Molecular Plant Advance Access originally published online on June 7, 2007
Molecular Plant 2008 1(1):15-26; doi:10.1093/mp/ssm001
| ||||||||||||||||||||||||||||||||||||||||||||||||||||
© The Author 2007. Published by Oxford University Press on behalf of CSPP and IPPE, SIBS, CAS.
Leaf Positioning of Arabidopsis in Response to Blue Light
a Department of Biology, Faculty of Science, Kyushu University, Ropponmatsu, Fukuoka, 810-8560 Japan
b Research and Development Center for Higher Education, Kyushu University, Ropponmatsu, Fukuoka, 810-8560 Japan
1 To whom correspondence should be addressed. E-mail kenrcb{at}mbox.nc.kyushu-u.ac.jp, fax 81-92-726-4758.
 |
Abstract |
|---|
 TOP Abstract INTRODUCTIONRESULTSDISCUSSIONMETHODS |
|---|
Appropriate leaf positioning is essential for optimizing photosynthesis and plant growth. However, it has not been elucidated how green leaves reach and maintain their position for capturing light. We show here the regulation of leaf positioning under blue light stimuli. When 1-week-old Arabidopsis seedlings grown under white light were transferred to red light (25 µmol m–2 s–1) for 5 d, new petioles that appeared were almost horizontal and their leaves were curled and slanted downward. However, when a weak blue light from above (0.1 µmol m–2 s–1) was superimposed on red light, the new petioles grew obliquely upward and the leaves were flat and horizontal. The leaf positioning required both phototropin1 (phot1) and nonphototropic hypocotyl 3 (NPH3), and resulted in enhanced plant growth. In an nph3 mutant, neither optimal leaf positioning nor leaf flattening by blue light was found, and blue light-induced growth enhancement was drastically reduced. When blue light was increased from 0.1 to 5 µmol m–2 s–1, normal leaf positioning and leaf flattening were induced in both phot1 and nph3 mutants, suggesting that phot2 signaling became functional and that the signaling was independent of phot1 and NPH3 in these responses. When plants were irradiated with blue light (0.1 µmol m–2 s–1) from the side and red light from above, the new leaves became oriented toward the source of blue light. When we transferred these plants to both blue light and red light from above, the leaf surface changed its orientation to the new blue light source within a few hours, whereas the petioles initially were unchanged but then gradually rotated, suggesting the plasticity of leaf positioning in response to blue light. We showed the tissue expression of NPH3 and its plasma membrane localization via the coiled-coil domain and the C-terminal region. We conclude that NPH3-mediated phototropin signaling optimizes the efficiency of light perception by inducing both optimal leaf positioning and leaf flattening, and enhances plant growth.
|
INTRODUCTION |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
|---|
Plants respond appropriately to ever-changing environments by morphogenesis, movement, changes in cellular components, and metabolic activity, thereby optimizing growth in natural environments. Plants respond by sensing changes in light, gravity, temperature, salt, and water status through individual receptors. Light is the most important factor influencing plant life, and wide ranges in wavelength from UV-A to far-red light are perceived by several photoreceptors to recognize the light environment. Blue light induces various developmental and movement responses, including phototropic bending, cotyledon opening, photoperiodic flowering, leaf flattening, de-etiolation, stomatal opening, chloroplast movements, anthocyanin accumulation, gene expression, and the inhibition of hypocotyl elongation (Cashmore et al., 1999; Briggs and Christie, 2002; Lin, 2002; Wang and Deng, 2002). In Arabidopsis plants, three classes of major blue light receptors—cryptochromes, phototropins, and FKF1/ZTL/LKP2 (Imaizumi et al., 2003)—are responsible for the responses mentioned above.
Cryptochrome was identified as the first plant blue light receptor using an Arabidopsis mutant that did not show hypocotyl growth inhibition in response to blue light (Ahmad and Cashmore, 1993), and later it turned out to act as an animal blue light receptor to regulate the circadian clock and other responses (Cashmore et al., 1999). Cryptochromes (cry1 and cry2) in plants act together with the red/far-red light receptor phytochromes to regulate photomorphogenic responses based on multiple gene expression (Lin, 2002; Nemhauser and Chory, 2002; Wang and Deng, 2002).
Phototropin1 (phot1) was identified as a plant-specific blue light receptor using an Arabidopsis mutant that showed impaired phototropic bending in response to blue light (Liscum and Briggs, 1995; Huala et al., 1997). Phototropin is a serine/threonine protein kinase in the C-terminus, with two LOV (light, oxygen, voltage) domains as the binding sites of the chromophore flavin mononucleotide (FMN) in the N-terminus. Later, phototropin2 (phot2) was found as a photoreceptor that mediates the photoavoidance response of chloroplasts to prevent strong light from damaging the photosynthetic machinery (Jarillo et al., 2001; Kagawa et al., 2001; Kasahara et al., 2002). In general, phot1 functions under a low intensity of blue light, and phot2 under a relatively high intensity. Phot1 and phot2 act redundantly and cover wide ranges of light intensity in phototropism, chloroplast accumulation, stomatal opening, and leaf flattening (Kagawa et al., 2001; Kinoshita et al., 2001; Sakai et al., 2001; Sakamoto and Briggs, 2002). Furthermore, phot1 alone acts as a blue light receptor in the rapid inhibition of hypocotyl elongation, followed by the cryptochrome action in the much slower response (Folta and Spalding, 2001), and is required for blue light-mediated destabilization of Lhcb and rbcL transcripts at high intensities (Folta and Kaufman, 2003). All these responses probably serve to optimize photosynthesis, and a dramatic plant growth enhancement mediated by phototropin is demonstrated under a low intensity of photosynthetically active radiation (PAR) (Takemiya et al., 2005).
