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Article

Impacts of Light Exposure and Soil Covering on Sweet Potato Storage Roots in a Novel Soilless Culture System

by
Masaru Sakamoto
* and
Takahiro Suzuki
Faculty of Biology-Oriented Science and Technology, Kindai University, Wakayama 649-6493, Japan
*
Author to whom correspondence should be addressed.
AgriEngineering 2024, 6(4), 3912-3930; https://doi.org/10.3390/agriengineering6040222
Submission received: 13 September 2024 / Revised: 19 October 2024 / Accepted: 23 October 2024 / Published: 24 October 2024
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Abstract

:
Soilless culture systems, which promote plant growth and enable the precise control of the root-zone environment, have yet to be fully established for sweet potatoes. In this study, we developed a soilless culture system and examined the effects of soil covering and light exposure on the storage roots of sweet potatoes. Sweet potato seedlings with induced storage roots were transplanted into five systems: a previously developed pot-based hydroponics system (Pot), an improved version with storage roots enclosed in a plastic box and covered with a soil sheet (SS), the SS system without the soil sheet (SD), the SD system with light exposure to storage roots after 54 days (SL), and a deep flow technique (DFT) hydroponics system. Our study enabled the time-course observation of storage root enlargement in the SS, SD, and SL systems. In the SL system, light exposure suppressed the storage root enlargement and reduced epidermal redness. No storage root enlargement was observed in the DFT system, even at 151 days after transplantation. Light exposure in the SL system increased the chlorophyll and total phenolic contents in the cortex beneath the epidermis, while the starch content was the lowest in this system. These findings indicate that the developed system can induce normal storage root enlargement without soil. Additionally, the observed changes in growth and composition due to light exposure suggest that this system is effective for controlling the root-zone environment of sweet potatoes.

1. Introduction

Light is an essential factor influencing various developmental processes in plants, including germination, flowering, and metabolism [1,2,3]. In fruits, light exposure has been shown to induce significant compositional changes [2,4,5,6]. For example, UV-B irradiation promotes anthocyanin biosynthesis in apple skins, while red light enhances red color development and increases carotenoid concentrations in post-harvest tomatoes [7,8]. In sweet potato leaves, UV radiation is known to induce peroxidase activity, thereby activating defense mechanisms against oxidative stress [9]. Beyond its role in photosynthetic tissues, light also influences root development. Photoreceptors such as phytochromes mediate light responses in roots [10]. In Arabidopsis, the exposure of roots to light inhibits their elongation through the action of HY5, a key regulator of photomorphogenesis [11]. Additionally, exposure to specific wavelengths, such as 940 nm, has been shown to increase root dry weight in Brassica napus [12]. These findings suggest that under-ground organs, which are typically not exposed to light, may exhibit unique physiological responses when exposed to light.
The physical properties of soil, where plant roots are established, also play a significant role in plant growth. Variations in soil type have been reported to affect the growth of root crops such as sweet potatoes and potatoes [13,14]. High soil pressure can inhibit root penetration, as observed in radishes [15]. Similarly, root elongation into compacted subsoil layers has been restricted in crops such as potatoes, carrots, and sweet potatoes [16,17]. Conversely, low-pressure conditions have been found to suppress the enlargement of sweet potato storage roots [18]. These studies collectively demonstrate that soil compaction and physical pressure are key factors that influence the development and enlargement of root systems in root crops.
Soilless culture systems have been successfully applied to a variety of leafy vegetables, enabling the precise management of plant growth [19]. These systems enable precise control over the root-zone environment, including regulating temperature, the nutrient solution, and chemical treatments, thereby facilitating extensive research on controlling plant growth and composition through these treatments [20,21,22]. However, studies focused on controlling the light environment of root zones remain limited to a few specific examples [12]. Meanwhile, the application of soilless culture systems to root crops such as sweet potatoes (Ipomoea batatas) remains underexplored, as root enlargement is inhibited when submerged in water [23,24,25]. Recently, some cultivation methods that prevent the submersion of enlarging roots in nutrient solutions have been developed. For instance, aeroponic systems have been utilized for potatoes, cassava, and burdock [26,27,28]. In sweet potato cultivation, methods have been developed to expose storage roots to air by creating space above the nutrient solution, allowing for root enlargement [29]. This approach, however, requires a cultivation apparatus with a specific height due to the direct contact between the storage roots and the nutrient solution tank. At the research level, systems have also been designed to separate storage roots from the nutrient solution, allowing for periodic observations [23]. Despite these advancements, these systems are primarily limited to early-stage growth and indoor settings, and they have not yet been established as robust cultivation methods.
Previously, we developed a cultivation method for sweet potatoes that uses small pots to reduce the amount of substrate required [30]. In this system, the storage roots grow in vermiculite, making it difficult to observe root enlargement and control the root-zone environment during growth. To address this, we improved the system by creating a separate space for root enlargement, independent of the nutrient solution tank. Unlike previously developed soilless culture systems, this new setup places the sweet potato storage roots horizontally on a nonwoven fabric above the tank, allowing for the real-time observation of root growth from the aerial part. In this study, we compared this system with pot-based cultivation and investigated the effects of light exposure and soil covering on the growth and composition of the storage roots.

2. Materials and Methods

2.1. Experimental Conditions

The sweet potato (Ipomoea batatas) cultivar “Narutokintoki” was used in this study. Stem cuttings were immersed in water for 10 days to promote the formation of adventitious roots. After 10 days, the cuttings with adventitious roots were transplanted into plastic pots filled with vermiculite and cultivated for 32 days to induce the formation of storage roots. On 3 June 2023, these seedlings with storage roots were transplanted into five different cultivation systems to begin the experimental cultivation (Figure 1).
The first method (Pot) utilized a previously developed pot-based nutrient solution cultivation system [30]. In this system, the storage roots were grown within pots filled with vermiculite, while fibrous roots extended along nonwoven fabric from the bottom of the pots to efficiently absorb the nutrient solution from the tank. The second method (SS; soil sheet) was an improved version of the first system, where the pots were removed, and the storage roots were covered with a plastic box (50 cm × 37 cm × 11.4 cm). Inside this box, the storage roots grew on nonwoven fabric in 100% relative humidity. In this setup, the upper part of the storage roots was covered with soil sheets to simulate a soil-like environment. The soil sheet consisted of 400 mL of vermiculite wrapped in a net, with one soil sheet used per plant for cultivation. These sheets could be removed for observation. The third method (SD; soilless dark) used the same improved system, but the storage roots were exposed to air and grown in the dark on nonwoven fabric. In the fourth method (SL; soilless light), the improved system (SD) was used to grow storage roots in the dark for 54 days. After this period, the storage roots were exposed to light by replacing the top of the plastic box with a translucent white board. The fifth method employed the deep flow technique (DFT), where the storage roots were submerged in water throughout the cultivation period.
In each system, culture containers (59 cm × 39 cm × 18 cm) were filled with a half-strength nutrient solution based on the OAT House recipe A treatment [31,32]. The nutrient solution was prepared by combining OAT House 1 and OAT House 2 (OAT Agrio Co., Ltd., Tokyo, Japan) at a 3:2 ratio. The resulting mixed solution contained the following concentrations: N, 260 mg L−1; P2O5, 120 mg L−1; K2O, 405 mg L−1; CaO, 230 mg L−1; MgO, 60 mg L−1; MnO, 1.5 mg L−1; B2O3, 1.5 mg L−1; Fe, 2.7 mg L−1; Cu, 0.03 mg L−1; Zn, 0.09 mg L−1; and Mo, 0.03 mg L−1. Water was added to compensate for reductions in the nutrient solution due to plant uptake or evaporation. The nutrient solution was replaced every two weeks. Plastic boards were covered with insulation sheets to maximize the utilization of sunlight for photosynthesis by reflection. In the SL system, exposure to light was achieved by replacing the top of the plastic box with a translucent white board. Each box had a 3 cm diameter hole on the side through which the plant stems were passed into the box. The gap between the hole and the stem was sealed with black urethane to prevent light from entering the inside of the box. Four plants were transplanted per container, with a total of 12 plants used for each system. In the SS, SD, and SL systems, the number of storage roots was limited to two per plant 17 days after transplantation (DAT), with all the other storage roots removed.
The temperatures in the storage root zone in four of the systems (excluding the DFT system) and outside the apparatus were investigated on cloudy and sunny days (Figure 2A). During the day, temperatures were lower in the SS and SD systems, while at night, temperatures were lower in the Pot system and outside the apparatus. Additionally, the temperature difference during the day was more pronounced on sunny days. The light irradiating the storage roots in the SD system was approximately one-tenth of direct sunlight, containing almost no UV light and having a higher proportion of long-wavelength light (Figure 2B,C). The plants were cultivated for 151 days (from 3 June to 1 November 2023) in the open experimental field of Kindai University (Faculty of Biology-Oriented Science Technology, Wakayama, Japan). According to the Japan Meteorological Agency [33], the average temperatures in Wakayama City in 2023 were 19.7 °C in May, 23.3 °C in June, 28.3 °C in July, 29.1 °C in August, 27.2 °C in September, and 19.2 °C in October. The average relative humidity during these months was 69%, 78%, 75%, 76%, 74%, and 66%, respectively. The experimental field is located approximately 17 km from the meteorological station in Wakayama City and is 90 m higher in elevation.

