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Photosynthetic lighting is one of the main costs of running controlled environment agriculture facilities. To optimize photosynthetic lighting, it is important to understand how plants use the provided light. When photosynthetic pigments absorb photons, the energy from those photons is used to drive the light reactions of photosynthesis, thermally dissipated, or re-emitted by chlorophyll as fluorescence. Chlorophyll fluorescence measurements can be used to determine the quantum yield of photosystem II (FPSII) and nonphotochemical quenching (NPQ), which is indicative of the amount of absorbed light energy that is dissipated as heat. Our objective was to develop and test a biofeedback system that allows for the control of photosynthetic photon flux density (PPFD) based on the physiological performance of the plants. To do so, we used a chlorophyll fluorometer to measure FPSII, and used these data and PPFD to calculate the electron transport rate (ETR) through PSII. A datalogger then adjusted the duty cycle of the light-emitting diodes (LEDs) based on the ratio of the measured ETR to a predefined target ETR (ETRT). The biofeedback system was able to maintain ETRs of 70 or 100 mmol m s over 16-hour periods in experiments conducted with lettuce (Lactuca sativa). With an ETRT of 70 mmol m s,FPSII was stable throughout the 16 hour and no appreciable changes in PPFD were needed. At an ETRT of 100 mmol m s, FPSII gradually decreased from 0.612 to 0.582. To maintain ETR at 100 mmol m s, PPFD had to be increased from 389 to 409 mmol m s, resulting in a gradual decrease ofFPSII and an increase in NPQ. The ability of the biofeedback system to achieve a range of different ETRswithin a single day was tested using lettuce, sweetpotato (Ipomoea batatas), and pothos (Epipremnum aureum). As the ETRTwas gradually increased, the PPFD required to achieve that ETR also increased, whereas FPSII decreased. Surprisingly, a subsequent decrease in ETRT, and in the PPFD required to achieve that ETR, resulted in only a small increase in FPSII. This indicates that FPSII was reduced because of photoinhibition. Our results show that the biofeedback system is able to maintain a wide range of ETRs, while it also is capable of distinguishing between NPQ and photoinhibition as causes for decreases in FPSII. Controlled environment agriculture, including greenhouses and indoor production facilities, is becoming an increasingly important part of the global food system. Totally enclosed, indoor vegetable growing facilities were developed in Japan beginning in the 1970s. These ‘‘plant factories’’ and similar operations can now be found in or near major cities across the world (Despommier, 2013; Goto, 2012; Liu, 2012; Mok et al., 2014; Thomaier et al., 2015). Recently, there has been much interest in the potential of large-scale, indoor agricultural production, often referred to as vertical farming. Vertical farming has been heralded as a potential solution to feeding a rapidly growing population while minimizing the environmental impacts of crop production (Despommier, 2010). It is estimated that vertical farms could produce 200 to 1000 ·more food per unit land area than traditional agriculture (Germer et al., 2011; Zeidler et al., 2013). However, building and operating vertical farms is expensive and edible biomass produced in a vertical farmwould need to be sold for $13.75/kg to cover the cost of operations (Zeidler et al., 2013). The feasibility and sustainability of large-scale plant factories are questionable. Plant factories are expensive to operate, partly because of the large power requirements of electric lamps that provide the actinic light that drives the light reactions of photosynthesis. In enclosed plant factories, photosynthesis is driven exclusively by electric light, and sunlight can at best provide a small percentage of the light required by vertical farms (Germer et al., 2011; Goto, 2012; Mok et al., 2014; Watanabe, 2011). Light-emitting diodes (LEDs) have become a popular Received for publication 19 Nov. 2015. Accepted for publication 31 Dec. 2015. Funding for this research was provided by the Georgia Research Alliance, USDA-NIFA-SBIR award 2015-33610-23472, and the Fred C. Gloeckner Foundation. Corresponding author. E-mail: mvanier@uga.edu. J. AMER. SOC. HORT. SCI. 141(2):169–176. 2016. 