Extensive studies on phototropism were done using etiolated hypocotyls and coleoptiles as model systems, and in many cases blue light was provided from the lateral side because it is easy to measure and analyze the responses (Fankhauser and Casal, 2004; Vandenbussche et al., 2005). These investigations have provided detailed information on phototropic bending at the physiological and biochemical levels. Although phototropism, together with other phototropin-mediated responses, has an important role in maximizing light capture by green leaves, most of the experimental work has been done without considering green leaf behavior and development. Therefore, it becomes important to elucidate the functional roles of blue light in more developed stages of plants with green leaves. However, the behavior of green leaves in response to blue light has not been investigated, nor has an attempt been made to formulate the optimal position to maximize photosynthesis in response to blue light when leaves are newly developed.
In this study, we established the experimental conditions that allow the appearance of new leaves, and investigated blue light's effects on the development of green leaves when the light was provided from above. We showed that, in response to a weak blue light, newly emerged leaves exhibit the appropriate positioning and leaf flattening to increase light capturing efficiency. We also showed that these responses are mediated by nonphototropic hypocotyl 3 (NPH3) via the phot1 pathway and probably enhance growth.
|
RESULTS |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
|---|
Blue Light-Dependent Leaf Positioning Increases Light Capture
We grew Arabidopsis seedlings under white light at 50 µmol m–2 s–1 for 7 d and induced de-etiolation. The de-etiolated plants each had a pair of open cotyledons and undeveloped first true leaves (data not shown). We then transferred these green plants to red light from above at 25 µmol m–2 s–1 with or without low-intensity blue light (0.1 µmol m–2 s–1) and kept them growing for 5 d to allow the appearance of new first true leaves. Slightly arched new petioles grew nearly horizontally, and the first true leaflets slanted down without blue light (Figure 1A, left). However, straight new petioles grew obliquely upward, and the new leaflets faced toward the light source when the blue light was supplemented with red light (Figure 1A, right). These results suggest that blue light from above oriented the leaf surface perpendicular to the light direction by inducing both the straight and upward growth of petioles. We refer to these responses as leaf positioning.
We measured the angle of a petiole of a first true leaf from the horizontal (
), illustrated in Figure 1B, to express an index of leaf positioning. The angles were nearly 45° in the presence of blue light and <10° in the absence of blue light. The blue light-dependent leaf positioning increased the area of light interception 2-fold in each first leaf when the blue light was provided together with red light from the top (Figure 1C and D).
|
We next illuminated the plants with blue light (0.1 µmol m–2 s–1) from the side but red light from the top as before. New petioles emerged and the new leaflets became oriented toward the blue light source, but the face of the leaf was not completely perpendicular to that source (Figure 1E, solid arrowheads). The surface of a pair of open cotyledons became partially oriented to the blue light (Figure 1E, open arrowheads).
From these results, we conclude that the plant determines the orientation of a newly developed leaf through the perception of blue light.
Phototropin1 (phot1) Mediates the Optimal Leaf Positioning Under Low Blue Light
Phototropins optimize photosynthesis and promote plant growth by inducing blue light-mediated multiple physiological responses at the same time (Briggs and Christie, 2002; Takemiya et al., 2005). We thus expected that phototropins might function in the leaf-positioning response shown above. To test this hypothesis, we grew phototropin mutant plants under the same growth conditions. As expected, the optimal leaf positioning for capturing light was not found in either a phot1 phot2 double mutant (phot1-5 phot2-1) or a phot1 mutant (phot1-5), but was found in phot2 (phot2-1) and cry1 cry2 double mutants (hy4-3 cry2-1) (Figure 2B and C). Without blue light, none of these plants showed the normal leaf positioning and their leaves slanted down (Figure 2A). These results indicate that the blue light-induced leaf positioning is mediated by phot1, and neither phot2 nor cryptochromes are involved in the response under our growth conditions.
|
NPH3 Mediates Phot1-Dependent Leaf Positioning
Since blue light-dependent leaf positioning is mediated by phot1, we wished to identify the components downstream of phot1 by isolating the mutants that lack the upward petiole growth. We obtained two mutant lines: an ethylmethane sulfonate (EMS)-mutagenized plant and a T-DNA insertional plant, both of which showed impairment in the upward petiole growth (Figure 3A). By crossing the two mutants, we found that the two mutations are allelic to each other.
|
To identify the mutated gene, we performed thermal asymmetric interlaced (TAIL)-PCR using the genomic DNA prepared from the T-DNA insertional mutant. We found that T-DNA was in the fifth exon of the NPH3 gene and confirmed that this line was a null nph3 mutant by reverse transcription (RT)-PCR (Figure 3B and C). Because the EMS-mutagenized mutant is allelic to the T-DNA insertional line, we cloned and sequenced the full-length NPH3 cDNA from the EMS-mutagenized mutant and found that the mutant had a single nucleotide substitution of cytosine to thymine in the last exon (Figure 3B). This substitution produced a stop codon on Gln681 in the coiled-coil domain of the NPH3 protein.
We tested the functional complementation of the nph3 mutation by the wild-type genomic NPH3 gene. A 5400 bp genomic NPH3 fragment containing the 5’ and 3’ noncoding regions was introduced into the two distinct mutants. The transformed lines in the T3 generation restored normal leaf positioning with upward petioles (Figure 3D). The results demonstrate that our mutants are allelic to the nph3 mutant and that NPH3 functions as a signal component in phot1-mediated leaf positioning. We thus named the EMS-mutagenized and the T-DNA insertional mutants as nph3-201 and nph3-202, respectively (Figure 3).
