Chloroplast thylakoid structure in evergreen leaves employing strong thermal energy dissipation.

Journal of Photochemistry and Photobiology B: Biology xxx (2015) xxx–xxx

Contents lists available at ScienceDirect

Journal of Photochemistry and Photobiology B: Biology journal homepage: www.elsevier.com/locate/jphotobiol

Chloroplast thylakoid structure in evergreen leaves employing strong thermal energy dissipation Barbara Demmig-Adams ⇑, Onno Muller 1, Jared J. Stewart, Christopher M. Cohu 2, William W. Adams III Department of Ecology and Evolutionary Biology, University of Colorado, Boulder, CO 80309-0334, USA

a r t i c l e

i n f o

Article history: Received 21 January 2015 Received in revised form 16 March 2015 Accepted 18 March 2015 Available online xxxx

a b s t r a c t In nature, photosynthetic organisms cope with highly variable light environments – intensities varying over orders of magnitudes as well as rapid fluctuations over seconds-to-minutes – by alternating between (a) highly effective absorption and photochemical conversion of light levels limiting to photosynthesis and (b) powerful photoprotective thermal dissipation of potentially damaging light levels exceeding those that can be utilized in photosynthesis. Adjustments of the photosynthetic apparatus to changes in light environment involve biophysical, biochemical, and structural adjustments. We used electron micrographs to assess overall thylakoid grana structure in evergreen species that exhibit much stronger maximal levels of thermal energy dissipation than the more commonly studied annual species. Our findings indicate an association between partial or complete unstacking of thylakoid grana structure and strong reversible thermal energy dissipation that, in contrast to what has been reported for annual species with much lower maximal levels of energy dissipation, is similar to what is seen under photoinhibitory conditions. For a tropical evergreen with tall grana stacks, a loosening, or vertical unstacking, of grana was seen in sun-grown plants exhibiting pronounced pH-dependent, rapidly reversible thermal energy dissipation as well as for sudden low-to-high-light transfer of shade-grown plants that responded with photoinhibition, characterized by strong dark-sustained, pH-independent thermal energy dissipation and photosystem II (PSII) inactivation. On the other hand, full-sun exposed subalpine confers with rather short grana stacks transitioned from autumn to winter via conversion of most thylakoids from granal to stromal lamellae concomitant with photoinhibitory photosynthetic inactivation and sustained thermal energy dissipation. We propose that these two types of changes (partial or complete unstacking of grana) in thylakoid arrangement are both associated with the strong non-photochemical quenching (NPQ) of chlorophyll fluorescence (a measure of photoprotective thermal energy dissipation) unique to evergreen species rather than with PSII inactivation per se. Ó 2015 Published by Elsevier B.V.

1. Introduction Photosynthesis deals with a wide range of light intensities covering several orders of magnitude and, in most natural habitats, also responds to rapid pronounced fluctuations in light level over the course of a single day. Photosynthetic organisms can thus alternate between (a) highly effective absorption and photochemical conversion of light levels limiting to photosynthesis and (b) powerful dissipation of that fraction of light absorbed in high light environments that cannot be utilized in photosynthesis and could thus be potentially damaging. Highly variable light intensities are encountered on second-to-minute scales in natural locations with, ⇑ Corresponding author. E-mail address: [email protected] (B. Demmig-Adams). Present address: Institute of Bio- and Geosciences, IBG-2: Plant Sciences, Forschungszentrum Jülich GmbH, 52425 Jülich, Germany. 2 Present address: Dow AgroSciences, Portland, OR, USA. 1

e.g., overhead forest canopy [1–3] or variable cloud cover. On clear days in open locations, photosynthesis in exposed leaves must contend with light intensities that vary over two orders of magnitude from limiting light in early morning and late afternoon versus full sunlight during midday [3,4–6]. Adjustments of the photosynthetic apparatus to changes in light environment occur on many scales and involve biophysical, biochemical, and structural adjustments. One aspect of the structure–function relationship that serves to adjust the photosynthetic apparatus to growth light environment involves the photosynthetic (thylakoid) membrane. Thylakoids are organized into granal regions, where thylakoid membranes are stacked, versus stromal lamellae with individual unstacked thylakoids. The two photosystems exhibit an uneven distribution among stromal lamellae and grana; while the stacked granal regions house photosystem II (PSII) core complexes along with their associated light-harvesting complexes (LHCII), photosystem I (PSI) complexes and their

http://dx.doi.org/10.1016/j.jphotobiol.2015.03.014 1011-1344/Ó 2015 Published by Elsevier B.V.

Please cite this article in press as: B. Demmig-Adams et al., Chloroplast thylakoid structure in evergreen leaves employing strong thermal energy dissipation, J. Photochem. Photobiol. B: Biol. (2015), http://dx.doi.org/10.1016/j.jphotobiol.2015.03.014

2

B. Demmig-Adams et al. / Journal of Photochemistry and Photobiology B: Biology xxx (2015) xxx–xxx

light-harvesting complex (LHCI) reside preferentially in the stromal lamellae [7,8; for a recent review, see 9]. Sun-grown leaves typically have grana with relatively few stacked thylakoids, whereas shade-grown leaves have grana with many more thylakoids [8], and the number of thylakoid membranes per granum also differs among species with different shade tolerance. The highly shade-tolerant tropical evergreen Alocasia macrorrhiza was reported to feature as many as a hundred stacked thylakoid membranes in its grana when grown in deep shade [10], while annual herbaceous species do not feature this many membranes in their grana even when grown in the shade [8]. Superimposed upon these adjustments in response to growth light environment that occur over days and weeks, faster dynamic adjustments occur in response to sudden changes in light environment (see below; see also [9]). Various types of dynamic rearrangements of pigment-protein complexes in thylakoid organization have been recognized, including structural changes in the service of (i) optimizing utilization of limiting light levels for photosynthesis via balancing the delivery of light absorbed by LHCII to either PSII or PSI [11–13], (ii) facilitating protein removal and replacement in the rapid turn-over cycle of the proteins constituting the core of photosystem II [14], and (iii) photoprotective thermal dissipation of excess absorbed light – quantified from nonphotochemical quenching (NPQ) of chlorophyll fluorescence (see, e.g., [3,4]). Horton [15] stated that, ‘‘dynamic changes in the organization of the grana membrane’’ are ‘‘at the heart of NPQ.’’ Furthermore, Ruban and Mullineaux [16] maintain that, ‘‘protein mobility is significantly modulated by physiological adaptation’’ and may differ in ‘‘different plant species’’. Changes in thylakoid organization in the context of NPQ have thus far been studied predominantly in annual species with mesophytic leaves like spinach or Arabidopsis that exhibit much higher rates of electron transport and much lower maximal NPQ capacities than evergreens. Groundbreaking early work on thylakoid membrane structure by Murakami and Packer [17] was conducted using the annual species spinach. Plant lifespan is inversely associated with apparent growth rate and maximal photosynthesis rate. Rapidly growing annuals or short-lived (ephemeral) weedy (herbaceous) species with soft-tissue (mesophytic) leaves grow rapidly, have high maximal rates of photosynthesis, use a large fraction of the light they absorb in photosynthesis, and have relatively low maximal NPQ capacities; long-lived evergreen species with tough (sclerophytic) leaves grow more slowly, spend stressful seasons with arrested growth, and have low maximal photosynthesis rates and exceptionally high NPQ capacities [3,6,18–20], here referred to as strong NPQ. In the present study, we characterized strong thermal energy dissipation in species with sclerophytic leaves, including a tropical evergreen tolerant of deep shade and evergreen conifers that experience cold winters at high altitude in the Rocky Mountains, in order to test a series of hypotheses: In the tropical evergreen, growth under high versus low light will result not only in higher electron transport rates, a greater maximal capacity of rapidly reversible, pH-dependent NPQ [20], and grana stacks with fewer membranes, but also in a more fluid grana structure. Sudden transfer of deep-shade-grown leaves of the tropical evergreen to high growth light will induce – over the course of days – a decrease of grana stacks along with a more fluid appearance of the thylakoid membranes in the remaining grana, rendering thylakoid structure similar to that of sun-grown leaves. Temperate evergreens overwintering in sun-exposed locations, exhibiting photosynthetic down-regulation along with strong, continuously maintained, dark-sustained (locked-in) thermal

