Supplemental lighting can improve the profitability of greenhouse crop production, and a better quantitative understanding of plant responses to PPFD can facilitate the development of more efficient crop-specific control strategies for greenhouse supplemental lighting (van Iersel, 2017). Chlorophyll fluorescence measurements are a rapid and reliable means of probing the light reactions of photosynthesis directly (Baker, 2008). During the light reactions, some of the light energy absorbed by chlorophylls and accessory pigments migrates to photosystem II (PSII) reaction centers, resulting in the splitting of water molecules, liberating electrons and protons. The freed electrons are used to regenerate nicotinamide adenine dinucleotide phosphate hydrogen (NADPH) via the electron transport chain, and a proton gradient across the thylakoid membrane drives adenosine triphosphate (ATP) synthase, regenerating ATP. These energy-rich molecules—NADPH and ATP—provide the reducing power and chemical energy for carbohydrate production in the Calvin-Benson-Bassham cycle. However, not all light absorbed by photosynthetic pigments is used to drive the light reactions; a significant amount is dissipated as heat, and a small fraction is reemitted as fluorescence. By measuring the fluorescence emitted by chlorophyll a molecules before and during short exposure to a beam of light with sufficient intensity to saturate the PSII reaction centers completely (a “saturating pulse”), ΦPSII can be quantified directly. ΦPSII is a unitless measure of the efficiency with which absorbed light is used to drive photochemistry in the light-adapted state of PSII. The dark-adapted value of the quantum efficiency of PSII (Fv/Fm) is an indicator of maximum potential photochemical efficiency. Combined with PPFD, ΦPSII is used to calculate the rate of linear electron transport through PSII (the ETR), an estimate of the overall rate of the light reactions of photosynthesis (Baker and Rosenqvist, 2004; Genty et al., 1989; Maxwell and Johnson, 2000). To distinguish measurements based on chlorophyll fluorescence from other measures of photosynthesis such as gas exchange or oxygen evolution, data related to ΦPSII and ETR are referred to as photochemical rather than photosynthetic herein.
Chlorophyll fluorescence is an ideal tool for understanding crop-specific photochemical responses to PPFD. Chlorophyll fluorometers are generally small and portable, with simple operation that requires no recalibration. Measurements can be collected quickly in situ, and are noninvasive and accurate (Baker and Rosenqvist, 2004; Maxwell and Johnson, 2000). An exact correlation between ETR and CO2 fixation rates may be difficult to establish because the products of the light reactions can be used to drive processes other than the Calvin-Benson-Bassham cycle. Photorespiration is a major sink for NADPH and ATP in C3 plants (Krall and Edwards, 1992), and NADPH may be used as an electron donor for nitrate reduction (Tischner, 2000). Freed electrons may reduce O2 at photosystem I (Mehler reaction, or water–water cycle) rather than be used to produce NAPDH (Polle, 1996), and ATP can be used for chloroplast functions such as protein repair and nucleotide metabolism (Murata and Nishiyama, 2018; Spetea et al., 2004). Thus, the relationship between ETR and CO2 fixation depends on many factors, including temperature, relative humidity, CO2 concentration, and water and nutrient availability. However, ETR can be taken as a relative indicator of overall photosynthetic rates, and hence plant growth. Furthermore, compared with gas exchange, ETR of C3 plants is relatively insensitive to changes in environmental variables other than light (Murchie and Lawson, 2013). Thus, chlorophyll fluorescence measurements provide a convenient, rapid, accurate, and robust means of evaluating photochemical responses to PPFD.