169 light source for indoor vegetable cultivation because they are considered to be energy efficient (Bourget, 2008; Goto, 2012; Liu, 2012; Morrow, 2008; Yeh and Chung, 2009), although the actual efficiency is debated. Nelson and Bugbee (2014) found that the efficiency of LED lights is similar to that of good highpressure sodium lights. In contrast, Ouzounis et al. (2015) state that good LED lights are more efficient than high-pressure sodium lights. Ouzounis et al. (2015) also conclude that the payback time for LEDs is now realistic, although this depends on electricity pricing. Furthermore, they point out that LEDs have unique properties that can make them more cost-effective, such as the ability to control light intensity and spectra. In a conceptual cost analysis of building and operating a large vertical farm, it was shown that the greatest recurring cost would be electricity. Powering and cooling LED lights would account for more than 30% of the operation’s total electrical cost (Zeidler et al., 2013). In plant factories, lighting accounts for 40% of the total recurring costs (Watanabe, 2011). The overall efficiency of converting electric light energy into biomass is complicated by the nature of the light reactions of photosynthesis. Not all photons absorbed by chlorophyll and accessory pigments are used to drive the light reactions. Excess absorbed photons can cause damage within the chloroplast. The reaction center of PSII, particularly the D1 protein, is highly susceptible to light-induced damage (photoinhibition), which decreases quantum yield of PSII. Higher plants have developed a complex set of responses to excess light, which allows them to safely dissipate excess light energy as heat (resulting in NPQ) and minimize photoinhibition (Demmig-Adams et al., 2012; Dietz, 2015; Horton, 2012; Rochaix, 2014). Although most of the light energy absorbed in chloroplasts is either used to drive photosynthesis or is thermally dissipated, a small fraction is re-emitted by chlorophyll as fluorescence. Chlorophyll fluorescence measurements can be used to quickly and reliably determine FPSII (Maxwell and Johnson, 2000). Generally, a decrease in FPSII indicates that either plants dissipate an increasing fraction of the absorbed light energy as heat (an upregulation of NPQ) or photoinhibition has occurred. Chlorophyll fluorescence measurements can be used to quantify NPQ, based on changes in light-saturated fluorescence under actinic light. Combined with PPFD, FPSII can be used to estimate electron transport rate through PSII (Baker and Rosenqvist, 2004; Genty et al., 1989). Because chlorophyll fluorescence is relatively easy to measure and provides detailed physiological information, such measurements can be a valuable tool to optimize greenhouse production (Baker and Rosenqvist, 2004) and to monitor crop responses to light (Pocock, 2015). For example, Janka et al. (2015) used chlorophyll fluorescence measurements to monitor diurnal and dynamic fluctuations in FPSII of greenhouse crops. Our objective was to develop a biofeedback system that not only monitors FPSII and ETR, but that can also control the PPFD fromLED lights tomaintain a specific ETR. Controlling the intensity of actinic light based on plants’ ability to use it efficiently may substantially reduce the energy cost of LED lighting, and contribute to making large-scale controlled environment agriculture more profitable. Materials and Methods LEDS AND CONTROL SYSTEM. Photosynthetic light was provided using a custom-built 400-W LED array consisting of four 100-W warm-white LED modules (3000 K; EpiLEDs, Tainan, Taiwan) mounted on aluminum heat sinks with air circulation provided by two 120-mm cooling fans. Glass lenses (Satistronix Group, Shenzhen, China) were attached below the LED modules to focus the light on the plant production area below the LEDs. The lights were powered using a 30-V DC power supply (Fig. 1). The LED array lighted an area of 0.75 m. A custom-made control board was used to alter the duty cycle of the LEDs (Fig. 1). The LEDs were continuously turned on and off (at 1000 Hz in this study; i.e., each on/off cycle lasted 1 ms) and duty cycle is the proportion of time that the diodes are energized during each cycle. Thus, the intensity of the LED light was modified by changing the duty cycle. The control board altered the duty cycle of the LEDs proportionally in response to an external 0to 2-V DC signal supplied to the board. To provide this voltage signal, an analog output module (SDM-A04; Campbell Scientific, Logan, UT) was connected to a datalogger (CR1000; Campbell Scientific). CHLOROPHYLL FLUORESCENCE MEASUREMENTS. Chlorophyll fluorescence was measured with a pulse-amplitude modulated fluorometer (MINI-PAM; Heinz Walz, Effeltrich, Germany). The fluorometer provided an analog voltage signal to the datalogger, proportional to the measured fluorescence. Following measurements of steady-state fluorescence (F0 and Fs in the dark and light, respectively), the datalogger sent a serial signal to the fluorometer to trigger a saturating light pulse and then Fig. 1. Diagram of the biofeedback system (top) and a simplified diagram of the duty cycle control board (bottom). A fluorometer is used to measure the quantum yield of photosystem II and the datalogger calculates the electron transport rate from this and the photosynthetic photon flux density (PPFD), measured using a quantum sensor. Based on the measured and target electron transport rate, the datalogger then uses an analog output module to send a voltage signal to the duty cycle control board. The duty cycle control board has a

A Chlorophyll Fluorescence-based Biofeedback System to Control Photosynthetic Lighting in Controlled Environment Agriculture