energy dissipation, will also exhibit predominantly stromal thylakoids with no to very few grana during periods of growth arrest in the winter, but will exhibit more frequent grana during the growing period. 2. Methods 2.1. Plants and growth conditions Single-stem Monstera deliciosa Liebm. plants were grown in Canadian Growing Mix 2 (Conrad Fafard Inc., Agawam, MA, USA) in large 7.6 l pots in a sun-lit glasshouse (maximal photon flux density [PFD] = 1500 lmol m2 s1) or in 3.8 l pots under low, nonfluctuating light (PFD = 10 lmol m2 s1), respectively, receiving nutrients and water regularly. The third youngest leaf on each plant that was fully expanded and dark green was used for characterization. Two subalpine conifers, Picea engelmannii Parry ex Engelm. (Engelmann spruce) and Abies lasiocarpa (Hook.) Nutt. (subalpine fir), growing at approximately 2800 m in the Roosevelt National Forest (40°010 N, 105°310 W), were examined in early autumn (28 September 2011) and in the middle of winter (2 February 2012). Only needles that had emerged in 2011 and developed in full sunlight on south-facing branches were characterized. 2.2. Conifer needle collection and preparation Within a few hours following sunrise, fully sun-exposed (southfacing) branches of 4 trees were selected and one current year shoot from each branch was clipped off and placed between wet paper towels in a dark container. Branches were kept in the dark container for at least 1 h at room temperature. The needles from the sun-exposed side of the branch were clipped off and about 10 needles were aligned on a non-fluorescent, porous adhesive tape for fluorescence and photosynthetic analysis. 2.3. Light- and CO2-saturated photosynthetic capacity of oxygen evolution The capacity for photosynthetic oxygen evolution was measured using a leaf-disk oxygen electrode system (Model LD-2, equipped with an LS-2 halogen light source; Hansatech, King’s Lynn, Norfolk, UK) at 25 °C in 5% CO2, 21% O2, balance N2 [22,23]. For the conifers, the projected sample area was determined from a scanned image, taken after measurements, using ImageJ (Rasband W.S., ImageJ, U.S. National Institute of Health, Bethesda, MD, USA, http://imagej.nih.gov/ij/, 1997–2012). For Monstera, a 10-cm2 leaf disk was used. 2.4. Chlorophyll fluorescence Chlorophyll-fluorescence parameters of PSII efficiency (intrinsic efficiency in the dark, Fv/Fm, or effective efficiency under actinic light, ½F 0m F=F 0m ) and NPQ (F m =F 0m 1 from illuminated samples or Fm unquenched/Fm quenched1 from samples at the end of the nocturnal dark period) were measured as described in [6]. Unquenched Fm was determined at the end of the dark period prior to experimental high-light treatments. Maximal NPQ capacities from illuminated samples were ascertained from leaf disks maintained at 25 °C and exposed to 2000 lmol photons m2 s1 for 20 min under conditions (2% O2, balance N2) preventing high rates of linear electron transport but allowing build-up of a trans-thylakoid DpH. Darksustained NPQ was computed (as Fm unquenched/Fm quenched1) from Fm ascertained at the end of the nocturnal dark period both before and after experimental high-light exposure.



2.5. Nigericin treatment of Monstera leaves Treatments with the uncoupler nigericin were conducted as described in [21], using rapid vacuum-infiltration of small leaf disks (with buffer containing 50 mM Hepes [pH 7.6], 0.01% Tween, and 2 mM nigericin) via 3 consecutive vacuum treatments for 5 s each.

3

2.9.2. Subalpine conifers For dark-adapted needles, the same procedure was followed as for Monstera, except that needles were cut into 2-mm segments and, because the stain appeared to penetrate to a depth of less than one mm from the cut end of the needle, only the stained portion was sectioned and used for analysis. 2.10. Statistical analyses

2.6. Pigment extraction and quantification Pigments were extracted and quantified as described in [24].

2.7. Protein level quantification Proteins were extracted and quantified as described in [21,25]. Rabbit anti-PsbS antibody raised against the peptide sequence CGDRGKFVDDPPTG of Arabidopsis PsbS [26] was used for PsbS detection.

2.8. Monstera photoinhibition Whole plants grown either under low, non-fluctuating light indoors (10 lmol photons m2 s1) or in a sun-lit greenhouse (with a peak PFD of 1500 lmol m2 s1) were transferred to a temperature-controlled growth chamber (10-h photoperiod of 700 lmol photons m2 s1/14 h dark). A large water filter (between light source and plant) was used to maintain leaf temperature at 25 °C during exposure to 700 lmol photons m2 s1, while leaf temperature was kept at 22 °C during the dark period. Only fully expanded, green leaves were characterized. PSII efficiency was assessed near the end of the dark period (as intrinsic PSII efficiency Fv/Fm after 13 h in the dark) or before the end of the photoperiod (as effective PSII efficiency ½F 0m F=F 0m after 7 h in the light) and at the same time points over five days of subsequent recovery under constant exposure to 10 lmol photons m2 s1. NPQ at the end of the dark period (dark-sustained NPQ) was calculated as the ratio of dark Fm unquenched/dark Fm quenched1, where dark Fm unquenched was determined before the transfer to 700 lmol photons m2 s1 and dark Fm quenched after this transfer. During the subsequent recovery period at constant low light, dark Fm was assessed at the same time points after 5 min of darkening.

For comparisons between two means, a Student’s t-test was applied, whereas an ANOVA coupled with a Tukey–Kramer test for honestly significant differences was used for comparison of multiple means (JMP software, Pro 11.0.1, SAS Institute Inc., Cary, NC, USA). 3. Results 3.1. Thermal energy dissipation and thylakoid structure in sun-grown plants of the tropical evergreen Monstera exhibiting rapidly reversible strong NPQ Monstera plants grown in a sun-lit glasshouse exhibited severalfold higher levels of light- and CO2-saturated photosynthetic capacity than plants grown under a non-fluctuating low PFD of 10 lmol m2 s1 (Fig. 1A). At the end of each night, leaves of both sets of plants had a maximally high PSII efficiency (Fig. 1B). Sun-grown plants also exhibited much greater maximal levels of reversible NPQ, as a measure of photoprotective thermal energy dissipation, than low-light-grown plants (Fig. 1C; for time courses of chlorophyll fluorescence quenching upon exposure to high light, see Fig. 2). In response to a brief (20 min) high light exposure,

2.9. Chloroplast ultrastructure characterization 2.9.1. Monstera Leaf disks of 3 cm2 were obtained predawn or placed under high light for 20 min (see fluorescence analysis for details) and then, while continuing to be exposed to high light, rapidly (within one minute) cut up into small leaf-tissue cuttings of 2 mm 2 mm at room temperature into fixate of 10 ml of 10% (w/v) glutaraldehyde and 6.2 ml of 16% (w/v) paraformaldehyde in 25 ml of 70-mM sodium cacodylate buffer (pH 6.9) with 8.8 ml water, rinsed thrice in 70-mM sodium cacodylate buffer, and post-fixed for 1 h at room temperature in 4% osmium tetroxide in the same buffer. After rinsing in 70-mM sodium cacodylate buffer, samples were stained in 2% (w/v) uranyl acetate for 1 h, rinsed thoroughly in de-ionized water, dehydrated in acetone series, and embedded in Spurr resin (Electron Microscopy Sciences, Hatfield, Pennsylvania, USA). Ultrathin sections (70–80 nm) were cut with a diamond knife and stained with 2% (w/v) uranyl acetate and 0.2% (w/v) lead citrate. Digital pictures captured from a CM10 transmission electron microscope (Philips Electronic Instruments, Mahwah, NJ) were used for analysis.

Fig. 1. (A) Maximal photosynthetic capacity (light- and CO2-saturated rate of oxygen evolution) determined at 25 °C, (B) intrinsic PSII efficiency (dark Fv/Fm), and (C) maximal capacity of thermal energy dissipation (light-saturated NPQ, as F m =F 0m 1, determined in 2% O2, balance N2 at 25 °C) from leaves of Monstera deliciosa plants grown either under low non-fluctuating light (LL-grown under 10 lmol photons m2 s1; black columns) or in a sun-lit glasshouse (sun-grown with a midday peak PFD of 1500 lmol m2 s1; light gray columns). Means ± standard deviation (n = 3); statistically significant differences are indicated by asterisks (⁄⁄ = p < 0.01; ⁄⁄⁄ = p < 0.001; n.s. = not significantly different).