2.2. Measurement of Plant Growth and Yield

Time-course observations of the storage roots were conducted in the SS, SD, and SL systems. Two storage roots were measured per plant. The observations were made in the evening when direct sunlight no longer hit the cultivation apparatus. After removing the plastic boxes from each system, photographs were taken from the same position each time to investigate changes in the size and morphology of the storage roots. Consequently, during the photography and measurements, the storage roots in the SS and SD systems were also temporarily exposed to low light. The photographs were processed using ImageJ to correct for positional shifts. The diameter of the storage roots was measured in millimeters (mm) using a digital caliper gauge (EDC-A1150, Niigata Seiki Co., Ltd., Niigata, Japan). The measurements with calipers began at 42 DAT; prior to this, the root diameters were calculated from the captured photographs using ImageJ. Color difference measurements began at 52 DAT, and in the SL system, only the top-side epidermis of the storage roots directly exposed to light was measured.
The plants were harvested at 151 DAT. In the three systems (SS, SD, and SL) where the number of storage roots was limited to two 17 DAT, most plants formed only two storage roots, but some individuals formed storage roots under the nonwoven fabric. These storage roots were included in the total storage root weight per plant, measured in grams (g). The above-ground parts and storage roots of the plants were first weighed fresh. Half of these samples were then reweighed before and after being dried in an oven at 80 °C for over a week to determine their dry matter content. The moisture content was calculated by comparing the initial fresh weight with the dry weight. The diameter measurements and samples for observation were taken from the basal side of the stem of all five systems.

2.3. Measurement of the Color of Storage Roots

The color of the storage root epidermis was evaluated using a WR-10 colorimeter (FRU, China). The reflectance values for the L* (luminosity: black–white axis), a* (green–red axis), and b* (blue–yellow axis) coordinates were directly recorded at a single central point on each storage root. The color difference in the storage roots at 151 DAT was measured after gently washing the surface of the harvested storage roots with water to remove any surface dirt. In the SL system, the epidermis of the storage roots was measured separately for the top side exposed to direct light (SL-T) and the bottom side not exposed to light (SL-B).

2.4. Measurement of Anthocyanin Content

The anthocyanin content was spectrophotometrically measured as previously described in [34]. A 1 cm2 section of the storage root epidermis was peeled and immersed in 90% methanol containing 1% hydrochloric acid. After storing the samples at 4 °C for one week, the absorbance of the solution was measured at 533 nm.

2.5. Measurement of Total Phenolic Contents

The total phenolic content was measured using the Folin–Ciocâlteu method as previously described in [35]. The epidermis of the storage root (50 mg) was homogenized in 500 μL of 90% methanol and centrifuged at 10,000× g for 5 min. The supernatant (50 μL) was diluted to 630 μL with distilled water and mixed with 50 μL of phenol reagent. After adding 300 μL of 5% sodium carbonate, the mixture was incubated at 25 °C for 30 min. The absorbance of the supernatant was measured at 765 nm, and a calibration curve was created using gallic acid. The absorbance was converted to total phenolic concentration in milligrams of gallic acid per gram of fresh weight.

2.6. Measurement of Chlorophyll Content

A 1 cm2 section of the cortex, after peeling and removing the storage root epidermis, was sliced to a thickness of 1 mm and immersed in 80% methanol. After storing the samples at 4 °C for one week, the absorbance of the solution was measured at 647 nm and 664 nm. The total chlorophyll content was calculated based on the formula reported by Chazaux et al. [36].

2.7. Measurement of Starch Content

The total starch content was measured spectrophotometrically as previously described in [37]. The sweet potato storage roots were homogenized, and 10 mg of the sample was used to quantify the starch content using a starch colorimetric/fluorimetric assay kit (K647-100, BioVision, Mountain View, CA, USA). The starch content was determined by measuring the absorbance at 570 nm.

2.8. Starch Staining

The histological analysis of starch was conducted as previously described in [38]. The sweet potato stems were thinly sliced and washed three times with distilled water. The samples were then immersed in Lugol’s iodine solution for 10 min for staining. After staining, the samples were repeatedly washed with distilled water until they were completely decolorized. The stem samples were observed using a stereo microscope.

2.9. Lignin Staining

The histological analysis of lignin was conducted as previously described in [30]. The storage roots and stems were sectioned freehand using a razor blade. The sections were stained with 1% phloroglucinol HCl solution for 5 min and observed under a stereo microscope.

2.10. Data Analysis

Data analysis was conducted using the JMP software (version 8.0, SAS Institute, Cary, NC, USA). Significant differences among the systems were determined using a one-way analysis of variance (ANOVA), followed by the Tukey–Kramer honest significant difference (HSD) test for pairwise comparisons. Statistical significance was set at p < 0.05.

3. Results

3.1. Time-Course Observations of Storage Root Growth

The time-course observations of storage root development under the SS, SD, and SL systems showed enlargement across all the systems (Figure 3, Supplementary Videos S1–S3). In the SL system, a reduction in the redness of the storage root epidermis was observed at 58 DAT, four days after the start of light exposure, with noticeable greening by 96 DAT. Additionally, fibrous roots proliferated on the nonwoven fabric near the storage roots, particularly in the SS system. The measurements of the storage root diameter over time indicated that the SS system promoted early-stage enlargement compared to the other systems (Figure 4). After light exposure began, the SL system exhibited suppressed storage root enlargement compared to the SD system at 80 DAT. In all the systems, the storage root diameter increased at an accelerated rate during the later stages of growth, with this trend being most pronounced in the SD system.

3.2. Time-Course Observations of Storage Root Color

The storage root epidermis color was monitored over time using a colorimeter. At 52 DAT, before light exposure, the lightness (L* value) was higher in the SS system compared to the other systems (Figure 5A). Light exposure significantly reduced the L* value in the SL system compared to the SD system. As growth progressed, the L* value decreased across all the systems, with diminishing differences between them. The redness (a* value) at 52 DAT was also higher in the SS system (Figure 5B). Light exposure caused a significant decrease in the a* value in the SL system, which did not recover by the time of harvest. In contrast, the differences between the SS and SD systems nearly disappeared over time, with both showing a tendency for the a* value to increase. The yellowness (b* value) at 52 DAT was lower in the SS system (Figure 5C). Light exposure caused a significant increase in the b* value in the SL system, with differences between the systems persisting until harvest. At 58 DAT, four days after light exposure began, the color coordinates a* and b* showed a significant change in the SL system, with a decrease in the a* value and an increase in the b* value (Figure 5D,E). This change progressed until 96 DAT, with partial recovery towards the original color by 150 DAT, as indicated by an increase in the a* value and a decrease in the b* value (Figure 5F,G).