Expression of NPH3
We investigated the expression of NPH3 mRNA by RT-PCR using wild-type Arabidopsis plants. The NPH3 mRNA was highly expressed in mesophyll cells, leaves, stems, and roots, but only a small amount was expressed in guard cells (Figure 4A). The results agree with observations that NPH3 functions mainly in the leaf and petiole (Figure 3A), and that NPH3 does not act in stomata (Inada et al., 2004).
|
Subcellular Localization of NPH3
To investigate the subcellular localization of NPH3 protein, we transiently expressed NPH3 fused with green fluorescent protein (GFP) in epidermal cells and guard cells of Vicia faba by particle bombardment. The fluorescence from full-length NPH3 was found on the periphery of both epidermal and guard cells, suggesting plasma membrane localization of NPH3 as described previously (Motchoulski and Liscum, 1999; Lariguet et al., 2006; Figure 4B, full length). We then investigated the localization in more detail using guard cells because the transient expression of NPH3 is much easier in them than in epidermal cells. The fluorescence of mutant NPH3-201 protein from nph3-201 was observed as many particles in cytosolic compartments (Figure 4B, NPH3-201). Since the mutant NPH3-201 protein may lack a C-terminal region downstream from the coiled-coil domain (Figure 3B), it is possible that this region is required for the membrane localization of NPH3. To test this, we expressed the NPH3 C-terminal fragment containing the coiled-coil domain (coiled-coil-C) fused with GFP. As expected, the fluorescent signal of this region was found on the plasma membrane (Figure 4B, coiled-coil-C). We then divided this coiled-coil-C into a coiled-coil domain (coiled-coil) and a C-terminal region (C-terminus) and expressed these as above. The GFP fluorescence of the coiled-coil domain was detected mainly in the plasma membrane and slightly in the cytoplasm (Figure 4B, Coiled-coil). The fluorescence of the C-terminus was found in both the cytosol and the plasma membrane (Figure 4B, C-terminus), and the distribution was different from that of GFP alone, which showed a clear cytosolic localization (Figure 4B, sGFP). These observations suggest that both the conserved coiled-coil domain and the C-terminal region probably function to localize NPH3 protein on the plasma membrane, and the membrane localization may be needed for the function of NPH3 (Figure 3A).
Recovery of Leaf Positioning in nph3 Mutants Under High Intensity Blue Light
We found that the petioles in nph3-201 and nph3-202 grew upward and exhibited almost wild-type leaf positioning when supplemented blue light was increased to 5 µmol m–2 s–1 from 0.1 µmol m–2 s–1 (Figure 5A). Quantitative data indicate that phot1-5, nph3-201, and nph3-202 largely restored the wild-type leaf positioning at relatively high fluence rates of blue light, whereas phot1-5 phot2-1 did not (Figure 5B). These results suggest that phot2 becomes functional and mediates the leaf positioning in response to the higher intensity of blue light. They also suggest that NPH3 functions principally through the phot1-dependent pathway in the response.
|
NPH3 Mediates Leaf Flattening Only Under Low Blue Light
Under our low blue light growth conditions (25 µmol m–2 s–1 red light with 0.1 µmol m–2 s–1 blue light), leaves of nph3-201 and nph3-202 curled, as did leaves of phot1-5 and phot1-5 phot2-1 mutants. This phenotype became more prominent when these plants were further grown for another 5 d (Figure 6A). In contrast, gl1, Col, WS, and phot2-1 exhibited flattened leaves under the same conditions. All of these plants showed curled leaves under red light alone (data not shown). These results suggest that NPH3 functions in leaf flattening through the phot1-mediated pathway.
|
When the intensity of supplemental blue light was increased to 5 µmol m–2 s–1, leaves of nph3-201, nph3-202, and phot1-5 became flattened, but those of the phot1-5 phot2-1 double mutant remained curled (Figure 6B). These results indicate that leaf flattening is mediated by phot2 under a relatively high intensity of blue light, and that this phot2-dependent leaf flattening is not mediated by NPH3.
Contribution of NPH3 to Growth Enhancement Under Low Blue Light
NPH3 mediates both horizontal leaf positioning and leaf flattening in response to very weak blue light (Figures 3A and 6A), but does not mediate chloroplast movement or stomatal opening (Inada et al., 2004). All these blue light responses are known to increase photosynthesis and plant growth in a low-light environment in particular (Takemiya et al., 2005). Taking advantage of the properties of nph3 mutants, we evaluated the contributions of leaf positioning and flattening to growth enhancement. We measured the fresh weights of the wild type (gl1) and of nph3-201, nph3-6, and phot1-5 mutants that had been grown under our conditions for 5 weeks. As shown in Figure 7A and B, the wild-type plants showed 2.5-fold growth enhancement by the addition of 0.1 µmol m–2 s–1 blue light to the red light, but no actual growth enhancement was found in the phot1-5 mutant. Interestingly, the nph3-201 and nph3-6 mutants showed slight but significant growth enhancement in response to very weak blue light (Figure 7B). This slight growth enhancement may have been brought about by both chloroplast movement and stomatal opening, because in the nph3 mutants chloroplasts gathered at the surface of mesophyll cells and stomata opened in response to blue light (Figure 7C and D; Inada et al., 2004). The growth difference between wild-type plants and nph3 mutants is probably provided by the leaf positioning and leaf flattening that were mediated by NPH3. These results further suggest that growth enhancement in response to a weak blue light is brought about mainly through the function of NPH3, as both responses tend to maximize light interception.
|
Reversibility of Leaf Positioning in Response to Blue Light
It is unclear whether the leaf positioning responses shown above are reversible or not. To test this, we utilized plants that had been grown under irradiation with blue light from the side and red light from above, as indicated in Figure 1E. The surfaces of the first true leaves of the plants were oriented toward the source of blue light (Figure 8A, 0 h). Such leaf orientation in response to blue light was not found in the mutants of phot1-5 or nph3-201 (data not shown). Then, we transferred these plants to both red (25 µmol m–2 s–1) and blue (0.1 µmol m–2 s–1) light from above and kept them growing for another 5 d. After the second transfer, the leaf surface began to orient rapidly toward the new blue light source with a time delay of 20 min (Figure 8B, leaf angle in left graph;
L), and began a relatively slower phase after about 2 h (Figure 8A and B, 2
h). Then, the leaf surface gradually approached the maximum angle within 8 h (Figure 8B, leaf angle in left graph), and maintained this position thereafter with a very slight change (Figure 8A and B, right graph). The petiole angle in the projected image of the first leaf became almost zero in the time course, similar to the light behavior of the leaflet (Figure 8A and B, petiole angle; P). The rate of the rapid leaflet orientation was 15° h–1, which is almost the same value as that for solar-tracking responses as reported for Lavatera cretica leaves (Koller et al., 1985; Koller and Levitan, 1989; Koller, 2000). Our results suggests that the rapid leaflet orientation might be a solar-tracking response in Arabidopsis, and is mediated by phot1.