4


Fig. 2. Time course of chlorophyll fluorescence emission from leaves of Monstera deliciosa plants, grown either (A) under low non-fluctuating light (10 lmol photons m2 s1) or (B) in a sun-lit glasshouse (peak PFD of 1500 lmol m2 s1), that form the basis for the computation of maximal NPQ capacity shown in Fig. 1C (see legend of Fig. 1 for experimental details). Actinic light (PFD of 2000 lmol m2 s1) was switched on at minute 0. Fo, minimal fluorescence emission in darkness prior to experimental light exposure; F 0o , minimal fluorescence emission during actinic light exposure ascertained by rapid darkening of leaves; Fm, maximal fluorescence emission in darkness prior to experimental light exposure; F 0m , maximal fluorescence emission during actinic light exposure.

low-light-grown leaves exhibited NPQ levels comparable to those observed in leaves of high-light-exposed annuals [3], while sungrown Monstera leaves exhibited more than twice that NPQ level. Fig. 3 summarizes data obtained for a set of Monstera plants grown under identical conditions to those used with the plants for which results are shown in Figs. 1 and 2. Treatment with the uncoupler nigericin revealed that the rapidly reversible NPQ shown in Fig. 1 is pH-dependent (Fig. 3A). This strong NPQ in sun-grown leaves was, furthermore, accompanied by much greater levels of zeaxanthin and antheraxanthin (Z + A), as the de-epoxidized forms of the xanthophyll cycle consisting of violaxanthin + antheraxanthin + zeaxanthin, as well as much greater levels of the PsbS protein, in sun-grown compared to low-lightgrown leaves (Fig. 3B and C). Fig. 4 shows selected representative transmission–electronmicrographic images of grana stacks for chloroplasts from lowlight-grown (Fig. 4A–C) and sun-grown (Fig. 4D–F) leaves at three time points each, i.e., in darkness (dark; Fig. 4A and D), after 20 min of experimental exposure to high light (HL; Fig. 4B and E) in a temperature-controlled chamber, and after an additional 30-min exposure to low light (LL; Fig. 4C and F). The samples representing the state after 20 min in high light were cut into fixate while taking care to continuously expose them to high light. The total number of thylakoid membranes per granum was greater in chloroplasts from sun-grown versus low-light-grown leaves (Fig. 4). Fig. 5 shows the average number of thylakoid membranes per given height of a grana stack (per lm) for chloroplasts from low-lightgrown and sun-grown leaves in the three characterized states. In this evergreen species, the average number of thylakoid membranes per grana stack was similarly high in sun-grown versus low-light-grown leaves harvested predawn before exposure to light (Fig. 5). However, the average number of thylakoid membranes per grana stack was significantly lower in sun-grown versus low-light-grown leaves experimentally exposed to high light for 20 min and after a 30-min period in low light subsequent to experimental high-light exposure (Figs. 4 and 5). In response to high-light exposure, the thylakoid membranes were thus stacked more tightly in grana of chloroplasts from low-light-grown leaves and stacked more loosely (exhibited vertical unstacking under high-light exposure) in grana of chloroplast from sun-grown leaves (Fig. 4). Thus, vertical unstacking was seen only in the leaf exhibiting strong maximal NPQ, but not in low-light-grown leaves that exhibited much lower maximal NPQ. In response to brief high-light exposure, grana thylakoids thus remained tightly appressed in thylakoids from leaves of low-light-grown plants that exhibited a level of NPQ comparable to high-light-grown annual mesophytes [3,21], while the thylakoids of sun-grown Monstera leaves exhibited vertical unstacking along with an exceptionally high level of rapidly reversible, pH-dependent NPQ. 3.2. Thermal energy dissipation and thylakoid structure in low-lightgrown plants of Monstera transferred to high light and exhibiting strong sustained NPQ

Fig. 3. (A) The pH-dependent (nigericin-abolished) component of the capacity of thermal energy dissipation (light-saturated NPQ, as F m =F 0m 1 determined after a 20-min exposure to 2000 lmol photons m2 s1 in 2% O2, balance N2 at 25 °C), (B) maximal level of zeaxanthin + antheraxanthin (Z + A) on a chlorophyll basis in saturating light, and (C) level of the PsbS protein relative to the D1 protein of the PSII core for leaves of Monstera deliciosa plants grown either under low nonfluctuating light (LL-grown under 10 lmol photons m2 s1; black columns) or in a sun-lit glasshouse (sun-grown with a peak PFD of 1500 lmol m2 s1; light gray columns). Means ± standard deviation (n = 3); statistically significant differences are indicated by asterisks (⁄⁄ = p < 0.01; ⁄⁄⁄ = p < 0.001). Data from [21].

Monstera plants grown in a sun-lit glasshouse or grown under non-fluctuating low light were transferred to a temperature-controlled growth chamber under a 10-h photoperiod of 700 lmol photons m2 s1 (14 h dark) for five consecutive days, followed by lowering of the light level to 10 lmol photons m2 s1 (Fig. 6). Use of a heat filter (refrigerated circulating water) between the light source in the growth chambers and the plants allowed exposure to high light, while maintaining leaf temperatures at 25 °C (and thereby preventing the heat damage of leaves – in the form of necrosis – seen in the absence of the heat filter; data not shown). The intrinsic photon yield (at open traps) of PSII was ascertained at the end of the night (as Fv/Fm) and the effective photon



5

Fig. 4. Representative transmission electron micrographic images of grana stacks from chloroplasts of foliar palisade mesophyll cells for Monstera plants grown either in nonfluctuating low light (LL-grown; A–C) or in a sun-lit glasshouse (sun-grown; D–F) and examined at the end of the dark period (dark; A and D), after a 20 min experimental high light-treatment (HL, 1500 lmol photons m2 s1 at 25 °C) coupled with tissue fixation in HL (B and E), or following a subsequent 30-min exposure to low light (LL, 10 lmol photons m2 s1; C and F). All images are at the same scale (scale bar = 250 nm).

Fig. 5. Number of grana membranes per lm of grana height for leaves of Monstera deliciosa plants grown either under low non-fluctuating light (LL-grown under 10 lmol photons m2 s1) or in a sun-lit glasshouse (sun-grown with a midday peak PFD of 1500 lmol m2 s1) and measured at the end of the dark period (black bars), after a 20 min high light (HL) treatment with tissue fixation in HL (open bars), or following a subsequent 30-min exposure to low light (HL ? LL; gray bars). Seven to 15 grana stacks were measured from leaves of 3 plants for each condition. Means ± standard error. Statistically significant differences across all means indicated with different lower case letters.

efficiency of PSII (photon yield of PSII under the actual degree of trap closure) under growth-light conditions was determined at the end of the high-light period as ðF 0m FÞ=F 0m (which is the same as the product of intrinsic PSII photon yield at open traps in the light, F 0v =F 0m , and the actual degree of PSII trap closure in the light that can be assessed via the photochemical quenching coefficient [27]).

Fig. 6. Time course of changes in (A) intrinsic PSII efficiency at the end of the nocturnal dark period (black symbols) and effective PSII efficiency under actinic high-light (open symbols) or low-light (gray symbols) exposure and (B) levels of dark-sustained thermal energy dissipation (NPQ) from leaves of Monstera deliciosa plants grown either under low non-fluctuating light (circles; LL-grown under 10 lmol photons m2 s1) or in a sun-lit glasshouse (squares; sun-grown with a midday peak PFD of 1500 lmol m2 s1). Means ± standard deviation (n = 3 plants).