Light response curves collected using chlorophyll fluorescence measurements are typically performed over a relatively brief period (often just a few minutes) with a highly focused light source and may not represent photochemical responses accurately under variable ambient light conditions (Rascher et al., 2000). Photoprotective processes affect photochemical light use efficiency by reducing the amount of absorbed light energy transferred to PSII reaction centers, and may operate over longer timescales. Because the accumulation of excess light energy in the light-harvesting complexes can lead to light-induced damage of PSII reaction centers (photoinhibition), plants have evolved a variety of interrelated photoprotective mechanisms by which excess absorbed light energy can be dissipated safely as heat, including molecular reorganization of PSII and the xanthophyll cycle (Demmig-Adams et al., 2012; Horton, 2012; Rochaix, 2014; Ruban, 2015). As PPFD increases, a larger fraction of absorbed light is dissipated as heat, resulting in a decrease in ΦPSII (Baker, 2008; Maxwell and Johnson, 2000). Fluctuations in PPFD throughout the course of a day can lead to variations in ΦPSII as a result of the up- or downregulation of the xanthophyll cycle. The xanthophyll cycle is the process by which the accumulation of protons leads to acidification of the thylakoid lumen, activating violaxanthin de-epoxidase, which catalyzes the de-epoxidation of violaxanthin to form antheraxanthin and zeaxanthin. This chemical conversion of the xanthophyll pigments facilitates the dissipation of excess light energy as heat. It reverses relatively slowly, over a scale of several minutes, through epoxidation catalyzed by zeaxanthin epoxidase. Because of this slow relaxation, transient exposure to high light levels may lead to decreases in photochemical efficiency (relative decreases in ΦPSII and ETR) for several minutes even if PPFDs subsequently decrease to much lower levels (Demmig-Adams et al., 2012; Kaiser et al., 2018; Ruban, 2015). Photochemistry-induced acidification of the thylakoid lumen can further affect rates of electron transport by inhibiting plastohydroquinone oxidation by the cytochrome b6f complex, thereby decreasing the rate of linear electron transport through PSII, in a process known as photosynthetic control (Foyer et al., 2012).
Light response curves collected over a short period of time may also be inadequate to describe photochemical responses for an entire growing period because photosynthetic rates can vary with leaf or plant age (Locke and Ort, 2014; Salmon et al., 2011) and can be affected by slow acclimation to light intensities. Acclimation to light intensities over the course of hours or days can lead to changes in the overall light response through mechanisms such as chlorophyll antennae rearrangement or changes in cellular metabolism and translation, and nuclear transcription, induced by chloroplast signaling (Dietz, 2015; Ruban, 2015). If factors such as ontogeny or acclimation impact the overall photochemical light response significantly, light response curves collected over only a few minutes may not describe realistic photochemical responses for a crop sufficiently, and longer term monitoring would be needed to characterize the photochemical response during a production cycle. Diurnal chlorophyll fluorescence monitoring can be used to gain a more detailed understanding of the photochemical light response under greenhouse lighting conditions (Weaver and van Iersel, 2016). This method consists of measuring chlorophyll fluorescence and PPFD over the course of several days, with measurements taken at regular intervals. In general, a 15-min interval between chlorophyll fluorescence measurements is sufficiently long to avoid measurement-induced photoinhibition resulting from the repeated application of saturating light pulses (van Iersel et al., 2016).
Although supplemental lighting can improve the growth, quality, and profitability of greenhouse-grown crops, the electricity requirement of supplemental lights can account for as much as 30% of the operating cost of a greenhouse (van Iersel and Gianino, 2017; Watson et al., 2018). The advent of light-emitting diode (LED) technology for horticultural lighting has facilitated the development of innovative approaches to providing and controlling greenhouse supplemental lighting (Morrow, 2008; Pinho et al., 2012; Singh et al., 2015). LED fixtures have several distinct advantages over the high-intensity discharge (HID) lamps traditionally used for greenhouse lighting, including their relatively high efficacy, low radiant heat load, and variable spectra. Another unique feature of LEDs is that the intensity of their light output can be controlled precisely and rapidly in a manner that is not possible with HID lamps. Lighting control systems that use this dimmability have the potential to reduce the electricity costs associated with providing supplemental light, and to increase the efficiency with which supplemental light is used for promoting plant growth. These adaptive, or dynamic, supplemental LED lighting control systems operate by keeping the LED lights off when ambient PPFD exceeds a predefined threshold PPFD. When ambient PPFD falls below this level, supplemental light is provided so that the combined PPFD of the LED lights and sunlight reaches, but does not exceed, the threshold. This ensures that supplemental light is provided only when the overall PPFD is relatively low, and the supplemental light can be used more efficiently by plants, because plant light use efficiency invariably decreases at greater PPFDs (Pinho et al., 2013; van Iersel and Gianino, 2017).
Providing supplemental light in a manner that allows it to be used most efficiently by a crop has the potential to decrease the amount of supplemental light, and thus the total amount of electricity required, for crop growth. For example, using simulations based on historical weather data and cultivar-specific light responses, Weaver and van Iersel (2018) estimated that the amount of supplemental light required for early season production can be reduced by 24% for Petunia ×hybrida ‘Daddy Blue’ and 37% for Impatiens walleriana ‘Super Elfin XP Violet’ using an adaptive lighting control approach that accounts for crop light use efficiency. Thus, understanding species- or cultivar-specific photosynthetic or photochemical responses to PPFD can facilitate the implementation of lighting control strategies that use the dimmability of LEDs fully and reduce electricity costs by providing supplemental light according to a specific crop’s ability to use that light efficiently.