|
During the leaf repositioning responses, the petiole was arch-shaped from 4 to 24 h, a conformation that facilitated orienting the leaf surface perpendicular to the blue light from above. The petiole subsequently became straight after 48 h (Figure 8A). Although the leaf itself became oriented to the blue light source within 8 h, the petiole remained unchanged and the adaxial side was still toward the original source of blue light during this time (Figure 8A, 8 h; and C, 12 h). Afterwards, the petiole gradually rotated from 24 to 96 h, and completed its rotation within 120 h (Figure 8D). The petioles with leaves finally became aligned directly opposite each other (Figure 8C, 120 h; and D). These results suggested that the leaf positioning is plastic in response to blue light and is comprised of both a relatively rapid leaf orientation response (within 0.3–8 h) and a slow petiole rotation response (within 24–120 h). In contrast to the first true leaves, cotyledons maintained their original angles irrespective of the change in blue light direction (Figure 8A, 0–8 h).
|
DISCUSSION |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
|---|
Blue Light-Mediated Leaf Positioning Promotes Light-Capturing Efficiency
Plants control leaf position in response to environmental stimuli, such as light, gravity, and the circadian rhythm, to optimize their photosynthetic performance. However, it has not been elucidated how a plant maintains a leaf position that is optimal for capturing light energy efficiently for photosynthesis. In this study, we found that blue light induced the leaf surface into a perpendicular orientation to the light source and that the response increased the light interception (Figure 1). We also demonstrated that the response is mediated by phototropins (Figures 2 and 5). The leaf positioning was achieved by the regulation of the position of new emergent petioles and leaves (Figure 1A and E). When the source of blue light was changed from above to the side without changing the source of red light, plants oriented the new leaf surface to the source of blue light (Figure 1E). These results suggest that plants utilize blue light to determine leaf direction.
Importance of Leaf Positioning as a Means of Capturing Light
The Arabidopsis leaf positioning might comprise both rapid movement and a slow growth process, requiring a long time (several days) to establish the response (Figures 1A and 8). In this study, we grew plants for 5 d under definite conditions and determined the positions of newly emergent leaves (Figure 1). However, these experimental conditions did not produce a rapid change in position in response to blue light. To monitor the changes, we investigated the leaf positioning by moving the blue light source: plants that had been irradiated from the side were now irradiated from above (Figure 8). We found that the leaf changed its direction to the new blue light source within several hours, followed by a slow change in petiole direction after 24 h. These results suggest that the plants preferentially change leaf direction, and that such rapid regulation of leaf direction is suitable for maximizing light interception. The rapid leaf orientation Arabidopsis seems to be identical to the response reported as solar tracking in Lavatera leaves (Figure 8; Koller, 2000).
We recently reported that phototropins mediate the leaf movement of kidney bean and that the response greatly increased the light absorption of leaves (Inoue et al., 2005). The movement response is reversible and is completed in a short time (1.5 h), which is achieved by the water transport in specialized motor cells of the pulvinus (Inoue et al., 2005). Although the physiological roles of both plant responses seem to be similar (i.e. the enhancement of photosynthesis), and although the responses are mediated by the same photoreceptors, the mechanisms between leaf positioning and leaf movement may differ, since the complete Arabidopsis leaf positioning probably requires at least a few days to complete (Figure 8).
Very recently it was shown that Arabidopsis petioles move upward and that the leaf surface becomes more vertical when the plants are placed in the dark. This movement is suggested to be a shade-avoidance role in reaction to shading by neighboring leaves (Mullen et al., 2006); it is regulated by phytochrome action (Mullen et al., 2006) and/or negative gravitropism (Mano et al., 2006), and is distinct from the responses shown here.
Interestingly, the three distinct responses (two movements and positioning) mentioned above have a similar physiological role of increasing the light capture efficiency (Figure 1C–E; Mullen et al., 2006), but the reactions are induced by at least two different stimuli (blue light and darkness). It is likely that the appropriate leaf positioning is very important for plant survival and is finely controlled by the integration of various environmental stimuli including blue light, red/far-red light, and gravity in natural environments through movements and morphogenic processes.
Involvement of NPH3 in Leaf Positioning and Leaf Flattening
It has been demonstrated that NPH3 and its ortholog CPT1 are responsible for hypocotyl and coleoptile phototropism in Arabidopsis and Oryza, respectively (Motchoulski and Liscum, 1999; Haga et al., 2005). Another example of NPH3 involvement is phot1-mediated destabilization of Lhcb and rbcL transcripts (Folta and Kaufman, 2003). In the present study, we found for the first time that NPH3 mediated both leaf positioning and leaf flattening in the phot1-dependent pathway (Figures 5 and 6). In accord with these functional roles of NPH3, we showed that NPH3 is localized on the plasma membrane, on which phot1 also localizes (Sakamoto and Briggs, 2002), via the coiled-coil domain and the C-terminus (Figure 4B). The co-localization of NPH3 and phot1 on the same membrane may facilitate phot1–NPH3 complex formation and siging (Motchoulski and Liscum, 1999; Lariguet et al., 2006; Figure 3A).