6


Fig. 6A shows that the effective PSII efficiency in leaves of sungrown plants decreased strongly during each light period, but returned to a high level during the dark period. On the other hand, effective PSII efficiency in leaves of low-light-grown plants decreased more strongly during photoperiods than in sun-grown plants, showed little recovery over the course of the 14-h dark periods, and also rose only slowly during the subsequent extended exposure to low light (Fig. 6A). Consistent with the patterns of intrinsic PSII photon efficiency in darkness (Fv/Fm), levels of darksustained thermal energy dissipation were high in low-lightgrown leaves and negligible in sun-grown leaves after each 14-h dark period (Fig. 6B). In other words, low-light-grown leaves exhibited continuously high levels of dark-sustained NPQ upon sudden experimental transfer to a high light environment, while NPQ in sun-grown leaves under the same experimental exposure was reversible in darkness. Fig. 7 illustrates that the structure of the thylakoid membranes in grana of chloroplasts from lowlight-grown leaves following three days of exposure to high light is reminiscent of the loose structure in grana of sun-grown leaves under high-light exposure. 3.3. Thylakoid structure in naturally overwintering conifers exhibiting photosynthetic inactivation and strong dark-sustained thermal energy dissipation The light- and CO2-saturated rate of photosynthesis (Fig. 8A and B) and the intrinsic efficiency of PSII in darkness (Fv/Fm; Fig. 8C and D) were high in the autumn, but very low in the winter, in needles of two subalpine conifers. Furthermore, the relative number of thylakoids present in grana stacks versus as single stromal lamellae (ratio of stacked versus unstacked membranes) in chloroplasts from needles of Engelmann spruce and subalpine fir was relatively high in the autumn (September) but very low in the winter (February) (Fig. 8E and F). Selected transmission electron micrographs of chloroplasts from Engelmann spruce needles show frequent small grana stacks (as well as starch grains) in the autumn (Fig. 9A and B) and predominantly unstacked thylakoid membranes in the winter (Fig. 9C and D).

Fig. 7. Representative transmission electron micrographic image of a granal stack from a chloroplast of foliar palisade mesophyll cells for Monstera plants grown in non-fluctuating low light (LL-grown) harvested at the end of the fourth dark period after a transfer to high light (700 lmol photons m2 s1) for 10 h per day. Scale bar = 500 nm.

Fig. 8. (A and B) Maximal photosynthetic capacity (light- and CO2-saturated rate of oxygen evolution), (C and D) intrinsic PSII efficiency (dark Fv/Fm), and (E and F) ratio of thylakoid membranes that are part of grana stacks versus unstacked stromal lamellae in chloroplasts from needles of Engelmann spruce (A, C, and E) and subalpine fir (B, D, and F) collected from sun-exposed locations in a Colorado subalpine forest in autumn (September) or winter (February). Means ± standard deviation (n = 4–5 plants); statistically significant differences are indicated by asterisks (⁄⁄⁄ = p < 0.001).

4. Discussion The findings presented here indicate a relationship between pronounced rearrangement of thylakoid structure (in the form of partial or complete unstacking of grana) and strong thermal energy dissipation in evergreens. Our observations differ from those made on the annual species spinach. The groundbreaking work of Murakami and Packer [17] reported no unstacking under high-light conditions, but instead a tighter appression. Similar observations have since been reported by other groups, including Johnson et al. [28] who concluded that, ‘‘[p]hotoinhibition and NPQ therefore appear to have opposite effects’’. Our results from evergreen species with their typical very high levels of thermal dissipation (estimated from NPQ) differ from the results obtained with annual species (that feature much lower maximal NPQ levels). We find strong reversible NPQ in non-photoinhibited evergreens as well as strong sustained NPQ in photoinhibited evergreens to be associated with partial to complete unstacking of grana. A loosening, or vertical unstacking, of thylakoids in grana was seen in chloroplasts from leaves of sun-grown Monstera plants exhibiting strong pHdependent thermal energy dissipation and from leaves exhibiting dark-sustained, pH-independent thermal energy dissipation upon transfer of low-light-grown Monstera plants to high light for 3 days (see also [21]). Dark-sustained, pH-independent thermal energy dissipation in needles of overwintering subalpine conifers (see also [30]), on the other hand, was accompanied by conversion of grana thylakoids to stromal lamellae. The fact that a very similar vertical unstacking of grana is seen in both photoinhibited and non-photoinhibited Monstera leaves indicates that vertical unstacking is a feature mechanistically associated with strong thermal energy dissipation but not necessarily with lasting PSII inactivation



Fig. 9. Representative transmission electron micrographic images of chloroplasts from needles of Engelmann spruce collected from sun-exposed locations in a subalpine forest in autumn (September; A and B) or winter (February; C and D) at two magnifications (A and C at 19,000; B and D at 34,000). Scale bars = 500 nm.

per se (cf. discussion in [9]). It is, furthermore, clear that only strong NPQ, and not an NPQ level of a more commonly seen magnitude, is associated with the vertical unstacking described here: In response to a brief (20-min-long) light exposure, no vertical unstacking of grana thylakoids was observed in chloroplasts from leaves of low-light-grown Monstera plants that exhibited NPQ levels comparable to those in high-light-exposed annuals [3,21]. What dynamic processes could be involved in such an unstacking of granal thylakoids? Several different types of thylakoid reorganizations have thus far been characterized. Chief among these previously characterized adjustments are (i) rearrangement of pigment-protein complexes between granal and stromal regions in the service of optimizing the utilization of limiting light levels for photosynthesis via ‘‘state shifts’’, balancing delivery of light absorbed by LHCII to either PSII or PSI and without net dissipation of excitation energy [11–13]. (ii) A shift in the fractions of excitation energy delivered to the two photosystems under excess light in drought-stressed leaves (involving decreased absorption by PSII, increased absorption by PSI, increased cyclic electron flow, and increased NPQ) was demonstrated by Zivak et al. [29]. (iii) With respect to thermal dissipation of excess light (assessed as NPQ), Betterle et al. [31], furthermore, provided direct evidence of an NPQ-related structural change in PSII involving redistribution of PSII core complexes within granal membranes when NPQ develops (reviewed by [15]). The focus of the latter discussion has been on reorganization of PSII and its associated pigment complexes into two major super-complexes, one consisting largely of PSII complexes with minor antenna proteins (Lhc4 and Lhc5) and with some LHCII and the other consisting mainly of LHCII complexes with the minor complex Lhc6 (see [32]). Ruban and Mullineaux [16] maintain that, ‘‘the fact that the NPQ-photoprotective state appears to require significant rearrangement of protein complexes in the grana raises the possibility that the mobility of chlorophyllproteins in the grana may be a major factor controlling the kinetics of NPQ formation.’’ While such rearrangements may occur within individual grana, Garab [33] stated that, ‘‘in unstacked thylakoid membranes more than 50% of protein complexes are mobile, whereas this number drops to about 20% in stacked grana regions [34]. Conversely, in order to allow dynamic reorganization in the lamellae, membranes might need to be unstacked.’’