3.3. Storage Root Growth and Morphology at Harvest

At 151 DAT, storage roots from each system were harvested and their morphology was observed. In the Pot system, consistent with previous findings, the storage roots exhibited irregular shapes, with both enlarged and non-enlarged sections (Figure 6). In the three systems using the newly developed cultivation apparatus (SS, SD, and SL), normal-shaped enlarged roots were formed, with most systems resulting in only two enlarged roots due to the limitations imposed. In the SL system, the storage roots were smaller and less red compared to the SD system, where the roots were not exposed to light. Conversely, in the DFT system, only fibrous roots developed, and the storage roots present at the time of transplantation did not enlarge, with many red pencil roots observed. In the SL system, the storage root diameter and weight per root were significantly reduced compared to the other three systems (Figure 7A,B). The total storage root weight was highest in the Pot system (Figure 7C). There were no significant differences in the moisture content of the storage roots among the systems (Figure 7D).
Next, changes in the surface and internal characteristics of the storage roots were investigated after gently washing the surface with water to remove any adhered dirt. In the SL system, the storage roots were divided into the top side, exposed to direct light (SL-T), and the bottom side, not exposed to light (SL-B), for analysis. The L* value was highest in the Pot system, followed by the SS and SD systems. Within the SL system, the L* value was lower, with the SL-B condition higher than the SL-T condition (Figure 8A). The a* value followed a similar trend, being highest in the Pot system, followed by the SS and SD systems, with lower values in the SL-B condition and a markedly lower value in the SL-T condition (Figure 8B). In contrast, the b* value was highest in the SL-T condition, followed by the SL-B, SD, and SS systems, with the Pot system showing the lowest value (Figure 8C). These results indicate that within the SL system, the SL-T condition resulted in less redness (a* value) and more yellowness (b* value) compared to the SL-B condition, consistent with our visual observations (Figure 9). The cross-sectional observations of the storage roots revealed green pigmentation in the cortex beneath the epidermis in the SL-T condition, as confirmed by peeling the epidermis (Figure 9). This result is also consistent with the decrease in the a* value, which is the complementary color of green, under the SL-T condition. Additionally, the color parameters in the SS, SD, and SL-T conditions were nearly identical to the values observed at 150 days after plantation, just before harvest (Figure 5).
Chlorophyll quantification in the cortex beneath the epidermis showed significant chlorophyll production in the SL-T condition (Figure 10A). Plants synthesize antioxidant phenolic compounds in response to oxidative stress, so we measured the total phenolic and anthocyanin contents [39,40]. The total phenolic content in the cortex beneath the epidermis was the highest for the SL-T condition, followed by the SL-B, Pot, SS, and SD systems, in that order (Figure 10B). The anthocyanin content in the epidermis was highest in the Pot system, followed by the SS and SD systems, with the lowest levels in the SL-B and SL-T conditions (Figure 10C). The starch content in the storage roots was highest in the Pot system, followed by the SD and SS systems, with the lowest levels in the SL system (Figure 10D). Phloroglucinol staining for lignin accumulation in the storage roots showed no significant lignification, as observed in the pencil roots, with no differences among the systems (Figure 11).

3.4. Above-Ground Growth and Morphology at Harvest

The maximum stem length was measured as a parameter for the time-course growth of the above-ground parts. At 139 DAT, just before harvest, stem growth was suppressed in the DFT system compared to the other systems (Figure 12A). Conversely, at 151 DAT, the stem diameter was significantly larger in the DFT system (Figure 12B). The fresh weight of the above-ground parts was highest in the Pot system and lowest in the DFT system (Figure 12C). The moisture content of the above-ground parts was significantly lower in the SL system compared to the Pot, SS, and SD systems, and even lower in the DFT system (Figure 12D). Given the lack of storage root enlargement and the increase in the stem diameter observed in the DFT system, stem cross-sections were examined histologically (Figure 13). The results showed that lignin staining in the stems was suppressed in the DFT system compared to the other four systems. In the systems where strong lignin staining was observed, vessel development was noted in the lignified areas. In contrast, Lugol’s reagent revealed that starch accumulated throughout the stems in the DFT system. In the other four systems, starch accumulation was suppressed in the lignified areas.

4. Discussion

4.1. Comparison of Pot and Novel Soilless Culture System

This study examined the growth of sweet potatoes using a novel soilless culture system. Unlike the Pot system, the new methods (SS, SD, and SL) supported the normal formation of storage roots. In the Pot system, the restricted growth space led to lignification and inhibited the enlargement of some parts of the storage roots, as previously reported in [30]. Given that limited substrate volume is known to reduce storage root length [41], the roots in the new system, exposed to air, were free from physical constraints, allowing for the development of normally shaped storage roots.
Temporal observations revealed that this system promoted increases in the storage root diameter during both the early and late growth stages, aligning with previous findings on sweet potato storage root weight gain during these stages [42]. Therefore, this system holds the potential for facilitating the real-time monitoring of storage root enlargement and morphological changes. No new storage roots formed on the upper part of the nonwoven fabric; only fibrous roots proliferated. This suggests that while the system supports the normal enlargement of the existing storage roots, it does not induce the formation of new storage roots. The formation of sweet potato storage roots has been shown to be suppressed by low-moisture conditions [43]. In our study, the root zone’s relative humidity in the novel system was maintained at 100%. Therefore, it is likely that other environmental factors, such as the physical interactions between the soil and storage roots, rather than humidity alone, impeded the formation of storage roots.
The total sweet potato yield was lower in the new systems compared to the Pot system, likely due to the differences in rhizosphere substrate conditions [44,45]. In the new systems, only two enlarged roots were retained during the early growth stage, with the others removed. Reducing the sink size is known to limit the translocation of photosynthates to the sink in crops such as wheat, tomatoes, and carrots [46,47,48]. This reduction in sink size likely contributed to the lower total storage root weight observed in this experiment.

4.2. Effect of Soil Covering

The enlargement of sweet potato storage roots is influenced by various external factors, including nutrients, temperature, and humidity [43,49,50,51]. In our study, it was found that covering the storage roots with a soil sheet did not affect their enlargement compared to the uncovered roots. Previous experiments using the DFT system have demonstrated that sweet potato storage roots can enlarge in the air space above the nutrient solution [23,29], suggesting that physical stimulation from the soil is not required for storage root enlargement. Furthermore, covering the storage roots with a soil sheet promoted the growth of fibrous roots on the nonwoven fabric. In maize and barley, it has been reported that root exposure to air spaces in the soil inhibits lateral root formation [52]. This suggests that fibrous roots exposed to air may have restricted growth on the nonwoven fabric due to inhibited lateral root formation. Given that the initial enlargement of the storage roots was promoted by the soil sheet, it is likely that exposure to air in the SD system suppressed the establishment and proliferation of fibrous roots on the nonwoven fabric, leading to inhibited initial storage root enlargement. Additionally, the continuous increase in fibrous roots in the SS system may have slightly reduced the translocation of photosynthates to the storage roots, resulting in a lower growth rate of storage root diameter in the SS system compared to the SD system by the end of the cultivation period. In contrast, at 54 DAT, the presence of the soil sheet increased the L* and a* values while decreasing the b* value, leading to a redder color. This suggests that contact between soil and storage roots influences the color and, consequently, the composition of the storage root epidermis.

