mol. plant 2008-inoue-15-26

Molecular Plant • Volume 1 • Number 1 • Pages 15–26 • January 2008

Leaf Positioning of Arabidopsis in Response toBlue Light

Shin-ichiro Inouea, Toshinori Kinoshitaa,2, Atsushi Takemiyaa, Michio Doib and Ken-ichiro Shimazakia,1

a Department of Biology, Faculty of Science, Kyushu University, Ropponmatsu, Fukuoka, 810-8560 Japanb Research and Development Center for Higher Education, Kyushu University, Ropponmatsu, Fukuoka, 810-8560 Japan

ABSTRACT Appropriate leaf positioning is essential for optimizing photosynthesis and plant growth. However, it has not

beenelucidatedhowgreen leaves reachandmaintain their position for capturing light.Weshowhere the regulationof leaf

positioning under blue light stimuli. When 1-week-old Arabidopsis seedlings grown under white light were transferred to

red light (25 mmol m22 s21) 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 mmol m22 s21) was superimposed on red light, the

new petioles grew obliquely upward and the leaves were flat and horizontal. The leaf positioning required both photo-

tropin1 (phot1) andnonphototropic hypocotyl 3 (NPH3), and resulted in enhancedplant growth. In annph3mutant, neither

optimal leaf positioning nor leaf flattening by blue lightwas found, and blue light-induced growthenhancementwas dras-

tically reduced.When blue lightwas increased from0.1 to 5mmolm22 s21, normal leaf positioning and leaf flatteningwere

induced in both phot1 and nph3 mutants, suggesting that phot2 signaling became functional and that the signaling was

independentofphot1andNPH3 in these responses.Whenplantswere irradiatedwithblue light (0.1mmolm22 s21) fromthe

side and red light from above, the new leaves became oriented toward the source of blue light.Whenwe 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 plasmamembrane localization

via the coiled-coil domain and theC-terminal region.We conclude thatNPH3-mediatedphototropin signalingoptimizes the

efficiency of light perception by inducing both optimal leaf positioning and leaf flattening, and enhances plant growth.

INTRODUCTION

Plants respond appropriately to ever-changing environments

by morphogenesis, movement, changes in cellular compo-

nents, and metabolic activity, thereby optimizing growth in

natural environments. Plants respond by sensing changes in

light, gravity, temperature, salt, and water status through in-

dividual receptors. Light is the most important factor influenc-

ing 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 developmen-

tal and movement responses, including phototropic bending,

cotyledon opening, photoperiodic flowering, leaf flattening,

de-etiolation, stomatal opening, chloroplast movements,

anthocyanin accumulation, gene expression, and the inhibi-

tion of hypocotyl elongation (Cashmore et al., 1999; Briggs

and Christie, 2002; Lin, 2002; Wang and Deng, 2002). In Arabi-

dopsis plants, three classes of major blue light receptors—cryp-

tochromes, 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 re-

ceptor using an Arabidopsis mutant that did not show hypo-

cotyl 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 im-

paired 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

1 To whom correspondence should be addressed. E-mail [email protected].

kyushu-u.ac.jp, fax 81-92-726-4758.

2 Present address: Division of Biological Science, Graduate School of Science,

Nagoya University, Chikusa, Nagoya, 464-8602, Japan.

ª The Author 2007. Published by Oxford University Press on behalf of CSPP

and IPPE, SIBS, CAS.

doi: 10.1093/mp/ssm001, Advance Access publication 7 June 2007

by guest on June 8, 2012http://m

plant.oxfordjournals.org/D

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(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 pre-

vent strong light from damaging the photosynthetic machin-

ery (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 photosyn-

thesis, and a dramatic plant growth enhancement mediated by

phototropin is demonstrated under a low intensity of photo-

synthetically active radiation (PAR) (Takemiya et al., 2005).