7

Ruban and Mullineaux [16] furthermore pointed out that, ‘‘granal protein mobility may be significantly different in different plant species, and in differently acclimated plants.’’ As stated above, previous characterizations were conducted with annual species that grow rapidly throughout their short life cycle and exhibit very high maximal photosynthetic capacities along with very low maximal NPQ capacities [3,6,18–21]. In contrast, evergreens undergo multiple season-long periods of growth arrest over their long life span and, even during favorable climatic periods, feature rather low growth rates, low maximal capacities of photosynthesis and high maximal capacities for NPQ [3,6,18,20,21]. Strong thermal energy dissipation is associated with greater levels of both zeaxanthin and PsbS in Monstera grown in high light and exhibiting strong pH-dependent, reversible NPQ [21]. On the other hand, strong dark-sustained thermal energy dissipation – that is neither pH-dependent nor rapidly reversible – has been observed in (i) overwintering evergreens [2,3,18,19,21,23,25, 28,35–47] and (ii) shade-grown tropical evergreens suddenly transferred to high light environments for prolonged periods [21,48,49]. This strong energy dissipation – continuously maintained (even in low light or darkness) and pH-independent – is associated with high levels of continuously retained zeaxanthin and accumulation of members (ELIPs, OHPs, HLIPs) of the stressinducible sub-family of light-harvesting proteins to which PsbS belongs, but not of PsbS itself [20,21,28,45,47,50]. We have referred to leaves exhibiting high NPQ (or low Fv/Fm) that is continuously maintained for 24 h per day (throughout dark, lowand high-light periods) as having dark-sustained thermal energy dissipation [2,18,21,36,37,42,51]; this dark-sustained energy dissipation is associated with an arrest of conversions in the xanthophyll cycle (consisting of violaxanthin, antheraxanthin, and zeaxanthin) in the state of the high-light forms zeaxanthin and antheraxanthin. In leaves with dark-sustained thermal energy dissipation, both high NPQ levels (manifest as low Fv/Fm levels since NPQ cannot be calculated from such leaves; see [2,37,52–55]) and high zeaxanthin + antheraxanthin levels (similar to those under peak PFD under field conditions) remain high (are locked-in) 24 h per day [2,3,18,23,37–40,42–45,51,56,57]. The roles of PsbS and zeaxanthin in the reorganization of PSII into two super-complexes have been reviewed, e.g., by Horton [15] and Morosinotto and Bassi [32]. Ruban and Mullineaux [16] concluded that, ‘‘looser packing of PSII complexes in the presence of PsbS could facilitate protein mobility in the grana membranes, and this could, in turn, allow a much more rapid rearrangement of complexes following light exposure and the formation of a pH gradient across the thylakoid membrane. In this model, PsbS acts primarily as a ‘lubricant’ of the membrane, promoting flexibility and rapid adaptation by preventing PS II super-complexes from locking into a rigid macro-organization.’’ Horton [15; see also 32] emphasized ‘‘the controlling influence of VAZ and PsbS on this structural change.’’ The possibility that some form of interaction between PSII and PSI systems may be involved in strong thermal energy dissipation when absorbed light is highly excessive should also be considered. Interaction between PSII and PSI in experimentally unstacked isolated thylakoids has thus far been shown to involve photochemical quenching of excitation energy transferred from PSII to PSI; it was concluded that unstacking causes spillover of excitation energy from PSII to PSI (and increased fluorescence emission from PSI) and involves LHCII-LHCI-PSI super-complexes [58]. However, can spillover-like changes also involve non-photochemical quenching, i.e., NPQ? Interactions between PSI and PSII systems are clearly fostered by the high-energy-state under high light and involve a thylakoid lumen expansion triggered by the trans-thylakoid pH gradient and STN7-kinase-dependent phosphorylation of LHCII [59]. Furthermore, protonation of PsbS causes energy spillover to PSI [60].


8


Moreover, Ruban and Mullineaux [16] concluded that, ‘‘protein mobility is significantly modulated by physiological adaptation. For example, mobility increases following photoinhibition . . ., and this effect is dependent on the presence of the Stn7 and Stn8 protein kinases [61,62].’’ Our results suggest that the very high levels of NPQ in non-photoinhibited evergreens may require similar changes. Species-dependent differences exist in thylakoid protein-phosphorylation patterns in response to high light. Unlike the annual, herbaceous species soybean, the evergreen, sclerophytic species Monstera maintained LHCII phosphorylation in high light [63]. Continuously high, dark-sustained energy dissipation in the overwintering conifer Douglas fir was, furthermore, associated with sustained phosphorylation of D1 and other PSII core proteins and downregulation of a protein phosphatase [25,45]. Likewise, dark-sustained energy dissipation in low-light-grown Monstera transferred to high light was associated with sustained D1 phosphorylation [44]. May a form of spillover quenching in PSI that is associated with NPQ and PsbS also involve zeaxanthin? Conversion of violaxanthin to zeaxanthin has been demonstrated in LHCI [64], and zeaxanthin quenches PSI-LHCI fluorescence in a photoprotective process [65]. 30% of the xanthophyll cycle pool in cotton leaves is located in PSIassociated pigment complexes [66]. We suggest that facile conversion of violaxanthin to zeaxanthin in LHCI and strong NPQ in LHCI could be a distinguishing feature between annual and evergreen species. While annual species growing in full sun maximally convert about 70% of their xanthophyll cycle pool under daily exposure to peak irradiance and show only modest levels of NPQ, evergreens convert close to 100% of the xanthophyll cycle pool to zeaxanthin and show strong NPQ on a daily basis. Although the NPQ data presented in the current report are derived from roomtemperature fluorescence that emanates mainly from PSII, we have previously shown that high-light-induced NPQ involves concomitant, proportional non-photochemical quenching of PSII and PSI fluorescence measured from frozen (77 K) leaf tissues of Monstera [48], several other evergreens [67,68], and the annual species soybean [63]. The findings presented here indicate a relationship between strong thermal energy dissipation and rearrangements of thylakoid structure that differ not only between annual and evergreen species, but also among evergreen species. While loosening, or vertical unstacking, of thylakoids in grana was seen in Monstera as a species with remarkably tall grana stacks irrespective of growth light environment, chloroplasts of sun-exposed needles from subalpine confers did not exhibit very tall grana stacks in either autumn or winter, and their transition from autumn to winter was accompanied by conversion of most thylakoids from granal to stromal lamellae. We propose that the differences seen in thylakoid structural changes of different evergreen species depend on the growth light environments to which these species are adapted – sclerophytic tropical evergreens with very tall grana stacks are well adapted to deep-shade environments and subalpine evergreens with few grana stacks are adapted to high-light environments with pronounced excess light over entire winter seasons. As a climbing vine (and hemi-epiphyte), Monstera begins its life cycle in deep shade on a forest floor and later climbs to the forest canopy where it tolerates full sunlight presumably by virtue of employing strong thermal energy dissipation (see discussion in [21]). In conclusion, evergreen species exhibit unique features in that they maintain green leaves or needles throughout periods of active growth as well as periods of growth arrest, and in that they exhibit exceptionally strong thermal dissipation of excess absorbed light accompanied by pronounced thylakoid structural rearrangements that differ in their specific manifestation between tropical evergreens and temperate conifers.

As stated above, vertical thylakoid unstacking in commonly studied annual species has been discussed in the context of lasting photoinhibitory inactivation of PSII (see, e.g., [9,28]). We speculate that the highly excessive light levels commonly experienced by evergreens even during periods of active growth are experienced by annuals only under conditions strongly inhibiting these species’ rapid growth and high electron transport rates. We assume that the latter conditions will lead to simultaneous PSII inactivation, chlorophyll degradation, and enhanced NPQ in annual species, thus complicating a dissection of the features of PSII inactivation from those of NPQ in annuals. In reviewing PSII inactivation in the context of the whole plant in its natural environment, we previously also pointed out that photoinhibition in evergreens involves carbohydrate (starch and sugar) accumulation in leaves along with strong, continuously maintained thermal energy dissipation and long-term arrest of xanthophyll cycle conversion [2,51,57,69]. We proposed that a combination of sugar signaling and redox signaling actively induces PSII inactivation and/or removal of PSII cores and oxygen-evolving complexes in order to limit reactive-oxygen formation and to facilitate strong thermal dissipation of all light absorbed in remaining chlorophyllprotein complexes – as a prerequisite to the maintenance of green leaves in (evergreen) plants throughout periods of season-long growth arrest [2,40,51,57,70,71].

Acknowledgments Supported by the National Science Foundation (Award Numbers IOS-0841546 and DEB-1022236 to B.D.-A. and W.W.A.) and the University of Colorado at Boulder. We also thank Emmo Scherbatskoy for assistance with the anatomical characterizations.