4.3. Effects of Light Exposure

Light exposure during the cultivation of root crops, such as carrots and sweet potatoes, has been reported to inhibit root enlargement [53,54]. Studies have shown that illuminated carrot root sections exhibit significantly less enlargement compared to roots grown in darkness—a process that is also associated with a reduction in carotenoid synthesis [53,55]. In sweet potatoes, exposing the upper part of the storage root to sunlight by removing the soil cover similarly hinders root enlargement [54]. However, this inhibition is reversible; when the exposed root is re-covered with soil, enlargement resumes. The permanent inhibition of root growth is associated with the lignification of the central stele [56,57]. In the SL system, no lignification was observed, and the storage root growth continued into the late stages of cultivation, indicating that the inhibition caused by light exposure was reversible and not linked to lignification.
In the SL system, light exposure reduced the moisture content of the above-ground parts at harvest. Various stresses, including heat, drought, and salinity, are known to reduce moisture content [58,59,60,61]. Additionally, Arabidopsis roots have been shown to produce reactive oxygen species in response to light-induced stress on the roots [62]. In our study, the increase in total phenolics, which act as antioxidants, in illuminated storage roots suggests that light exposure triggered oxidative stress, contributing to the decreased moisture content in the above-ground parts and inhibiting storage root growth.
Significant changes in root color were observed within two days of light exposure, with a rapid decrease in redness and an increase in yellowness in the SL system. In carrots, exposing the taproot to light increases chlorophyll and decreases carotenoid contents, altering the epidermal color from orange to green [53]. Chlorophyll accumulation occurs in the thickened stele of Arabidopsis roots exposed to light, but not in the outer cell layers [63]. In our study, chlorophyll was confirmed in the outer cortex of sweet potato storage roots, suggesting that the tissue’s response to light varies between plant species. Although anthocyanin synthesis is generally induced by light [64], our study found a decrease in anthocyanin content in the epidermis of light-exposed storage roots. Interestingly, in Medicago truncatula, sucrose treatment increases the anthocyanin content in the roots [65]. Considering the inhibition of storage root enlargement and the decrease in starch content due to exposure to light, it is suggested that the latter inhibits the translocation of photosynthetic products to the storage roots, resulting in reduced sucrose content and the suppression of anthocyanin synthesis. However, anthocyanin synthesis is influenced by various factors, including temperature and hormones, in addition to light [66,67]. The higher anthocyanin content in the epidermis of storage roots in the Pot system compared to the other systems suggests that environmental factors in the storage root zone also play a role in anthocyanin accumulation. Importantly, at each timepoint during root measurement, the storage roots in the SS and SD systems were temporarily exposed to light. This temporary exposure may have contributed to the reduced anthocyanin accumulation compared to the Pot system.

4.4. DFT Hydroponics

In sweet potatoes, submerging the roots in water is known to induce the lignification of the central stele in storage roots, thereby inhibiting their enlargement [23]. A similar inhibitory effect on root enlargement has been reported in carrots when the taproot is submerged [24]. In our experiment, the DFT system also completely inhibited the enlargement of the storage roots. However, an interesting observation in the DFT system was the increase in stem diameter compared to the other systems, accompanied by starch accumulation and suppression of lignification. The negative correlation between lignification and starch accumulation is well documented in sweet potatoes and cassava [68,69]. This suggests that in the DFT system, the photosynthetic products, which could not be directed to the inhibited storage roots, were instead diverted to the stems. As a result, the stems, which would typically lignify and harden, did not lignify, leading to their enlargement. Additionally, it has been reported that under cadmium stress, the common reed accumulates starch in its lower stem [70]. Considering the significant decrease in water content in the above-ground parts observed in the DFT system, it is plausible that stress, including the limitation of sinks due to the inhibition of storage root enlargement, induced starch accumulation in the stems.

5. Conclusions

To avoid submerging roots in water, various soilless cultivation methods, such as aeroponics, have been developed for root vegetables. Experimental cultivation using aeroponic systems has been researched for crops such as burdock, potato, and cassava; however, these systems require space for misting the root zone [26,28,71]. Similarly, DFT systems for sweet potatoes and potatoes require a designated space above the nutrient solution [18,23]. The soilless culture system developed in this study represents a substantial improvement over conventional methods by enabling the horizontal arrangement of storage roots, thereby enhancing spatial efficiency. While the current design includes vertical space for the temporary observation of the roots, further advancements in root-covering techniques are expected to optimize space utilization and boost cultivation efficiency. This space-saving feature makes our system particularly suitable for plant factories, where space is a limiting factor, and it holds significant potential for both large-scale and commercial farming applications.
In the experiment reported here, we utilized seedlings that had already been induced to form storage roots before transplantation into the system. It is believed that in sweet potatoes, the processes of storage root formation and the subsequent growth are distinct [72]. Thus, the induction of storage roots from sweet potato stem cuttings may require the development of a different induction method for storage root enlargement, which will need to be explored in future studies.
Recent efforts have focused on lunar farming as a part of expanding human activities beyond Earth [73]. Sweet potatoes have been identified as a viable crop for such endeavors by NASA, JAXA, and others [74,75,76,77]. Although lunar regolith, the moon’s soil, is known to impede the growth of plants such as Arabidopsis [78], importing soil from Earth is not feasible. The innovative, space-efficient, soilless cultivation system developed in this study offers a promising solution for growing root crops in extraterrestrial environments.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/agriengineering6040222/s1. Video S1: Time-course change in storage roots of sweet potatoes cultivated in the Soil Sheet (SS) system. The numbers represent the days post transplantation (dpt) to the system. Video S2: Time-course change in storage roots of sweet potatoes cultivated in the Soilless Dark (SD) system. The numbers represent the days post transplantation (dpt) to the system. Video S3. Time-course change in storage roots of sweet potatoes cultivated in the Soilless Light (SL) system. The numbers represent the days post transplantation (dpt) to the system.

Author Contributions

Conceptualization, M.S. and T.S.; formal analysis, M.S.; investigation, M.S.; writing—original draft preparation, M.S.; writing—review and editing, M.S. and T.S.; funding acquisition, M.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research was supported by JSPS Grant-in-Aid for Scientific Research C, grant number 23K05474.

Data Availability Statement

The original contributions presented in the study are included in the article, further inquiries can be directed to the corresponding author.

Conflicts of Interest

The authors declare no conflicts of interests.