Extensive studies on phototropism were done using etio-

lated hypocotyls and coleoptiles as model systems, and in

many cases blue light was provided from the lateral side be-

cause it is easy to measure and analyze the responses (Fank-

hauser and Casal, 2004; Vandenbussche et al., 2005). These

investigations have provided detailed information on photo-

tropic 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 devel-

opment. 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 po-

sition 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

Blue Light-Dependent Leaf Positioning Increases

Light Capture

We grew Arabidopsis seedlings under white light at 50 lmol

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 lmol m�2 s�1 with or

without low-intensity blue light (0.1 lmol 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 (Fig-

ure 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 ori-

ented 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 (h), 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 inter-

ception 2-fold in each first leaf when the blue light was pro-

vided together with red light from the top (Figure 1C and D).

We next illuminated the plants with blue light (0.1 lmol m�2

s�1) from the side but red light from the top as before. New

petioles emergedandthe newleafletsbecameorientedtoward

theblue light source,but theface of the leafwasnot completely

perpendicular to that source (Figure 1E, solid arrowheads). The

surfaceofapairofopencotyledonsbecamepartiallyorientedto

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 percep-

tion 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 physiologi-

cal responses at the same time (Briggs and Christie, 2002; Take-

miya 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 posi-

tioning 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 dou-

ble 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 indi-

cate 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

16 | Inoue et al. d Blue Light-Mediated Leaf Positioning



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growth. We obtained two mutant lines: an ethylmethane sul-

fonate (EMS)-mutagenized plant and a T-DNA insertional

plant, both of which showed impairment in the upward pet-

iole 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 asym-

metric 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 (Fig-

ure 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 substitu-

tion 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 mu-

tation by the wild-type genomic NPH3 gene. A 5400 bp geno-

mic 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 po-

sitioning with upward petioles (Figure 3D). The results demon-

strate 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, respec-

tively (Figure 3).

Expression of NPH3

We investigated the expression ofNPH3mRNA 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 pro-

tein (GFP) in epidermal cells and guard cells ofVicia fababy par-

ticle bombardment. The fluorescence from full-length NPH3

was found on the periphery of both epidermal and guard cells,

Figure 1. Leaf Positioning in Response to a VeryLow Intensity of Blue Light.

Wild-type (Col-0) plants of Arabidopsis weregrown under white light (50 lmol m�2 s�1) for7 d and then transferred to red light (25 lmolm�2 s�1) with or without blue light (0.1 lmolm�2 s�1). The plants were further grown for 5 d.The supplemental blue light was applied fromabove (A–D) or from the side (E). White solidarrowheads show the first true leaves. Whiteopen arrowheads show cotyledons. White arrowsshow the direction of blue light.(A) Side view of plants after growth for 5 d withor without blue light. The white bar represents1 cm.(B) Angles (h) of petioles from the horizontal line.Values presented are means of 25 seedlings withstandard errors.(C) Pictures taken from above. The black bar rep-resents 1 cm.(D) Area of light perception in the first leaf. Areasof projections by the first leaves were measuredby taking pictures from above. Bars representmeans 6 SE (n = 32).(E) Side view of plants after growth for 5 d. Sideview is perpendicular to the applied blue light.Right view is from the same direction as the bluelight source.

Inoue et al. d Blue Light-Mediated Leaf Positioning | 17



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suggestingplasmamembrane localizationofNPH3asdescribed

previously (Motchoulski and Liscum, 1999; Lariguet et al., 2006;

Figure 4B, full length). We then investigated the localization in

moredetailusingguardcellsbecausethetransientexpressionof

NPH3 is mucheasier in themthan inepidermal cells. The fluores-

cenceofmutantNPH3-201proteinfromnph3-201wasobserved

as many particles in cytosolic compartments (Figure 4B, NPH3-

201). Since the mutant NPH3-201 protein may lack a C-terminal

regiondownstreamfromthecoiled-coildomain (Figure3B), it is

possible that this region is required for the membrane localiza-

tion of NPH3. To test this, we expressed the NPH3 C-terminal

fragment containing the coiled-coil domain (coiled-coil-C)

fusedwithGFP.Asexpected, thefluorescentsignalofthis 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)andaC-terminal region(C-terminus)andexpressed

these as above. The GFP fluorescence of the coiled-coil domain

wasdetectedmainly in theplasmamembraneandslightly inthe

cytoplasm(Figure4B,Coiled-coil). ThefluorescenceoftheC-ter-

minus 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 con-

served 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

Figure 2. Leaf Positioning Mediated by phot1.