NPH3 is suggested to function as a common signal component in both phot1- and phot2-dependent pathways in phototropism, since nph3 mutants showed no hypocotyl phototropism under high irradiation with blue light (Sakai et al., 2000; Inada et al., 2004). Unexpectedly, we found that the leaf positioning and leaf flattening responses were lost in nph3 mutants under a very low intensity of blue light (Figures 3A and 6A), but both responses were restored by high-intensity blue light in both the nph3 and phot1 mutants (Figures 5A and B, and 6B). These results suggest that the responses observed under a high blue light intensity might be mediated by phot2, and that an additional signal component other than NPH3 must be involved downstream from phot2.
Contribution of Responses to Phot1-Mediated Growth Enhancement
We demonstrated that the leaf positioning and leaf flattening responses actually contribute to blue light-dependent growth enhancement by increasing the amount of light captured (Figures 1C–E and 7). Our findings add a means by which to optimize photosynthesis through phototropin functions, in addition to an understanding of the physiological and morphological changes in photosynthetic tissues under various light environments (Niklas and Owens, 1989; Ballaré and Scopel, 1997).
In a previous work we demonstrated that phot1 dramatically enhances plant growth in response to a very low intensity of blue light, and that the enhancement is achieved by integrating phot1-mediated responses, including those of chloroplast accumulation (Jarillo et al., 2001; Kagawa et al., 2001; Sakai et al., 2001), stomatal opening (Kinoshita et al., 2001; Doi et al., 2004), and leaf flattening (Sakamoto and Briggs, 2002; Takemiya et al., 2005). Although we suggested that leaf flattening was the largest factor responsible for growth enhancement, we could not evaluate the contributions to growth by these distinct responses. In the present study, we found that NPH3 mediates leaf positioning and flattening but does not mediate chloroplast movement or stomatal opening. Taking advantage of this property of the nph3 mutant, we showed that this mutant slightly enhanced plant growth under our growth conditions, with active chloroplast movement and stomatal opening in the mutant (Figure 7). These results indicate that leaf flattening and positioning play an important role in maximizing photosynthesis, and that chloroplast movement and stomatal opening contribute only slightly to the enhancement of photosynthesis, particularly under the low light environments.
Signaling Mechanism of Leaf Positioning and Leaf Flattening
Without blue light, petioles were arched (Figure 1A, left). This suggests that the upper side of the petiole might elongate more than the lower side. When blue light was superimposed on red light, the epinastic growth of petioles was inhibited and caused the petioles to grow straight (Figure 1A, right). A similar differential growth between irradiated and shaded sides was previously reported in the coleoptile phototropism in monocotyledons (Iino and Briggs, 1984; Haga et al., 2005). Such differential growth is induced by a lateral translocation of auxin to the shaded side, and CPT1 is reported to function in this process (Friml et al., 2002; Haga et al., 2005). Moreover, the mutants defective in auxin sensitivity, such as msg1/nph4 and axr4, have strongly curled leaves (Hobbie and Estelle, 1995; Watahiki and Yamamoto, 1997), as has been found in the phenotype of the phot1 phot2 mutant (Sakai et al., 2001; Sakamoto and Briggs, 2002). The leaf curling of the msg1/nph4 mutant is attributed to the differential growth between the upper and lower sides (Stowe-Evans et al., 1998). It is likely that the leaf positioning and leaf flattening shown in this study are also achieved by the differential growth in both the petioles and leaves, which might be achieved via the alteration of auxin distribution. Further studies are needed to clarify the participation of auxin in these responses using transgenic plants in which auxin distribution can be visualized (Friml et al., 2002).
|
METHODS |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
|---|
Plant Materials and Growth Conditions
Arabidopsis thaliana wild-type and mutants plants were grown under white fluorescent lamps at 50 µmol m–2 s–1 for 7 d under a 14/10 h light–dark cycle. The plants were then transferred to red light (25 µmol m–2 s–1) with or without blue light (0.1 or 5 µmol m–2 s–1) under continuous light. All plants were grown at 24°C with a relative humidity of 55–75% in growth rooms. To determine growth, plants were grown under red light (25 µmol m–2 s–1) with or without blue light (0.1 µmol m–2 s–1). The T-DNA insertional mutant pool CS22830, of M. Sussman and R. Amasino, was obtained from the Arabidopsis Biological Research Center (The Ohio State University, Columbus, OH, USA). We used nph3-6 as a null mutant instead of the WS background nph3-202 mutant to compare growth on the Col background (Motchoulski and Liscum, 1999; Figure 7).
Isolation of Mutants Lacking Blue Light-Induced Leaf Positioning
We screened 34 000 EMS-mutagenized Arabidopsis seedlings of the M2 population and 30 000 T-DNA insertion seedlings by isolating the mutant lacking upward petiole growth under our experimental conditions. We obtained 32 mutants (23 lines of the EMS-mutagenized population and nine lines of the T-DNA insertional population) that showed horizontal petiole growth. Of these, 11 lines were fertile and heritable phenotypes in M3 generations. We found that one EMS mutant and one T-DNA mutant expressed wild-type levels of phot1 protein by immunoblotting using these mutants. The phot1 proteins in these two mutants exhibited autophosphorylation in response to blue light, and no mutation in the genomic PHOT1 of either mutant was found (data not shown). When the two mutants were crossed with each other, upward petiole growth was impaired in all of the obtained F1 seedlings (data not shown), suggesting that the two mutations are allelic to each other. After three backcrosses to the wild type (Col-0 and WS, respectively), these two mutants were used in all experiments.
Preparation of Protoplasts from Guard Cells and Mesophyll Cells
Protoplasts of guard and mesophyll cells from Arabidopsis were prepared enzymatically as reported by Ueno et al. (2005) with slight modifications. The amount of protein was determined as described previously (Bradford, 1976).