References

  1. Fankhauser, C.; Chory, J. Light Control of Plant Development. Annu. Rev. Cell Dev. Biol. 1997, 13, 203–229. [Google Scholar] [CrossRef] [PubMed]
  2. Ntagkas, N.; de Vos, R.C.H.; Woltering, E.J.; Nicole, C.C.S.; Labrie, C.; Marcelis, L.F.M. Modulation of the Tomato Fruit Metabolome by LED Light. Metabolites 2020, 10, 266. [Google Scholar] [CrossRef] [PubMed]
  3. Liu, Y.; Singh, S.K.; Pattanaik, S.; Wang, H.; Yuan, L. Light Regulation of the Biosynthesis of Phenolics, Terpenoids, and Alkaloids in Plants. Commun. Biol. 2023, 6, 1055. [Google Scholar] [CrossRef]
  4. Zoratti, L.; Karppinen, K.; Luengo Escobar, A.; Häggman, H.; Jaakola, L. Light-Controlled Flavonoid Biosynthesis in Fruits. Front. Plant Sci. 2014, 5, 534. [Google Scholar] [CrossRef]
  5. Chen, S.; Wang, X.; Cheng, Y.; Gao, H.; Chen, X. Effects of Supplemental Lighting on Flavonoid and Anthocyanin Biosynthesis in Strawberry Flesh Revealed via Metabolome and Transcriptome Co-Analysis. Plants 2024, 13, 1070. [Google Scholar] [CrossRef]
  6. An, J.-P.; Liu, Y.-J.; Zhang, X.-W.; Bi, S.-Q.; Wang, X.-F.; You, C.-X.; Hao, Y.-J. Dynamic Regulation of Anthocyanin Biosynthesis at Different Light Intensities by the BT2-TCP46-MYB1 Module in Apple. J. Exp. Bot. 2020, 71, 3094–3109. [Google Scholar] [CrossRef]
  7. Ubi, B.E.; Honda, C.; Bessho, H.; Kondo, S.; Wada, M.; Kobayashi, S.; Moriguchi, T. Expression Analysis of Anthocyanin Biosynthetic Genes in Apple Skin: Effect of UV-B and Temperature. Plant Sci. 2006, 170, 571–578. [Google Scholar] [CrossRef]
  8. Panjai, L.; Röhlen-Schmittgen, S.; Ellenberger, J.; Noga, G.; Hunsche, M.; Fiebig, A. Effect of Postharvest Irradiation with Red Light on Epidermal Color and Carotenoid Concentration in Different Parts of Tomatoes. Food Meas. 2021, 15, 1737–1746. [Google Scholar] [CrossRef]
  9. Kim, Y.-H.; Lim, S.; Han, S.-H.; Lee, J.-C.; Song, W.-K.; Bang, J.-W.; Kwon, S.-Y.; Lee, H.-S.; Kwak, S.-S. Differential Expression of 10 Sweetpotato Peroxidases in Response to Sulfur Dioxide, Ozone, and Ultraviolet Radiation. Plant Physiol. Biochem. 2007, 45, 908–914. [Google Scholar] [CrossRef]
  10. Mo, M.; Yokawa, K.; Wan, Y.; Baluška, F. How and Why Do Root Apices Sense Light under the Soil Surface? Front. Plant Sci. 2015, 6, 775. [Google Scholar] [CrossRef]
  11. Zhang, Y.; Wang, C.; Xu, H.; Shi, X.; Zhen, W.; Hu, Z.; Huang, J.; Zheng, Y.; Huang, P.; Zhang, K.-X.; et al. HY5 Contributes to Light-Regulated Root System Architecture Under a Root-Covered Culture System. Front. Plant Sci. 2019, 10, 1490. [Google Scholar] [CrossRef] [PubMed]
  12. Kon, S.; Toyofuku, K.; Muto, D.; Kimura, S.; Ogawa, A. Irradiating Roots of Komatsuna (Brassica napus) with Various Light Qualities Affects Growth and Nutrient Content in Leaves, Stems, and Roots. Sci. Hortic. 2024, 331, 113179. [Google Scholar] [CrossRef]
  13. Agbede, T.M.; Oyewumi, A. Soil Properties, Sweet Potato Growth and Yield under Biochar, Poultry Manure and Their Combination in Two Degraded Alfisols of Humid Tropics. Sci. Hortic. 2022, 304, 111331. [Google Scholar] [CrossRef]
  14. Hajiaghaei Kamrani, M.; Rahimi Chegeni, A.; Hosseinniya, H. Effects of Different Growing Media on Yield and Growth Parameters of Potato Minitubers (Solanum tuberosum L.). Commun. Soil Sci. Plant Anal. 2019, 50, 1838–1853. [Google Scholar] [CrossRef]
  15. Tomobe, H.; Tsugawa, S.; Yoshida, Y.; Arita, T.; Tsai, A.Y.-L.; Kubo, M.; Demura, T.; Sawa, S. A Mechanical Theory of Competition between Plant Root Growth and Soil Pressure Reveals a Potential Mechanism of Root Penetration. Sci. Rep. 2023, 13, 7473. [Google Scholar] [CrossRef]
  16. Johansen, T.J.; Thomsen, M.G.; Løes, A.-K.; Riley, H. Root Development in Potato and Carrot Crops–Influences of Soil Compaction. Acta Agric. Scand. Sect. B Soil Plant Sci. 2015, 65, 182–192. [Google Scholar] [CrossRef]
  17. Pérez, M.; García, A.; Paredes, A.; Luna, J.; Madriz, P. Soil Mechanical Resistance to Root Penetration and Shape of the Reserve Root of Sweet Potato from the Huaman Descriptor. Agron. Costarric. 2016, 40, 147–159. [Google Scholar] [CrossRef]
  18. Kudo, K.; Sato, N.; Noborio, K. Tuberous Root Enlargement of Sweet Potato under Low Soil Presser Conditions. JASMAC Proc. 2023, 35, OR1-3. Available online: https://www.jasma.info/jasmac-35/wp-content/uploads/sites/15/2023/09/OR1-3-1.pdf (accessed on 12 September 2024).
  19. Kozai, T. Sustanable Plant Factory: Closed Plant Production System with Articicial Light for High Resource Use Efficiencies and Quality Produce. Acta Hortic. 2013, 1004, 27–40. [Google Scholar] [CrossRef]
  20. Sakamoto, M.; Suzuki, T. Effect of Root-Zone Temperature on Growth and Quality of Hydroponically Grown Red Leaf Lettuce (Lactuca sativa L. Cv. Red Wave). Am. J. Plant Sci. 2015, 6, 2350. [Google Scholar] [CrossRef]
  21. Thapa, U.; Nandi, S.; Rai, R.; Upadhyay, A. Effect of Nitrogen Levels and Harvest Timing on Growth, Yield and Quality of Lettuce under Floating Hydroponic System. J. Plant Nutr. 2022, 45, 2563–2577. [Google Scholar] [CrossRef]
  22. Lau, V.; Mattson, N. Effects of Hydrogen Peroxide on Organically Fertilized Hydroponic Lettuce (Lactuca sativa L.). Horticulturae 2021, 7, 106. [Google Scholar] [CrossRef]
  23. Eguchi, T.; Yoshida, S. A Cultivation Method to Ensure Tuberous Root Formation in Sweetpotatoes (Ipomoea batatas (L.) Lam.). Environ. Control. Biol. 2004, 42, 259–266. [Google Scholar] [CrossRef]
  24. Kusakawa, T.; Inoue, M. Damage to Pot-cultured Carrot Growth due to a Temporarily Raised Groundwater Level and Flooding Period. Hort. Res. 2010, 9, 495–500. [Google Scholar] [CrossRef]
  25. Siqinbatu; Kitaya, Y.; Hirai, H.; Shibuya, T.; Endo, R. Effects of Soil Water Content on the Growth of Sweet Potato Grown in Sandy Soil. Eco Eng. 2014, 26, 75–80. [Google Scholar] [CrossRef]
  26. Hayden, A. Aeroponic and Hydroponic Systems for Medicinal Herb, Rhizome, and Root Crops. HortScience 2006, 41, 536–538. [Google Scholar] [CrossRef]
  27. Tunio, M.H.; Gao, J.; Shaikh, S.A.; Lakhiar, I.A.; Qureshi, W.A.; Solangi, K.A.; Chandio, F.A. Potato Production in Aeroponics: An Emerging Food Growing System in Sustainable Agriculture for Food Security. Chil. J. Agric. Res. 2020, 80, 118–132. [Google Scholar] [CrossRef]
  28. Selvaraj, M.G.; Montoya-P, M.E.; Atanbori, J.; French, A.P.; Pridmore, T. A Low-Cost Aeroponic Phenotyping System for Storage Root Development: Unravelling the below-Ground Secrets of Cassava (Manihot esculenta). Plant Methods 2019, 15, 131. [Google Scholar] [CrossRef]
  29. Uewada, T. The solution culture of sweet potatoes. Environ. Control. Biol. 1990, 28, 135–140. [Google Scholar] [CrossRef]
  30. Sakamoto, M.; Suzuki, T. Effect of Pot Volume on the Growth of Sweetpotato Cultivated in the New Hydroponic System. Sustain. Agric. Res. 2018, 7, 137–145. [Google Scholar] [CrossRef]
  31. Sakamoto, M.; Komatsu, Y.; Suzuki, T. Nutrient Deficiency Affects the Growth and Nitrate Concentration of Hydroponic Radish. Horticulturae 2021, 7, 525. [Google Scholar] [CrossRef]
  32. Lee, J.-H.; Goto, E. Ozone Control as a Novel Method to Improve Health-Promoting Bioactive Compounds in Red Leaf Lettuce (Lactuca sativa L.). Front. Plant Sci. 2022, 13, 1045239. [Google Scholar] [CrossRef] [PubMed]