References [1] W.W. Adams III, B. Demmig-Adams, B.A. Logan, D.H. Barker, C.B. Osmond, Rapid changes in xanthophyll cycle-dependent energy dissipation and photosystem II efficiency in two vines, Stephania japonica and Smilax australis, growing in the understory of an open Eucalyptus forest, Plant Cell Environ. 22 (1999) 125–136, http://dx.doi.org/10.1046/j.13653040.1999.00369.x. [2] W.W. Adams III, C.R. Zarter, K.E. Mueh, V. Amiard, B. Demmig-Adams, Energy dissipation and photoinhibition: a continuum of photoprotection, in: B. Demmig-Adams, W.W. Adams III, A.K. Mattoo (Eds.), Photoprotection, Photoinhibition, Gene Regulation, and Environment, Advances in Photosynthesis and Respiration, vol. 21, Springer, Dordrecht, 2006, pp. 49– 64, http://dx.doi.org/10.1007/1-4020-3579-9_5. [3] B. Demmig-Adams, C.M. Cohu, O. Muller, W.W. Adams III, Modulation of photosynthetic energy conversion efficiency in nature: from seconds to seasons, Photosynth. Res. 113 (2012) 75–88, http://dx.doi.org/10.1007/ s11120-012-9761-6. [4] B. Demmig-Adams, W.W. Adams III, The role of xanthophyll cycle carotenoids in the protection of photosynthesis, Trends Plant Sci. 1 (1996) 21–26, http:// dx.doi.org/10.1016/S1360-1385(96)80019-7. [5] B. Demmig-Adams, W.W. Adams III, Xanthophyll cycle and light stress in nature: uniform response to excess direct sunlight among higher plant species, Planta 198 (1996) 460–470, http://dx.doi.org/10.1007/BF00620064. [6] B. Demmig-Adams, W.W. Adams III, D.H. Barker, B.A. Logan, A.S. Verhoeven, D.R. Bowling, Using chlorophyll fluorescence to assess the allocation of absorbed light to thermal dissipation of excess excitation, Physiol. Plant. 98 (1996) 253–264, http://dx.doi.org/10.1034/j.1399-3054.1996.980206.x. [7] B. Andersson, J.M. Anderson, Lateral heterogeneity in the distribution of chlorophyll-protein complexes of the thylakoid membranes of spinach chloroplasts, Biochim. Biophys. Acta 593 (1980) 427–440, http://dx.doi.org/ 10.1016/0005-2728(80)90078-X. [8] J.M. Anderson, W.S. Chow, D.J. Goodchild, Thylakoid membrane organization in sun/shade acclimation, Aust. J. Plant Physiol. 15 (1988) 11–26, http:// dx.doi.org/10.1071/PP9880011. [9] M. Pribil, M. Labs, D. Leister, Structure and dynamics of thylakoids in land plants, J. Exp. Bot. 65 (2014) 1955–1972, http://dx.doi.org/10.1093/jxb/eru090. [10] W.S. Chow, L. Qian, D.J. Goodchild, J.M. Anderson, Photosynthetic acclimation of Alocasia macrorrhiza (L.) G. Don to growth irradiance: structure, function and composition of the chloroplasts, Aust. J. Plant Physiol. 15 (1988) 107–122, http://dx.doi.org/10.1071/PP9880107.


B. Demmig-Adams et al. / Journal of Photochemistry and Photobiology B: Biology xxx (2015) xxx–xxx [11] W.P. Williams, J.F. Allen, State 1/State 2 changes in higher plants and algae, Photosynth. Res. 13 (1987) 19–45, http://dx.doi.org/10.1007/BF00032263. [12] J.F. Allen, Protein phosphorylation in regulation of photosynthesis, Biochim. Biophys. Acta 1098 (1992) 275–335, http://dx.doi.org/10.1016/S00052728(09)91014-3. [13] G.H. Krause, P. Jahns, Non-photochemical energy dissipation determined by chlorophyll fluorescence quenching: characterization and function, in: G.C. Papageorgiou, Govindjee (Eds.), Chlorophyll a Fluorescence: A Signature of Photosynthesis, Advances in Photosynthesis and Respiration, vol. 19, Springer, Dordrecht, 2004, pp. 463–495, http://dx.doi.org/10.1007/978-1-4020-32189_18. [14] M. Edelman, A.K. Mattoo, The D1 protein past and future perspectives, in: B. Demmig-Adams, W.W. Adams III, A.K. Mattoo (Eds.), Photoprotection, Photoinhibition, Gene Regulation, and Environment, Advances in Photosynthesis and Respiration, vol. 21, Springer, Dordrecht, 2006, pp. 23– 38, http://dx.doi.org/10.1007/1-4020-3579-9_2. [15] P. Horton, Developments in research on non-photochemical fluorescence quenching: emergence of key ideas, theories and experimental approaches, in: B. Demmig-Adams, G. Garab, W.W. Adams III, Govindjee (Eds.), NonPhotochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 73–95, http://dx.doi.org/10.1007/978-94-017-9032-1_3. [16] A.V. Ruban, C.W. Mullineaux, Non-photochemical fluorescence quenching and the dynamics of photosystem II structure, in: B. Demmig-Adams, G. Garab, W.W. Adams III, Govindjee (Eds.), Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 373– 386, http://dx.doi.org/10.1007/978-94-017-9032-1_17. [17] S. Murakami, L. Packer, Protonation and chloroplast membrane structure, J. Cell Biol. 47 (1970) 332–351. [18] B. Demmig-Adams, V. Ebbert, C.R. Zarter, W.W. Adams III, Characteristics and species-dependent employment of flexible versus sustained thermal dissipation and photoinhibition, in: B. Demmig-Adams, W.W. Adams III, A.K. Mattoo (Eds.), Photoprotection, Photoinhibition, Gene Regulation, and Environment, Advances in Photosynthesis and Respiration, vol. 21, Springer, Dordrecht, 2006, pp. 39–48, http://dx.doi.org/10.1007/1-4020-3579-9_4. [19] W.W. Adams III, B. Demmig-Adams, Lessons from nature: a personal perspective, in: B. Demmig-Adams, G. Garab, W. Adams III, Govindjee (Eds.), Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 45–72, http://dx.doi.org/10.1007/978-94-017-9032-1_2. [20] B. Demmig-Adams, S.C. Koh, C.M. Cohu, O. Muller, J.J. Stewart, W.W. Adams III, Non-photochemical quenching in contrasting plant species and environments, in: B. Demmig-Adams, G. Garab, W.W. Adams III, Govindjee (Eds.), NonPhotochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 531–552, http://dx.doi.org/10.1007/978-94-017-90321_24. [21] B. Demmig-Adams, V. Ebbert, D.L. Mellman, K.E. Mueh, L. Schaffer, C. Funk, C.R. Zarter, I. Adamska, S. Jansson, W.W. Adams III, Modulation of PsbS and flexible versus sustained energy dissipation by light environment in different species, Physiol. Plant. 127 (2006) 670–680, http://dx.doi.org/10.1111/j.13993054.2006.00698.x. [22] T. Delieu, D.A. Walker, Polarographic measurement of photosynthetic oxygen evolution by leaf disks, New Phytol. 86 (1981) 165–178, http://dx.doi.org/ 10.1111/j.1469-8137.1981.tb07480.x. [23] W.W. Adams III, B. Demmig-Adams, T.N. Rosenstiel, A.K. Brightwell, V. Ebbert, Photosynthesis and photoprotection in overwintering plants, Plant Biol. 4 (2002) 545–557, http://dx.doi.org/10.1055/s-2002-35434. [24] W.W. Adams III, B. Demmig-Adams, Operation of the xanthophyll cycle in higher plants in response to diurnal changes in incident sunlight, Planta 186 (1992) 390–398, http://dx.doi.org/10.1007/BF00195320. [25] V. Ebbert, W.W. Adams III, A.K. Mattoo, A. Sokolenko, B. Demmig-Adams, Upregulation of a PSII core protein phosphatase inhibitor and sustained D1 phosphorylation in zeaxanthin-retaining, photoinhibited needles of overwintering Douglas fir, Plant Cell Environ. 28 (2005) 232–240, http:// dx.doi.org/10.1111/j.1365-3040.2004.01267.x. [26] X.-P. Li, A. Phippard, J. Pasari, K.K. Niyogi, Structure-function analysis of photosystem II subunit S (PsbS) in vivo, Funct. Plant Biol. 29 (2002) 1131– 1139, http://dx.doi.org/10.1071/FP02065. [27] B. Genty, J.M. Briantais, N.R. Baker, The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence, Biochim. Biophys. Acta 990 (1989) 87–92, http://dx.doi.org/ 10.1016/S0304-4165(89)80016-9. [28] M.P. Johnson, T.K. Goral, C.D.P. Duffy, A.P.R. Brain, C.W. Mullineaux, A.V. Ruban, Photoprotective energy dissipation involves the reorganization of photosystem II light-harvesting complexes in the grana membranes of spinach chloroplasts, Plant Cell 23 (2011) 1468–1479, http://dx.doi.org/ 10.1105/tpc.110.081646. [29] M. Zivak, M. Brestic, Z. Balatova, P. Drevenkova, K. Olsovska, H.M. Kaalji, X Yang, S.I. Allakhverdiev, Photosynthetic electron transport and specific photoprotective responses in wheat leaves under drought, Photosynth. Res. 117 (2013) 529–546, http://dx.doi.org/10.1007/s11120-013-9885-3. [30] C.R. Zarter, W.W. Adams III, V. Ebbert, D. Cuthbertson, I. Adamska, B. DemmigAdams, Winter down-regulation of intrinsic photosynthetic capacity coupled with up-regulation of Elip-like proteins and persistent energy dissipation in a