Lettuce is an important greenhouse crop because there is a continuous demand for a supply of fresh leafy greens, production cycles are relatively short, and lettuce can be produced year-round in greenhouses if appropriate environmental conditions (e.g., light, temperature) are provided. Supplemental lighting for hydroponic greenhouse lettuce production has been the subject of a great deal of research, and some of the most advanced supplemental lighting strategies developed to date have focused on lettuce production (Albright et al., 2000; Bumgarner and Buck, 2016; Seginer et al., 2006). In our study, in situ diurnal chlorophyll fluorescence monitoring was used to evaluate the photochemical performance of a greenhouse-grown crop of a romaine-type lettuce cultivar (Lactuca sativa L. ‘Green Towers’) under growing conditions comparable to a commercial production environment. Specific hypotheses tested were whether the current ETR is affected by previous PPFDs during a day, and whether photochemical efficiency is affected by plant age or previous day’s DLI. In addition to quantifying instantaneous photochemical responses to PPFD, the integral of ETR over individual measurement days was calculated and defined as the DPI (mol·m−2·d−1), the integral of ETR over a 24-h period. Last, we conducted simulations to demonstrate how these data can be used to develop energy-efficient supplemental lighting strategies, and outline general methods for using adaptive lighting control to improve crop light use efficiency by decreasing the DLI required to achieve a given DPI, or increasing the resulting DPI for a fixed DLI.
Aaslyng, J.M., Lund, J.B., Ehler, N. & Rosenqvist, E. 2003 IntelliGrow: A greenhouse component-based climate control system Environ. Model. Softw. 18 657 666
Aikman, D.P. 1989 Potential increase in photosynthetic efficiency from the redistribution of solar radiation in a crop J. Expt. Bot. 40 855 864
Baker, N.R. & Rosenqvist, E. 2004 Applications of chlorophyll fluorescence can improve crop production strategies: An examination of future possibilities J. Expt. Bot. 55 1607 1621
Both, A.J., Albright, L.D., Langhans, R.W., Reiser, R.A. & Vinzant, B.G. 1997 Hydroponic lettuce production influenced by integrated supplemental light levels in a controlled environment agriculture facility: Experimental results Acta Hort. 418 45 52
Björkman, O. & Demmig, B. 1987 Photon yield of O2 evolution and chlorophyll fluorescence at 77k among vascular plants of diverse origins Planta 170 489 504
Bumgarner, N. & Buck, J. 2016 Light emitting diode and metal halide supplemental lighting for greenhouse Bibb lettuce production in the Midwestern United States J. Appl. Hort. 18 128 134
Clausen, A., Maersk-Moeller, H.M., Soerensen, J.C., Joergensen, B.N., Kjaer, K.H. & Ottosen, C.O. 2015 Integrating commercial greenhouses in the smart grid with demand response based control of supplemental lighting. Intl. Conf. Ind. Technol. Mgt. Sci. (ITMS 2015) 199–213.