  33. Japan Meteorological Agency Official Website. Past Weather Data of Wakayama City in 2023. Available online: https://www.data.jma.go.jp/stats/etrn/view/annually_s.php?prec_no=65&block_no=47777&year=2023&month=&day=&view= (accessed on 12 September 2024).
  34. Sakamoto, M.; Suzuki, T. Synergistic Effects of a Night Temperature Shift and Methyl Jasmonate on the Production of Anthocyanin in Red Leaf Lettuce. Am. J. Plant Sci. 2017, 8, 1534. [Google Scholar] [CrossRef]
  35. Sakamoto, M.; Suzuki, T. Methyl Jasmonate and Salinity Increase Anthocyanin Accumulation in Radish Sprouts. Horticulturae 2019, 5, 62. [Google Scholar] [CrossRef]
  36. Chazaux, M.; Schiphorst, C.; Lazzari, G.; Caffarri, S. Precise Estimation of Chlorophyll a, b and Carotenoid Content by Deconvolution of the Absorption Spectrum and New Simultaneous Equations for Chl Determination. Plant J. 2022, 109, 1630–1648. [Google Scholar] [CrossRef]
  37. Korte, P.; Unzner, A.; Damm, T.; Berger, S.; Krischke, M.; Mueller, M.J. High Triacylglycerol Turnover Is Required for Efficient Opening of Stomata during Heat Stress in Arabidopsis. Plant J. 2023, 115, 81–96. [Google Scholar] [CrossRef]
  38. Wang, Y.; Yu, B.; Zhao, J.; Guo, J.; Li, Y.; Han, S.; Huang, L.; Du, Y.; Hong, Y.; Tang, D.; et al. Autophagy Contributes to Leaf Starch Degradation. Plant Cell 2013, 25, 1383–1399. [Google Scholar] [CrossRef]
  39. Valifard, M.; Mohsenzadeh, S.; Kholdebarin, B.; Rowshan, V. Effects of Salt Stress on Volatile Compounds, Total Phenolic Content and Antioxidant Activities of Salvia mirzayanii. S. Afr. J. Bot. 2014, 93, 92–97. [Google Scholar] [CrossRef]
  40. Gharibi, S.; Tabatabaei, B.E.S.; Saeidi, G.; Goli, S.A.H. Effect of Drought Stress on Total Phenolic, Lipid Peroxidation, and Antioxidant Activity of Achillea Species. Appl. Biochem. Biotechnol. 2016, 178, 796–809. [Google Scholar] [CrossRef]
  41. Villordon, A.; Gregorie, J.C.; LaBonte, D.; Khan, A.; Selvaraj, M. Variation in ‘Bayou Belle’ and ‘Beauregard’ Sweetpotato Root Length in Response to Experimental Phosphorus Deficiency and Compacted Layer Treatments. HortScience 2018, 53, 1534–1540. [Google Scholar] [CrossRef]
  42. Liu, H.; Si, C.; Shi, C.; Wang, S.; Sun, Z.; Shi, Y. Switch from Apoplasmic to Symplasmic Phloem Unloading during Storage Roots Formation and Bulking of Sweetpotato. Crop Sci. 2019, 59, 675–683. [Google Scholar] [CrossRef]
  43. Gajanayake, B.; Reddy, K.R.; Shankle, M.W.; Arancibia, R.A. Early-Season Soil Moisture Deficit Reduces Sweetpotato Storage Root Initiation and Development. HortScience 2013, 48, 1457–1462. [Google Scholar] [CrossRef]
  44. Sinkovič, L.; Pipan, B.; Meglič, V.; Kunstelj, N.; Nečemer, M.; Zlatić, E.; Žnidarčič, D. Genetic Differentiation of Slovenian Sweet Potato Varieties (Ipomoea batatas) and Effect of Different Growing Media on Their Agronomic and Nutritional Traits. Ital. J. Agron. 2017, 12, 350–356. [Google Scholar] [CrossRef]
  45. Siose, T.K.; Guinto, D.F. Performance of Improved Sweetpotato (Ipomoea batatas L.) Cultivars under Different Soil Types of Samoa. S. Pac. J. Nat. App. Sci. 2017, 35, 1–9. [Google Scholar] [CrossRef]
  46. Wang, Z.; Fu, J.; He, M.; Tian, Q.; Cao, H. Effects of Source/Sink Manipulation on Net Photosynthetic Rate and Photosynthate Partitioning during Grain Filling in Winter Wheat. Biol. Plant. 1997, 39, 379–385. [Google Scholar] [CrossRef]
  47. Bertin, N.; Gautier, H.; Roche, C. Number of Cells in Tomato Fruit Depending on Fruit Position and Source-Sink Balance during Plant Development. Plant Growth Regul. 2002, 36, 105–112. [Google Scholar] [CrossRef]
  48. Sakamoto, M.; Suzuki, T. Effect of Nutrient Solution Concentration on the Growth of Hydroponic Sweetpotato. Agronomy 2020, 10, 1708. [Google Scholar] [CrossRef]
  49. Gajanayake, B.; Reddy, K.R.; Shankle, M.W.; Arancibia, R.A.; Villordon, A.O. Quantifying Storage Root Initiation, Growth, and Developmental Responses of Sweetpotato to Early Season Temperature. Agron. J. 2014, 106, 1795–1804. [Google Scholar] [CrossRef]
  50. Eguchi, T.; Kitano, M.; Eguchi, H. Growth of Tuberous Root as Affected by the Ambient Humidity in Sweetpotato (Ipomoea batatas Lam.). Environ. Control. Biol. 1999, 37, 197–201. [Google Scholar] [CrossRef]
  51. Noda, T.; Kobayashi, T.; Suda, I. Effect of Soil Temperature on Starch Properties of Sweet Potatoes. Carbohydr. Polym. 2001, 44, 239–246. [Google Scholar] [CrossRef]
  52. Orman-Ligeza, B.; Morris, E.C.; Parizot, B.; Lavigne, T.; Babé, A.; Ligeza, A.; Klein, S.; Sturrock, C.; Xuan, W.; Novák, O.; et al. The Xerobranching Response Represses Lateral Root Formation When Roots Are Not in Contact with Water. Curr. Biol. 2018, 28, 3165–3173.e5. [Google Scholar] [CrossRef] [PubMed]
  53. Rodriguez-Concepcion, M.; Stange, C. Biosynthesis of Carotenoids in Carrot: An Underground Story Comes to Light. Arch. Biochem. Biophys. 2013, 539, 110–116. [Google Scholar] [CrossRef] [PubMed]
  54. Hozyo, Y.; Kato, S. Thkenig Growth and Re-thickening Growth of Tuberous Roots of Sweet Potato Pants (Ipomoea batatas Poiret). Jpn. J. Crop Sci. 1976, 45, 131–138. [Google Scholar] [CrossRef]
  55. Fuentes, P.; Pizarro, L.; Moreno, J.C.; Handford, M.; Rodriguez-Concepcion, M.; Stange, C. Light-Dependent Changes in Plastid Differentiation Influence Carotenoid Gene Expression and Accumulation in Carrot Roots. Plant Mol. Biol. 2012, 79, 47–59. [Google Scholar] [CrossRef] [PubMed]
  56. Noh, S.A.; Lee, H.-S.; Kim, Y.-S.; Paek, K.-H.; Shin, J.S.; Bae, J.M. Down-Regulation of the IbEXP1 Gene Enhanced Storage Root Development in Sweetpotato. J. Exp. Bot. 2013, 64, 129–142. [Google Scholar] [CrossRef]
  57. Yang, Y.; Zhu, J.; Sun, L.; Kong, Y.; Chen, J.; Zhu, M.; Xu, T.; Li, Z.; Dong, T. Progress on Physiological and Molecular Mechanisms of Storage Root Formation and Development in Sweetpotato. Sci. Hortic. 2023, 308, 111588. [Google Scholar] [CrossRef]
  58. Wahid, A.; Close, T.J. Expression of Dehydrins under Heat Stress and Their Relationship with Water Relations of Sugarcane Leaves. Biol. Plant 2007, 51, 104–109. [Google Scholar] [CrossRef]
  59. Rodríguez, P.; Torrecillas, A.; Morales, M.A.; Ortuño, M.F.; Sánchez-Blanco, M.J. Effects of NaCl Salinity and Water Stress on Growth and Leaf Water Relations of Asteriscus maritimus Plants. Environ. Exp. Bot. 2005, 53, 113–123. [Google Scholar] [CrossRef]
  60. Egilla, J.N.; Davies, F.T.; Boutton, T.W. Drought Stress Influences Leaf Water Content, Photosynthesis, and Water-Use Efficiency of Hibiscus Rosa-Sinensis at Three Potassium Concentrations. Photosynthetica 2005, 43, 135–140. [Google Scholar] [CrossRef]
  61. Ghori, N.-H.; Ghori, T.; Hayat, M.Q.; Imadi, S.R.; Gul, A.; Altay, V.; Ozturk, M. Heavy Metal Stress and Responses in Plants. Int. J. Environ. Sci. Technol. 2019, 16, 1807–1828. [Google Scholar] [CrossRef]
  62. Yokawa, K.; Fasano, R.; Kagenishi, T.; Baluška, F. Light as Stress Factor to Plant Roots–Case of Root Halotropism. Front. Plant Sci. 2014, 5, 718. [Google Scholar] [CrossRef] [PubMed]
  63. Kobayashi, K.; Baba, S.; Obayashi, T.; Sato, M.; Toyooka, K.; Keränen, M.; Aro, E.-M.; Fukaki, H.; Ohta, H.; Sugimoto, K.; et al. Regulation of Root Greening by Light and Auxin/Cytokinin Signaling in Arabidopsis. Plant Cell 2012, 24, 1081–1095. [Google Scholar] [CrossRef] [PubMed]
  64. Ma, Y.; Ma, X.; Gao, X.; Wu, W.; Zhou, B. Light Induced Regulation Pathway of Anthocyanin Biosynthesis in Plants. Int. J. Mol. Sci. 2021, 22, 11116. [Google Scholar] [CrossRef]