[31]

[32]

[33]

[34]

[35]

[36]

[37]

[38]

[39]

[40]

[41]

[42]

[43]

[44]

[45]

[46]

[47]

[48]

[49]

[50]

9

subalpine forest, New Phytol. 172 (2006) 272–282, http://dx.doi.org/10.1111/ j.1469-8137.2006.01815.x. N. Betterle, M. Ballattori, S. Zorzan, S. de Bianchi, S. Cazzaniga, L. Dall’Osto, T. Morosinotto, R. Bassi, Light-induced dissociation of an antenna heterooligomer is needed for non-photochemical quenching induction, J. Biol. Chem. 284 (2009) 15255–15266, http://dx.doi.org/10.1074/jbc.M808625200. T. Morosinotto, R. Bassi, Molecular mechanisms for activation of nonphotochemical fluorescence quenching: from unicellular algae to mosses and higher plants, in: B. Demmig-Adams, G. Garab, W.W. Adams III, Govindjee (Eds.), Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 315–331, http://dx.doi.org/10.1007/ 978-94-017-9032-1_14. G. Garab, Structural changes and non-photochemical quenching of chlorophyll a fluorescence in oxygenic photosynthetic organisms, in: B. Demmig-Adams, G. Garab, W.W. Adams III, Govindjee (Eds.), Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 343– 371, http://dx.doi.org/10.1007/978-94-017-9032-1_16. H. Kirchhoff, R.M. Sharpe, M. Herbstová, R. Yarbrough, G.E. Edwards, Differential mobility of pigment-protein complexes in granal and agranal thylakoid membranes of C3 and C4 plants, Plant Physiol. 161 (2013) 497–507, http://dx.doi.org/10.1104/pp.112.207548. W.W. Adams III, B. Demmig-Adams, Carotenoid composition and down regulation of photosystem II in three conifer species during the winter, Physiol. Plant. 92 (1994) 451–458, http://dx.doi.org/10.1034/j.1399-3054. 1994.920313.x. W.W. Adams III, B. Demmig-Adams, The xanthophyll cycle and sustained thermal energy dissipation activity in Vinca minor and Euonymus kiautschovicus in winter, Plant Cell Environ. 18 (1995) 117–127, http:// dx.doi.org/10.1111/j.1365-3040.1995.tb00345.x. W.W. Adams III, B. Demmig-Adams, A.S. Verhoeven, D.H. Barker, ‘Photoinhibition’ during winter stress: involvement of sustained xanthophyll cycle-dependent energy dissipation, Aust. J. Plant Physiol. 22 (1995) 261–276, http://dx.doi.org/10.1071/PP9950261. W.W. Adams III, D.H. Barker, Seasonal changes in xanthophyll cycle-dependent energy dissipation in Yucca glauca Nuttall, Plant Cell Environ. 21 (1998) 501– 512, http://dx.doi.org/10.1046/j.1365-3040.1998.00283.x. W.W. Adams III, B. Demmig-Adams, T.N. Rosenstiel, V. Ebbert, Dependence of photosynthesis and energy dissipation activity upon growth form and light environment during the winter, Photosynth. Res. 67 (2001) 51–62, http:// dx.doi.org/10.1023/A:1010688528773. W.W. Adams III, C.R. Zarter, V. Ebbert, B. Demmig-Adams, Photoprotective strategies of overwintering evergreens, Bioscience 54 (2004) 41–49, http:// dx.doi.org/10.1641/0006-3568(2004)054[0041:PSOOE]2.0.CO;2. A.S. Verhoeven, W.W. Adams III, B. Demmig-Adams, Close relationship between the state of the xanthophyll cycle pigments and photosystem II efficiency during recovery from winter stress, Physiol. Plant. 96 (1996) 567– 576, http://dx.doi.org/10.1111/j.1399-3054.1996.tb00228.x. A.S. Verhoeven, W.W. Adams III, B. Demmig-Adams, Two forms of sustained xanthophyll cycle-dependent energy dissipation in overwintering Euonymus kiautschovicus, Plant Cell Environ. 21 (1998) 893–903, http://dx.doi.org/ 10.1046/j.1365-3040.1998.00338.x. A.S. Verhoeven, W.W. Adams III, B. Demmig-Adams, The xanthophyll cycle and acclimation of Pinus ponderosa and Malva neglecta to winter stress, Oecologia 118 (1999) 277–287, http://dx.doi.org/10.1007/s004420050728. V. Ebbert, B. Demmig-Adams, W.W. Adams III, K.E. Mueh, L.A. Staehelin, Correlation between persistent forms of zeaxanthin-dependent energy dissipation and thylakoid protein phosphorylation, Photosynth. Res. 67 (2001) 63–78, http://dx.doi.org/10.1023/A:1010640612843. B. Demmig-Adams, W.W. Adams III, Photoprotection in an ecological context: the remarkable complexity of thermal energy dissipation, New Phytol. 172 (2006) 11–21, http://dx.doi.org/10.1111/j.1469-8137.2006.01835.x. C.R. Zarter, B. Demmig-Adams, V. Ebbert, I. Adamska, W.W. Adams III, Photosynthetic capacity and light harvesting efficiency during the winter-tospring transition in subalpine conifers, New Phytol. 172 (2006) 283–292, http://dx.doi.org/10.1111/j.1469-8137.2006.01816.x. C.R. Zarter, W.W. Adams III, V. Ebbert, I. Adamska, S. Jansson, B. DemmigAdams, Winter acclimation of PsbS and related proteins in the evergreen Arctostaphylos uva-ursi as influenced by altitude and light environment, Plant Cell Environ. 29 (2006) 869–878, http://dx.doi.org/10.1111/j.13653040.2005.01466.x. B. Demmig, O. Björkman, Comparison of the effect of excessive light on chlorophyll fluorescence (77K) and photon yield of O2 evolution in leaves of higher plants, Planta 171 (1987) 171–184, http://dx.doi.org/10.1007/BF003 91092. B. Demmig-Adams, D.L. Moeller, B.A. Logan, W.W. Adams III, Positive correlation between levels of retained zeaxanthin plus antheraxanthin and degree of photoinhibition in shade leaves of Schefflera arboricola (Hayata) Merrill, Planta 205 (1998) 367–374, http://dx.doi.org/10.1007/s0042500 50332. X. Wang, Y. Peng, J.W. Singer, A. Fessehaie, S.L. Krebs, R. Arora, Seasonal changes in photosynthesis, antioxidant systems, and ELIP expression in a thermonastic and non-thermonastic Rhododendron species: a comparison of photoprotective strategies in overwintering plants, Plant Sci. 177 (2009) 607– 617, http://dx.doi.org/10.1016/j.plantsci.2009.08.009.


