conductivity Cockburn et al., 1979 
Ananas comosus  Bromeliaceae 0.13 3.3 Gas chromatography Spalding et al., 1979 
Cattleya sp.  Orchidaceae 0.15 3.8 Thermal conductivity Cockburn et al., 1979 
Phalaenopsis sp.  Orchidaceae 0.23 5.8 Thermal conductivity Cockburn et al., 1979 
Hoya carnosa  Asclepiadaceae 0.08 2.0 Gas chromatography Spalding et al., 1979 
Huernia sp.  Asclepiadaceae 0.14 3.5 Gas chromatography Spalding et al., 1979 
Kalanchoë gastonis‐bonnieri Crassulaceae 0.27 6.8 Gas chromatography Spalding et al., 1979 
Kalanchoë tomentosa  Crassulaceae 0.35 8.8 Gas chromatography Spalding et al., 1979 
Kalanchoë daigremontiana Crassulaceae 0.50 12.5 Gas chromatography Kluge et al., 1981 
Kalanchoë tubiflora  Crassulaceae 0.09 2.3 Gas exchange Friemert et al., 1986 
Sedum praealtum  Crassulaceae 0.29 7.3 Gas chromatography Spalding et al., 1979 
Sedum morganianum  Crassulaceae 0.26 6.5 Gas exchange Friemert et al., 1986 
Sempervivum tectorum  Crassulaceae 0.12 3.0 Gas exchange Friemert et al., 1986 
Table 2.

Ratios of internal concentrations of O2:CO2at midday in various CAM plants (Spalding et al., 1979)

 O2/CO2 
Ambient air 633 
Opuntia monacantha 205 
Ananas comosus 161 
Hoya carnosa 285 
Huernia sp 174 
Kalanchoë gastonis‐bonnieri 155 
Kalanchoë tomentosa 88 
Sedum praealtum 81 
 O2/CO2 
Ambient air 633 
Opuntia monacantha 205 
Ananas comosus 161 
Hoya carnosa 285 
Huernia sp 174 
Kalanchoë gastonis‐bonnieri 155 
Kalanchoë tomentosa 88 
Sedum praealtum 81 
Table 3.

O2‐evolution and SO2‐oxidation by acidified (loaded with malic acid) and deacidified (malic acid broken down) protoplasts of Kalanchoë daigremontiana (Miszalski et al., 1997) In O2‐evolution measurements either no bicarbonate or 10 mmol l–1 bicarbonate was added to the medium. This is not specified by the authors for the SO2‐oxidation measurements. For direct comparison with rates of O2‐evolution, rates of SO2‐oxidation h–1 were calculated taking four times the SO2 oxidized in the first 15 min of the experiment shown.

Rate (µmol mg–1 chlorophyll h–1Deacidified Acidified 
O2‐evolution without HCO3 8.6 15.7 
O2‐evolution with 10 mM HCO3 13.2 32.3 
SO2‐oxidation 9.2 22.0 
Rate (µmol mg–1 chlorophyll h–1Deacidified Acidified 
O2‐evolution without HCO3 8.6 15.7 
O2‐evolution with 10 mM HCO3 13.2 32.3 
SO2‐oxidation 9.2 22.0 

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