  65. Ruan, Y.; Chen, K.; Su, Y.; Jiang, S.; Xu, P.; Murray, J.D. A Root Tip-Specific Expressing Anthocyanin Marker for Direct Identification of Transgenic Tissues by the Naked Eye in Symbiotic Studies. Plants 2021, 10, 605. [Google Scholar] [CrossRef]
  66. Shi, L.; Li, X.; Fu, Y.; Li, C. Environmental Stimuli and Phytohormones in Anthocyanin Biosynthesis: A Comprehensive Review. Int. J. Mol. Sci. 2023, 24, 16415. [Google Scholar] [CrossRef]
  67. Gao, H.-N.; Jiang, H.; Cui, J.-Y.; You, C.-X.; Li, Y.-Y. Review: The Effects of Hormones and Environmental Factors on Anthocyanin Biosynthesis in Apple. Plant Sci. 2021, 312, 111024. [Google Scholar] [CrossRef]
  68. Sun, J.; Hui, K.; Guo, Z.; Li, Y.; Fan, X. Cellulose and Lignin Contents Are Negatively Correlated with Starch Accumulation, and Their Correlation Characteristics Vary Across Cassava Varieties. J. Plant Growth Regul. 2023, 42, 658–669. [Google Scholar] [CrossRef]
  69. Singh, V.; Sergeeva, L.; Ligterink, W.; Aloni, R.; Zemach, H.; Doron-Faigenboim, A.; Yang, J.; Zhang, P.; Shabtai, S.; Firon, N. Gibberellin Promotes Sweetpotato Root Vascular Lignification and Reduces Storage-Root Formation. Front. Plant Sci. 2019, 10, 1320. [Google Scholar] [CrossRef]
  70. Higuchi, K.; Kanai, M.; Tsuchiya, M.; Ishii, H.; Shibuya, N.; Fujita, N.; Nakamura, Y.; Suzui, N.; Fujimaki, S.; Miwa, E. Common Reed Accumulates Starch in Its Stem by Metabolic Adaptation under Cd Stress Conditions. Front. Plant Sci. 2015, 6, 138. [Google Scholar] [CrossRef]
  71. Buckseth, T.; Sharma, A.K.; Pandey, K.K.; Singh, B.P.; Muthuraj, R. Methods of Pre-Basic Seed Potato Production with Special Reference to Aeroponics—A Review. Sci. Hortic. 2016, 204, 79–87. [Google Scholar] [CrossRef]
  72. Ravi, V.; Chakrabarti, S.K.; Makeshkumar, T.; Saravanan, R. Molecular Regulation of Storage Root Formation and Development in Sweet Potato. In Horticultural Reviews: Volume 42; John Wiley & Sons, Ltd.: Hoboken, NJ, USA, 2014; pp. 157–208. ISBN 978-1-118-91682-7. [Google Scholar]
  73. Michael Turner, G.; Tran, V.; Sun, W.; Federico Rosas, O.; Ravindran, H.; Neculăescu, A.-M.; Kougianos, A.; Keys, S.; Hawkey, M.; Alexandrou, N. Lunar Agriculture: Farming for the Future. In Earth and Space 2021; American Society of Civil Engineers: Reston, VA, USA, 2021; pp. 639–652. [Google Scholar] [CrossRef]
  74. Wilson, C.D.; Pace, R.D.; Bromfield, E.; Jones, G.; Lu, J.Y. Sweet Potato in a Vegetarian Menu Plan for NASA’s Advanced Life Support Program. Life Support Biosph. Sci. 1998, 5, 347–351. [Google Scholar] [PubMed]
  75. JAXA (Japan Aerospace Exploration Agency). Release of the Lunar Farm Working Group Review Report. 2019. Available online: https://www.ihub-tansa.jaxa.jp/english/Lunarfarming_en.html (accessed on 12 September 2024).
  76. Kitaya, Y.; Higashi, K.; Shibuya, T.; Endo, R. Fundamental Study on Plant-Based Regenerative Life-Support Systems with Sweetpotato Culture in Space. Aerosp. Technol. Jpn. 2021, 19, 889–891. [Google Scholar] [CrossRef]
  77. Miyajima, H. 7. Study of the Lunar Farming System. Study Working Group. 2023, pp. 68–83. Available online: https://jaxa.repo.nii.ac.jp/record/2000110/files/AA2330006000.pdf#page=73 (accessed on 12 September 2024).
  78. Paul, A.-L.; Elardo, S.M.; Ferl, R. Plants Grown in Apollo Lunar Regolith Present Stress-Associated Transcriptomes That Inform Prospects for Lunar Exploration. Commun. Biol. 2022, 5, 382. [Google Scholar] [CrossRef] [PubMed]
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Figure 1. (A) Hydroponic systems used in this study: five cultivation systems were used, including Pot, SS (soil sheet), SD (soilless dark), SL (soilless light), and DFT (deep flow technique). (B) Experimental timeline in this study.
Figure 1. (A) Hydroponic systems used in this study: five cultivation systems were used, including Pot, SS (soil sheet), SD (soilless dark), SL (soilless light), and DFT (deep flow technique). (B) Experimental timeline in this study.
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Figure 2. Environmental conditions in different cultivation systems: (A) typical daily temperature fluctuations and (B) light intensity (lux, visible light intensity) changes under varying weather conditions for each system. (C) The spectral properties of light wavelengths were compared between the SL system and direct sunlight (outside).
Figure 2. Environmental conditions in different cultivation systems: (A) typical daily temperature fluctuations and (B) light intensity (lux, visible light intensity) changes under varying weather conditions for each system. (C) The spectral properties of light wavelengths were compared between the SL system and direct sunlight (outside).
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Figure 3. Time-course changes in the morphology of the sweet potato storage roots cultivated in three different systems (SS, SD, and SL). The scale bars represent 5 cm.
Figure 3. Time-course changes in the morphology of the sweet potato storage roots cultivated in three different systems (SS, SD, and SL). The scale bars represent 5 cm.
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Figure 4. Time-course changes in the diameter of the sweet potato storage roots cultivated in three different systems. The red arrow indicates the start of exposure to light in the SL system. The vertical bars represent ± SE (n = 24). The different letters at each timepoint indicate significant differences between the systems (p < 0.05, Tukey–Kramer test). The absence of letters indicates no significant differences.
Figure 4. Time-course changes in the diameter of the sweet potato storage roots cultivated in three different systems. The red arrow indicates the start of exposure to light in the SL system. The vertical bars represent ± SE (n = 24). The different letters at each timepoint indicate significant differences between the systems (p < 0.05, Tukey–Kramer test). The absence of letters indicates no significant differences.
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Figure 5. Color parameters of the sweet potato storage root epidermis cultivated in three different systems. Time-course changes in the (A) L*, (B) a*, and (C) b* values of the storage root epidermis. The red arrow indicates the start of exposure to light in the SL system. The vertical bars represent ± SE (n = 24). The different letters at each timepoint indicate significant differences between systems (p < 0.05, Tukey–Kramer test). The absence of letters indicates no significant differences. The color coordinates (a* = red–green axis and b* = yellow–blue axis) of the storage root epidermis were measured at (D) 54, (E) 58, (F) 96, and (G) 150 days after transplantation.
Figure 5. Color parameters of the sweet potato storage root epidermis cultivated in three different systems. Time-course changes in the (A) L*, (B) a*, and (C) b* values of the storage root epidermis. The red arrow indicates the start of exposure to light in the SL system. The vertical bars represent ± SE (n = 24). The different letters at each timepoint indicate significant differences between systems (p < 0.05, Tukey–Kramer test). The absence of letters indicates no significant differences. The color coordinates (a* = red–green axis and b* = yellow–blue axis) of the storage root epidermis were measured at (D) 54, (E) 58, (F) 96, and (G) 150 days after transplantation.