10


[51] W.W. Adams III, O. Muller, C.M. Cohu, B. Demmig-Adams, May photoinhibition be a consequence, rather than a cause, of limited plant productivity?, Photosynth Res. 117 (2013) 31–44, http://dx.doi.org/10.1007/s11120-0139849-7. [52] W.W. Adams III, A. Hoehn, B. Demmig-Adams, Chilling temperatures and the xanthophyll cycle. A comparison of warm-grown and overwintering spinach, Aust. J. Plant Physiol. 22 (1995) 75–85, http://dx.doi.org/10.1071/PP9950075. [53] W.W. Adams III, B. Demmig-Adams, Chlorophyll fluorescence as a tool to monitor plant response to the environment, in: G.C. Papageorgiou, Govindjee (Eds.), Chlorophyll a Fluorescence: A Signature of Photosynthesis, Advances in Photosynthesis and Respiration, vol. 19, Springer, Dordrecht, 2004, pp. 583– 604, http://dx.doi.org/10.1007/978-1-4020-3218-9_22. [54] B.A. Logan, W.W. Adams III, B. Demmig-Adams, Avoiding common pitfalls of chlorophyll fluorescence analysis under field conditions, Funct. Plant Biol. 34 (2007) 853–859, http://dx.doi.org/10.1071/FP07113. [55] B.A. Logan, B. Demmig-Adams, W.W. Adams III, W. Bilger, Context, quantification, and measurement guide for non-photochemical quenching of chlorophyll fluorescence, in: B. Demmig-Adams, G. Garab, W.W. Adams III, Govindjee (Eds.), Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 187–201, http://dx.doi.org/10.1007/ 978-94-017-9032-1_7. [56] B. Demmig-Adams, W.W. Adams III, V. Ebbert, B.A. Logan, Ecophysiology of the xanthophyll cycle, in: H.A. Frank, A.J. Young, G. Britton, R.J. Cogdell (Eds.), The Photochemistry of Carotenoids, Advances in Photosynthesis, vol. 8, Kluwer Academic Publishers, Dordrecht, 1999, pp. 245–269, http://dx.doi.org/ 10.1007/0-306-48209-6_14. [57] W.W. Adams III, O. Muller, C.M. Cohu, B. Demmig-Adams, Photosystem II efficiency and non-photochemical quenching in the context of source-sink balance, in: B. Demmig-Adams, G. Garab, W.W. Adams III, Govindjee (Eds.), Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, Advances in Photosynthesis and Respiration, vol. 40, Springer, Dordrecht, 2014, pp. 503–529, http://dx.doi.org/10.1007/978-94-017-90321_23. [58] C.D. van der Weij-de Wit, J.A. Ihalainen, R. van Grondelle, J.P. Dekker, Excitation energy transfer in native and unstacked thylakoid membranes studied by low temperature and ultrafast fluorescence spectroscopy, Photosynth. Res. 93 (2007) 173–182, http://dx.doi.org/10.1007/s11120-0079157-1. [59] C.H. Clausen, M.D. Brooks, T.D. Li, P. Grob, G. Kemalyan, E. Nogales, K.K. Niyogi, D.A. Fletcher, Dynamic mechanical responses of Arabidopsis thylakoid membranes during PSII-specific illumination, Biophys. J. 106 (2014) 1864– 1870, http://dx.doi.org/10.1016/j.bpj.2014.03.016. [60] A. Jajoo, N.R. Mekala, T. Tongra, A. Tiwari, M. Grieco, M. Tikkanen, E.-M. Aro, Low pH-induced regulation of excitation energy between the two

[61]

[62]

[63]

[64]

[65]

[66]

[67]

[68]

[69]

[70]

[71]

photosystems, FEBS Lett. 588 (2014) 970–974, http://dx.doi.org/10.1016/ j.febslet.2014.01.056. T.K. Goral, M.P. Johnson, A.P.R. Brain, H. Kirchhoff, A.V. Ruban, C.W. Mullineaux, Visualizing the mobility and distribution of chlorophyll-proteins in higher plant thylakoid membranes: effects of photoinhibition and protein phosphorylation, Plant J. 62 (2010) 948–959, http://dx.doi.org/10.1111/j.1365313X.2010.04207.x. M. Herbstová, S. Tietz, C. Kinzel, M.V. Turkina, H. Kirchhoff, Architectural switch in plant photosynthetic membranes induced by light stress, Proc. Natl. Acad. Sci. USA 109 (2012) 20130–20135, http://dx.doi.org/10.1073/pnas. 1214265109. B. Demmig, R.E. Cleland, O. Björkman, Photoinhibition, 77K chlorophyll fluorescence quenching and phosphorylation of the light-harvesting chlorophyll-protein complex of photosystem II in soybean leaves, Planta 172 (1987) 378–385, http://dx.doi.org/10.1007/BF00398667. A. Wehner, S. Storf, P. Jahns, V.H.R. Schmid, De-epoxidation of violaxanthin in light-harvesting complex I proteins, J. Biol. Chem. 279 (2004) 26823–26829, http://dx.doi.org/10.1074/jbc.M402399200. M. Ballottari, M.J.P. Alcocer, C. D’Andrea, D. Viola, T.K. Ahn, A. Petrozza, D. Polli, F.R. Fleming, G. Cerullo, R. Bassi, Regulation of photosystem I light harvesting by zeaxanthin, Proc. Natl. Acad. Sci. USA 111 (2014) E2431–E2438, http:// dx.doi.org/10.1073/pnas.1404377111. S.S. Thayer, O. Björkman, Carotenoid distribution and deepoxidation in thylakoid pigment-protein complexes from cotton leaves and bundle-sheath cells of maize, Photosynth. Res. 33 (1992) 213–225, http://dx.doi.org/10.1007/ BF00030032. W.W. Adams III, I. Terashima, E. Brugnoli, B. Demmig, Comparisons of photosynthesis and photoinhibition in the CAM vine Hoya australis and several C3 vines growing on the coast of eastern Australia, Plant Cell Environ. 11 (1988) 173–181, http://dx.doi.org/10.1111/j.1365-3040.1988.tb01134.x. O. Björkman, B. Demmig, T.J. Andrews, Mangrove photosynthesis: response to high-irradiance stress, Aust. J. Plant Physiol. 15 (1988) 43–61, http:// dx.doi.org/10.1071/PP9880043. W.W. Adams III, A.M. Watson, K.E. Mueh, V. Amiard, R. Turgeon, V. Ebbert, B.A. Logan, A.F. Combs, B. Demmig-Adams, Photosynthetic acclimation in the context of structural constraints to carbon export from leaves, Photosynth. Res. 94 (2007) 455–466, http://dx.doi.org/10.1007/s11120-006-9123-3. B. Demmig-Adams, J.J. Stewart, W.W. Adams III, Multiple feedbacks between chloroplast and whole plant in the context of plant adaptation and acclimation to the environment, Philos. Trans. R. Soc. B 269 (2014) 20130244, http:// dx.doi.org/10.1098/rstb.2013.0244. B. Demmig-Adams, J.J. Stewart, T.A. Burch, W.W. Adams III, Insights from placing photosynthetic light harvesting into context, J. Phys. Chem. Lett. 5 (2014) 2880–2889, http://dx.doi.org/10.1021/jz5010768.


Phosphorylation of thylakoid proteins during chloroplast biogenesis in greening etiolated and light-grown wheat leaves.

Photothermal beam deflection: a new method for in vivo measurements of thermal energy dissipation and photochemical energy conversion in intact leaves.

Live-cell visualization of excitation energy dynamics in chloroplast thylakoid structures.

Thylakoid protein kinase activity and associated control of excitation energy distribution during chloroplast biogenesis in wheat.

Dissipation of excess excitation energy of the needle leaves in Pinus trees during cold winters.

A Putative Chloroplast Thylakoid Metalloprotease VIRESCENT3 Regulates Chloroplast Development in Arabidopsis thaliana.

Energy dissipation in flows through curved spaces.

Does the touch of cold make evergreen leaves tougher?

Energy dissipation in multifrequency atomic force microscopy.

The action of proteolytic enzymes on chloroplast thylakoid membranes.

Physiological effects of sublethal atrazine on barley chloroplast thylakoid membranes.

On the molecular nature of chloroplast thylakoid membranes.

Vectorial discharge of nascent polypeptides attached to chloroplast thylakoid membranes.

Chloroplast thylakoid membrane fluidity and its sensitivity to temperature.

The relationship between maximum tolerated light intensity and photoprotective energy dissipation in the photosynthetic antenna: chloroplast gains and losses.

The thermal conductivity of leaves.

Cell size and chloroplast size in relation to chloroplast replication in light-grown wheat leaves.

Reversibility and energy dissipation in adiabatic superconductor logic.

Energy dissipation in human hand-arm exposed to random vibration.

Energy dissipation and noise correlations in biochemical sensing.

Energy Dissipation Pathways in Few-Layer MoS2 Nanoelectromechanical Systems.

Chloroplast DNA in mature and senescing leaves: a reappraisal.

Intercalated water layers promote thermal dissipation at bio-nano interfaces.

Structural development and energy dissipation in simulated silicon apices.