Expression of NPH3 Transcripts Determined by RT-PCR
Total RNAs were extracted from guard cell protoplasts, mesophyll cell protoplasts, leaves, stems, and roots of 4-week-old plants with ISOGEN (Nippon Gene, Tokyo, Japan). First-strand cDNAs were synthesized from 5 µg of each total RNA by SuperScript III reverse transcriptase using oligo(dT)12–18 primer (Invitrogen, Carlsbad, CA, USA). A 500 bp fragment of NPH3 cDNA was amplified with the primers 5'-GGTTGGAGTTGGAGGTGGAG-3’ and 5'-GATCGTCGGGTCAGGATCTC-3'. As an internal standard, a 350 bp fragment of ACT8 cDNA was used with the primers 5'-ACTTTACGCCAGTGGTCGTACAAC-3’ and 5'-AAGGACTTCTGGGCACCTGAATCT-3'. The PCR was obtained after 27 cycles for Figure 4A.
For amplification of the full-length NPH3 cDNA from the wild types (Col and WS) and from nph3-201 and nph3-202 mutants, total RNAs were prepared and first-strand cDNAs were synthesized as described above. For PCRs, two pairs of oligonucleotide primers were used: 5'-TTCCCTTGGTCCTTTCTTGCTTCC-3’ and 5'-CTATCACTTCATGAAATTGAGTTCCTCC-3’ (for NPH3), and 5'-CTCAAGAGGTTCTCAGCAGTA-3’ and 5'-TCACCTTCTTCATCCGCAGTT-3’ (for TUB2).
Thermal Asymmetric Interlaced (TAIL)-PCR
To identify the T-DNA insertion site of the nph3-202 mutant, we performed TAIL-PCR using genomic DNA from the mutant seedlings. The PCR and thermal cycler programs were performed according to the method of Liu et al. (1995) with a minor modification. For the gene-specific primers, 5'-CCTATAAATACGACGGATCG-3', 5'-ATAACGCTGCGGACATCTAC-3', and 5'-TGATCCATGTAGATTTCCCG-3’ were used. The primers were designed at the right border of the T-DNA region on the pD991 vector. For arbitrary degenerate primers, 5'-NTCGASTWTSGWGTT-3', 5'-NGTCGASWGANAWGAA-3', 5'-WGTGNAGWANCANAGA-3', 5'-TGWGNAGWANCASAGA-3', 5'-AGWGNAGWANCAWAGG-3', 5'-CAWCGICNGAIASGAA-3', 5'-TCSTICGNACITWGGA-3', and 5'-GTNCGASWCANAWGTT-3’ were used. The amplified genomic DNA fragments were obtained by nested PCR twice, and were cloned into a pCR4-TOPO vector (Invitrogen) and sequenced.
Construction of Plant Transformation Vector
To complement our nph3 mutants with the wild-type NPH3 gene, we constructed a gene transfer vector bearing the genomic NPH3 gene under the control of the native NPH3 promoter. The genomic NPH3 gene, including 5’ and 3’ noncoding sequences, was partially amplified by PCR from genomic DNA of the wild type (Col-0) using oligonucleotide primers 5'-CCGGGAGCTCTCTCGCTAGCATAACCATAAACCCC-3’ and 5'-TTGTTCGAATTGCATCCCTACGCG-3’ (for the first half of NPH3), and 5'-CGTCTTCTTAGAGCAGCAAACATGC-3’ and 5'-CGCGGATCCGAAATCTGCAGACAGATAAGGCGTG-3’ (for the second half of NPH3). These amplified DNA fragments were treated with SacI, or SacI and BamHI, respectively, and subcloned into pBluescript II KS (+) (Stratagene, La Jolla, CA, USA), respectively. The latter half of the NPH3 fragment was cloned into the gene transfer vector pCAMBIA1300 (Cambia, Canberra, Australia) with SacI and BamHI sites. Then, the first half of the NPH3 fragment was cloned into pCAMBIA1300 containing the latter half of the NPH3 fragment with the SacI site. The resulting vector was verified by DNA sequencing.
Transformation of Arabidopsis
The gene transfer vector was introduced into Agrobacterium tumefaciens (GV3101), and the Agrobacterium was transformed into the nph3-201 and nph3-202 mutants by an A. tumefaciens-mediated method (Clough and Bent, 1998). Transformed plants were selected on a half-strength MS plate containing 2% (w/v) sucrose and 30 µg ml–1 hygromycin. The complementation test was performed using independent transgenic lines from the T3 generation.
Transient Expression Assays by Particle Bombardment
The cDNAs encoding the full-length, NPH3-201 fragment, and coiled-coil-C fragment of NPH3 protein were amplified by RT-PCR using the total RNA from wild-type seedlings with oligonucleotide primers 5'-CCATGGGGGAATCTGAGAGCGAC-3’ and 5'-CCGGCCATGGCTGAAATTGAGTTCCTCCATCGTCTTG-3’ (for full length), 5'-CCATGGGGGAATCTGAGAGCGAC-3’ and 5'- CCGGCCATGGCCATCACTTCCATCTCGTTCTGAAGC-3’ (for NPH3-201), and 5'-CCGGCCATGGCCTTTCAGGAAGGATGGGCTGCAG-3’ and 5'- CCGGCCATGGCTGAAATTGAGTTCCTCCATCGTCTTG-3’ (for coiled-coil-C). The obtained cDNAs were cloned into the CaMV35S-sGFP(S65T)-NOS3’ vector with NcoI (Niwa et al., 1999). Plasmids expressing the coiled-coil and C-terminus fragments were constructed from the plasmid of coiled-coil-C by inverse PCR with oligonucleotide primers 5'-GCCATGGTGAGCAAGGGC-3’ and 5'-AGAAGATGGCGTGTTCTTCACTTTCC-3’ (for coiled-coil), and 5'-ACGCCATCTTCTTCGGCTTGGACC-3’ and 5'-CCATCCTTCCTGAAAGGCCATGG-3’ (for C-terminus). After the inverse PCR, reaction mixtures were treated with DpnI for the degradation of template DNA and then with T4 polynucleotide kinase for phosphorylation of the 5’ ends. The phosphorylated linear DNAs were self-ligated. Plasmid DNAs were prepared for the particle bombardment and transfected as described previously (Emi et al., 2005). The transfected Vicia leaves were kept in darkness for 6–10 h at room temperature. Epidermal peels were obtained from the leaves, and epidermal cells and stomata were examined by a confocal laser-scanning microscope (Digital Eclipse C1; Nikon, Tokyo, Japan).