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Figure 6. Sweet potato roots cultivated in five different systems at 151 DAT. In the Pot, SS, SD, and SL systems, most fibrous roots were removed before photographing, whereas in the DFT system, all the fibrous roots were included in the image.
Figure 6. Sweet potato roots cultivated in five different systems at 151 DAT. In the Pot, SS, SD, and SL systems, most fibrous roots were removed before photographing, whereas in the DFT system, all the fibrous roots were included in the image.
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Figure 7. (A) Storage root diameter, (B) storage root weight, (C) storage root yield, and (D) storage root moisture content of sweet potatoes cultivated in four different systems at 151 DAT. Vertical bars represent ± SE (n = 12). Different letters indicate significant differences between systems (p < 0.05, Tukey–Kramer test).
Figure 7. (A) Storage root diameter, (B) storage root weight, (C) storage root yield, and (D) storage root moisture content of sweet potatoes cultivated in four different systems at 151 DAT. Vertical bars represent ± SE (n = 12). Different letters indicate significant differences between systems (p < 0.05, Tukey–Kramer test).
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Figure 8. Color parameters of sweet potato storage root epidermis at 151 days after transplantation: (A) L*, (B) a*, and (C) b* values were measured in four systems (Pot, SS, SD, and SL), with SL separated into top (SL-T) and bottom (SL-B) sides. Vertical bars represent ± SE (n = 24). Different letters indicate significant differences between systems (p < 0.05, Tukey–Kramer test).
Figure 8. Color parameters of sweet potato storage root epidermis at 151 days after transplantation: (A) L*, (B) a*, and (C) b* values were measured in four systems (Pot, SS, SD, and SL), with SL separated into top (SL-T) and bottom (SL-B) sides. Vertical bars represent ± SE (n = 24). Different letters indicate significant differences between systems (p < 0.05, Tukey–Kramer test).
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Figure 9. Morphological comparison of the sweet potato storage roots at 151 DAT in four different systems; the SL roots were analyzed separately for the top (SL-T) and bottom (SL-B) sides: (A) External view of the storage roots (scale bars: 3 cm). (B) Cross-sections (scale bars: 2 cm). (C) Stereo microscopic images of the cross-sections (scale bars: 100 μm). (D) Peeled surface of the storage roots (scale bars: 0.5 cm).
Figure 9. Morphological comparison of the sweet potato storage roots at 151 DAT in four different systems; the SL roots were analyzed separately for the top (SL-T) and bottom (SL-B) sides: (A) External view of the storage roots (scale bars: 3 cm). (B) Cross-sections (scale bars: 2 cm). (C) Stereo microscopic images of the cross-sections (scale bars: 100 μm). (D) Peeled surface of the storage roots (scale bars: 0.5 cm).
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Figure 10. (A) Total chlorophyll content, (B) total phenolic content, (C) anthocyanin content, and (D) total starch content of sweet potato storage roots at 151 DAT in four different systems (Pot, SS, SD, and SL). SL roots were measured separately for SL-T and SL-B in (A–C). Vertical bars represent ± SE (n = 4 for (A,C); n = 6 for (B,D)). Different letters indicate significant differences between systems (p < 0.05, Tukey–Kramer test).
Figure 10. (A) Total chlorophyll content, (B) total phenolic content, (C) anthocyanin content, and (D) total starch content of sweet potato storage roots at 151 DAT in four different systems (Pot, SS, SD, and SL). SL roots were measured separately for SL-T and SL-B in (A–C). Vertical bars represent ± SE (n = 4 for (A,C); n = 6 for (B,D)). Different letters indicate significant differences between systems (p < 0.05, Tukey–Kramer test).
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Figure 11. Phloroglucinol staining for lignin in cross-sections of sweet potato storage roots at 151 DAT in four different systems. Scale bars represent 1 cm.
Figure 11. Phloroglucinol staining for lignin in cross-sections of sweet potato storage roots at 151 DAT in four different systems. Scale bars represent 1 cm.
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Figure 12. Above-ground growth of sweet potato plants cultivated in five different systems: (A) Time-course changes in maximum stem length. (B) Maximum stem length, (C) above-ground fresh weight, and (D) above-ground moisture content of sweet potato plants measured at 151 DAT. Vertical bars represent ± SE (n = 12). Different letters (at each timepoint) indicate significant differences between systems (p < 0.05, Tukey–Kramer test). Absence of letters indicates no significant differences.
Figure 12. Above-ground growth of sweet potato plants cultivated in five different systems: (A) Time-course changes in maximum stem length. (B) Maximum stem length, (C) above-ground fresh weight, and (D) above-ground moisture content of sweet potato plants measured at 151 DAT. Vertical bars represent ± SE (n = 12). Different letters (at each timepoint) indicate significant differences between systems (p < 0.05, Tukey–Kramer test). Absence of letters indicates no significant differences.
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Agriengineering 06 00222 g013
Figure 13. Stem sections of sweet potato plants at 151 DAT in four different systems: (A) External view of stem sections (scale bars: 1 cm). (B) Stereo microscopic images of lignin-stained stem sections by phloroglucinol (scale bars: 200 μm). (C) Stereo microscopic images of starch-stained stem sections by Lugol’s iodine (scale bars: 200 μm).
Figure 13. Stem sections of sweet potato plants at 151 DAT in four different systems: (A) External view of stem sections (scale bars: 1 cm). (B) Stereo microscopic images of lignin-stained stem sections by phloroglucinol (scale bars: 200 μm). (C) Stereo microscopic images of starch-stained stem sections by Lugol’s iodine (scale bars: 200 μm).
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MDPI and ACS Style

Sakamoto, M.; Suzuki, T. Impacts of Light Exposure and Soil Covering on Sweet Potato Storage Roots in a Novel Soilless Culture System. AgriEngineering 2024, 6, 3912-3930. https://doi.org/10.3390/agriengineering6040222

AMA Style

Sakamoto M, Suzuki T. Impacts of Light Exposure and Soil Covering on Sweet Potato Storage Roots in a Novel Soilless Culture System. AgriEngineering. 2024; 6(4):3912-3930. https://doi.org/10.3390/agriengineering6040222

Chicago/Turabian Style

Sakamoto, Masaru, and Takahiro Suzuki. 2024. "Impacts of Light Exposure and Soil Covering on Sweet Potato Storage Roots in a Novel Soilless Culture System" AgriEngineering 6, no. 4: 3912-3930. https://doi.org/10.3390/agriengineering6040222

APA Style

Sakamoto, M., & Suzuki, T. (2024). Impacts of Light Exposure and Soil Covering on Sweet Potato Storage Roots in a Novel Soilless Culture System. AgriEngineering, 6(4), 3912-3930. https://doi.org/10.3390/agriengineering6040222

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