Demmig-Adams, B., Cohu, C.M., Muller, O. & Adams, W.W. 2012 Modulation of photosynthetic energy conversion in nature: From seconds to seasons Photosynth. Res. 113 75 78
Dietz, K.J. 2015 Efficient high light acclimation involves rapid processes at multiple mechanistic levels J. Expt. Bot. 66 2401 2414
Foyer, C.H., Neukermans, J., Queval, G., Noctor, G. & Harbinson, J. 2012 Photosynthetic control of electron transport and the regulation of gene expression J. Expt. Bot. 63 1637 1661
Genty, B., Briantais, J. & Baker, N.R. 1989 The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence Biochim. Biophys. Acta 990 87 92
Horton, P. 2012 Optimization of light harvesting and photoprotection: Molecular mechanisms and physiological consequences Philos. Trans. R. Soc. Lond. B Biol. Sci. 367 3455 3465
Kaiser, E., Morales, A. & Harbinson, J. 2018 Fluctuating light takes crop photosynthesis on a rollercoaster ride Plant Physiol. 176 977 989
Kjaer, K.H., Ottosen, C.O. & Jørgensen, B.N. 2011 Cost-efficient light control for production of two Campanula species Scientia Hort. 129 825 831
Kjaer, K.H., Ottosen, C.O. & Jørgensen, B.N. 2012 Timing growth and development of Campanula by daily light integral and supplemental light level in a cost-efficient light control system Scientia Hort. 143 189 196
Locke, A.M. & Ort, D.R. 2014 Leaf hydraulic conductance declines in coordination with photosynthesis, transpiration and leaf water status as soybean leaves age regardless of soil moisture J. Expt. Bot. 65 6617 6627
Mauromicale, G., Ierna, A. & Marchese, M. 2006 Chlorophyll fluorescence and chlorophyll content in field-grown potato as affected by nitrogen supply, genotype, and plant age Photosynthetica 44 76 82
Murata, N. & Nishiyama, Y. 2018 ATP is a driving force in the repair of photosystem II during photoinhibition Plant Cell Environ. 41 285 299
Murchie, E.H. & Lawson, T. 2013 Chlorophyll fluorescence analysis: A guide to good practice and understanding some new applications J. Expt. Bot. 64 3983 3998
Pinho, P., Hytönen, T., Rantanen, M., Elomaa, P. & Halonen, L. 2013 Dynamic control of supplemental lighting intensity in a greenhouse environment Light. Res. Technol. 45 295 304
Rascher, U., Liebig, M. & Lüttge, U. 2000 Evaluation of instant light-response curves of chlorophyll fluorescence parameters obtained with a portable chlorophyll fluorometer on site in the field Plant Cell Environ. 23 1397 1405
Salmon, Y., Barnard, R.L. & Buchmann, N. 2011 Ontogeny and leaf gas exchange mediate the carbon isotopic signature of herbaceous plants Plant Cell Environ. 34 465 479
Seginer, I., Albright, L.D. & Ioslovich, I. 2006 Improved strategy for a constant daily light integral in greenhouses Biosyst. Eng. 93 69 80
Šesták, Z. 1999 Chlorophyll fluorescence kinetic depends on age of leaves and plants, p. 291–296. In: J.H. Argyroudi-Akoyunoglou and H. Senger (eds.). The chloroplast: From molecular biology to biotechnology. Springer, Dordrecht, Netherlands
Singh, D., Basu, C., Meinhardt-Wollweber, M. & Roth, B. 2015 LEDs for energy efficient greenhouse lighting Renew. Sustain. Energy Rev. 49 139 147
Slattery, R.A., Walker, B.J., Weber, A.P. & Ort, D.R. 2018 The impacts of fluctuating light on crop performance Plant Physiol. 176 990 1003
Spetea, C., Hundal, T., Lundin, B., Heddad, M., Adamska, I. & Andersson, B. 2004 Multiple evidence for nucleotide metabolism in the chloroplast thylakoid lumen Proc. Natl. Acad. Sci. USA 101 1409 1414
Torres, A.P. & Lopez, R.G. n.d Measuring daily light integral in a greenhouse. Dept. of Hort. and Landscape Architecture, Purdue Univ. Purdue Extension Bul. HO-238-W
van Iersel, M.W. 2017 Optimizing LED lighting in controlled environment agriculture, p. 59–80. In: S.D. Gupta (ed.). Light emitting diodes for agriculture: Smart lighting. Springer, Singapore
van Iersel, M.W. & Gianino, D. 2017 An adaptive control approach for light-emitting diode lights can reduce the energy costs of supplemental lighting in greenhouses HortScience 52 72 77
van Iersel, M.W., Weaver, G., Martin, M.T., Ferrarezi, R.S., Mattos, E. & . Haidekker, M 2016 A chlorophyll fluorescence-based biofeedback system to control photosynthetic lighting in controlled environment agriculture J. Amer. Soc. Hort. Sci. 141 169 176
Watson, R.T., Boudreau, M. & van Iersel, M.W. 2018 Simulation of greenhouse energy use: An application of energy informatics Energy Informatics 1 1
Weaver, G. & van Iersel, M.W. 2016 Screening photosynthetic performance of bedding plants using chlorophyll fluorescence Proc. SNA Res. Conf. 61 71 75
Weaver, G.M. & van Iersel, M.W. 2018 Modeling energy-efficient lighting strategies for petunia and impatiens using electron transport rate and historical weather data Proc. SNA Res. Conf. 62 29 34