Determination of Phototropin-Mediated Physiological Responses
Growth enhancement, chloroplast distribution, and stomatal apertures were measured according to a previous report (Takemiya et al., 2005).
Light Source
White light was produced by fluorescent lamps (FL 40S N-SDL; National, Tokyo, Japan), and both red and blue light were produced by light-emitting photodiodes (LED-R, maximum intensity at 660 nm; and Stick-B-32, maximum intensity at 470 nm; Eyela, Tokyo, Japan). Photon flux densities were determined with a quantum meter (LI-250; Li-Cor, Lincoln, NE, USA) equipped with a light sensor (LI-190 SA; Li-Cor).
|
Acknowledgements |
|---|
We thank M. Wada (National Institute for Basic Biology, Okazaki, Japan) for providing seeds of the nph3-6 mutant. This work was supported by the Ministry of Education, Science, Sports, and Culture of Japan (grant Nos 16207003, 17084005 to K.S. and 14704003 to T.K.).
|
Notes |
|---|
2 Present address: Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa, Nagoya, 464-8602, Japan.
-
Ahmad M, Cashmore AR. HY4 gene of A. thaliana encodes a protein with characteristics of a blue-light photoreceptor. Nature (1993) 366:162–166.[CrossRef][Medline]
Ballaré CL, Scopel AL. Phytochrome signaling in plant canopies: testing its population-level implications with photoreceptor mutants of Arabidopsis. Funct. Ecol. (1997) 11:441–450.[CrossRef]
Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Anal. Biochem. (1976) 72:248–254.[CrossRef][ISI][Medline]
Briggs WR, Christie JM. Phototropins 1 and 2: versatile plant blue-light receptors. Trends Plant Sci. (2002) 7:204–210.[CrossRef][ISI][Medline]
Cashmore AR, Jarillo JA, Wu YJ, Liu D. Cryptochromes: blue light receptors for plants and animals. Science (1999) 284:760–765.
Clough SJ, Bent AF. Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. (1998) 16:735–743.[CrossRef][ISI][Medline]
Doi M, Shigenaga A, Emi T, Kinoshita T, Shimazaki K. A transgene encoding a blue-light receptor, phot1, restores blue-light responses in the Arabidopsis phot1 phot2 double mutant. J. Exp. Bot. (2004) 55:517–523.
Emi T, Kinoshita T, Sakamoto K, Mineyuki Y, Shimazaki K. Isolation of a protein interacting with Vfphot1a in guard cells of Vicia faba. Plant Physiol. (2005) 138:1615–1626.
Fankhauser C, Casal JJ. Phenotypic characterization of a photomorphogenic mutant. Plant J. (2004) 39:747–760.[CrossRef][ISI][Medline]
Folta KM, Kaufman LS. Phototropin1 is required for high-fluence blue-light-mediated mRNA destabilization. Plant Mol. Biol. (2003) 51:609–618.[CrossRef][ISI][Medline]
Folta KM, Spalding EP. Unexpected roles for cryptochrome 2 and phototropin revealed by high-resolution hypocotyl growth analysis. Plant J. (2001) 26:471–478.[CrossRef][ISI][Medline]
Friml J, Wi
niewska J, Benková E, Mundgen K, Palme K. Lateral relocation of auxin efflux regulator PIN3 mediates tropism in Arabidopsis. Nature (2002) 415:806–809.[Medline]
Haga K, Takano M, Neumann R, Iino M. The rice COLEOPTILE PHOTOTROPISM1 gene encoding an ortholog of Arabidopsis NPH3 is required for phototropism of coleoptiles and lateral translocation of auxin. Plant Cell (2005) 17:103–115.
Hobbie L, Estelle M. The axr4 auxin-resistant mutants of Arabidopsis thaliana define a gene important for root gravitropism and lateral root initiation. Plant J. (1995) 7:211–220.[CrossRef][ISI][Medline]
Huala E, Oeller PW, Liscum E, Han I-S, Larsen E, Briggs WR. Arabidopsis NPH1: a protein kinase with a putative redox-sensing domain. Science (1997) 278:2120–2123.
Iino M, Briggs WR. Growth distribution during first positive phototropic curvature of maize coleoptiles. Plant Cell Environ. (1984) 7:97–104.[CrossRef]
Imaizumi T, Tran HG, Swartz TE, Briggs WR, Kay SA. FKF1 is essential for photoperiodic-specific light signalling in Arabidopsis. Nature (2002) 426:302–306.
Inada S, Ohgishi M, Mayama T, Okada K, Sakai T. RPT2 is a signal transducer involved in phototropic response and stomatal opening by association with phototropin1 in Arabidopsis thaliana. Plant Cell (2004) 16:887–896.
Inoue S, Kinoshita T, Shimazaki K. Possible involvement of phototropins in leaf movement of kidney bean in response to blue light. Plant Physiol. (2005) 138:1994–2004.
Jarillo JA, Gabrys H, Capel J, Alonso JM, Ecker JR, Cashmore AR. Phototropin-related NPL1 controls chloroplast relocation induced by blue light. Nature (2001) 410:952–954.[CrossRef][Medline]
Kagawa T, Sakai T, Suetsugu N, Oikawa K, Ishiguro S, Kato T, Tabata S, Okada K, Wada M. Arabidopsis NPL1: a phototropin homolog controlling the chloroplast high-light avoidance response. Science (2001) 291:2138–2141.
Kasahara M, Kagawa T, Oikawa K, Suetsugu N, Miyao M, Wada M. Chloroplast avoidance movement reduces photodamage in plants. Nature (2002) 420:829–832.[CrossRef][Medline]
Kinoshita T, Doi M, Suetsugu N, Kagawa T, Wada M, Shimazaki K. phot1 and phot2 mediate blue light regulation of stomatal opening. Nature (2001) 414:656–660.[CrossRef][Medline]
Koller D. Plants in search of sunlight. Adv. Bot. Res. (2000) 33:35–131.[CrossRef]
Koller D, Levitan I. Diurnal phototropism in leaves of Lavatera cretica L. under conditions of simulated solar-tracking. J. Exp. Bot. (1989) 40:1059–1064.
Koller D, Levitan I, Briggs WR. The vectorial photo-excitation in solar-tracking leaves of Lavatera cretica (Malvaceae). Photochem. Photobiol. (1985) 42:717–723.[CrossRef]
Lariguet P, et al. PHYTOCHROME KINASE SUBSTRATE 1 is a phototropin 1 binding protein required for phototropism. Proc. Natl Acad. Sci. USA (2006) 103:10134–10139.
Lin C. Blue light receptors and signal transduction. Plant Cell (2002) 14(suppl.):S207–S225.
Liscum E. Phototropism: mechanisms and outcomes. In: The Arabidopsis Book—Somerville CR, Meyerowitz EM, eds. (2002) Rockville, MD: American Society of Plant Biologists. doi/10.1199/tab.0042, http://www.aspb.org/publications/arabidopsis/.
Liscum E, Briggs WR. Mutations in the NPH1 locus of Arabidopsis disrupt the perception of phototropic stimuli. Plant Cell (1995) 7:473–485.[Abstract]
Liu Y-G, Mitsukawa N, Whitter RF. Efficient isolation and mapping of Arabidopsis thaliana T-DNA insert junctions by thermal asymmetric interlaced PCR. Plant J. (1995) 8:457–463.[CrossRef][ISI][Medline]
Mano E, Horiguchi G, Tsukaya H. Gravitropism in leaves of Arabidopsis thaliana (L.) Heynh. Plant Cell Physiol. (2006) 47:217–223.
Motchoulski A, Liscum E. Arabidopsis NPH3: a NPH1 photoreceptor-interacting protein essential for phototropism. Science (1999) 286:961–964.
Mullen JL, Weinig C, Hangarter RP. Shade avoidance and the regulation of leaf inclination in Arabidopsis. Plant Cell Environ. (2006) 29:1099–1106.[CrossRef][Medline]
Nemhauser J, Chory J. Photomorphogenesis. In: The Arabidopsis Book—Somerville CR, Meyerowitz EM, eds. (2002) Rockville, MD: American Society of Plant Biologists. doi/10.1199/tab.0054, http://www.aspb.org/publications/arabidopsis/.
Niklas KJ, Owens TG. Physiological and morphological modifications of Plantago major (Plantginaceae) in response to light conditions. Am. J. Bot. (1989) 76:370–382.[CrossRef][ISI]
Niwa Y, Hirano T, Yoshimoto K, Shimizu M, Kobayashi H. Non-invasive quantitative detection and applications of non-toxic, S65T-type green fluorescent protein in living plants. Plant J. (1999) 18:455–463.[CrossRef][ISI][Medline]
Sakai T, Kagawa T, Kasahara M, Swartz TE, Christie JM, Briggs WR, Wada M, Okada K. Arabidopsis nph1 and npl1: blue light receptors that mediate both phototropism and chloroplast relocation. Proc. Natl Acad. Sci. USA (2001) 98:6969–6974.
Sakai T, Wada T, Ishiguro S, Okada K. RPT2: a signal transducer of the phototropic response in Arabidopsis. Plant Cell (2000) 12:225–236.
Sakamoto K, Briggs WR. Cellular and subcellular localization of phototropin 1. Plant Cell (2002) 14:1723–1735.
Stowe-Evans EL, Harper RM, Motchoulski AV, Liscum E. NPH4, a conditional modulator of auxin-dependent differential growth responses in Arabidopsis. Plant Physiol. (1998) 118:1265–1275.
Takemiya A, Inoue S, Doi M, Kinoshita T, Shimazaki K. Phototropins promote plant growth in response to blue light in low light environments. Plant Cell (2005) 17:1120–1127.
Ueno K, Kinoshita T, Inoue S, Emi T, Shimazaki K. Biochemical characterization of plasma membrane H+-ATPase activation in guard cell protoplasts of Arabidopsis thaliana in response to blue light. Plant Cell Physiol. (2005) 46:955–963.
Vandenbussche F, Verbelen JP, Van Der Straeten D. Of light and length: regulation of hypocotyl growth in Arabidopsis. BioEssays (2005) 27:275–284.[CrossRef][ISI][Medline]
Wang H, Deng XW. Phytochrome signaling mechanism. In: The Arabidopsis Book—Somerville CR, Meyerowitz EM, eds. (2002) Rockville, MD: American Society of Plant Biologists. doi/10.1199/tab.0074, http://www.aspb.org/publications/arabidopsis/.
Watahiki MK, Yamamoto KT. The massugu1 mutation of Arabidopsis identified with failure of auxin-induced growth curvature of hypocotyl confers auxin insensitivity to hypocotyl and leaf. Plant Physiol. (1997) 115:419–426.[Abstract]
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  S.-i. Inoue, T. Kinoshita, M. Matsumoto, K. I. Nakayama, M. Doi, and K.-i. Shimazaki Blue light-induced autophosphorylation of phototropin is a primary step for signaling PNAS, April 8, 2008; 105(14): 5626 - 5631. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||