Wild-type(gl1andWS),phot1-5,phot2-1,phot1-5pho2-1, andhy4-3cry2-1 plants were grown and transferred as described in Figure 1.(A) Plants grown under red light at 25 lmol m�2 s�1.(B) Plants grown under red light with blue light at 0.1 lmol m�2 s�1.(C) Angles of petioles in these plants. The measurements were doneas in Figure 1. Values are the means of 25–38 seedlings with stan-dard errors. White bars represent 1 cm.

Figure 3. Involvement of NPH3 in Leaf Positioning.

(A) Isolation of mutants impaired in upward petiole growth underthe low blue light condition. The picture shows mutant plantsgrown under red light with low blue light. The white bar represents1 cm.(B)Determination of the mutated gene in the isolated mutants. Thegenomic structure of NPH3 on chromosome 5 is shown. Black boxesand bold lines represent exons and introns, respectively. An nph3-201 mutant has a C-to-T nucleotide substitution in the last exon.This nucleotide change causes the substitution of Gln681 by thestop codon. T-DNA insertion in nph3-202 was identified in the fifthexon.(C) Expression of NPH3 and TUB2 (b-tubulin) mRNAs analyzed byRT-PCR in 2-week-old seedlings of wild-type (Col and WS) plantsand of two nph3 mutants (nph3-201 and nph3-202).(D) Functionalcomplementationofnph3-201andnph3-202mutantswith wild-type genomic NPH3 genes. Plants of nph3-201, nph3-201transformed with wild-type genomic NPH3 (201-G), nph3-202, andnph3-202 transformed with wild-type genomic NPH3 (202-G) weregrown as in Figure 1. The white bar represents 1 cm.




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supplemented blue light was increased to 5 lmol m�2 s�1 from

0.1 lmol 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 lmol m�2 s�1

red light with 0.1 lmol 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 in-

creased to 5 lmol 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

Figure 5. Rescue of Leaf Positioning Under a Relatively High Inten-sity of Blue Light in phot1-5 and nph3 Mutants.

Wild-type (gl1, Col-0, and WS) plants and phot1-5, phot2-1, phot1-5phot2-1, nph3-201, and nph3-202 plants were grown under whitelight at 50 lmol m�2 s�1 from fluorescent lamps for 7 d and thentransferred under red light (25 lmol m�2 s�1) with blue light andallowed to grow for an additional 5 d for the determination of thepetiole angles.(A) Pictures indicate the leaf positioning in the mutant plants under5 lmol m�2 s�1 of blue light.(B) Angles of petioles were measured under 0.1 or 5 lmol m�2 s�1 ofbluelightasinFigure1.Valuesaremeansof21–28seedlingswithstan-dard errors.

Figure 4. Expression of NPH3 mRNAs and Subcellular Localizationof NPH3 Protein.

(A) Expression of NPH3 mRNAs in guard cell protoplasts (GCPs), me-sophyll cellprotoplasts (MCPs), leaves, stems,androots from4-week-old plants analyzed by RT-PCR. The purities of GCPs and MCPs were98 and 99%, respectively, on a cell number basis. ACT8 was used asan internal standard for cDNA amounts. Two separate experimentsgave similar results.(B) Transient expression of NPH3–GFP proteins in Vicia epidermalcells and guard cells. The primary structure of NPH3 protein andstructures of fusion proteins are illustrated. Four conserved domainsin the NPH3/RPT2 family are shown in light gray open blocks as de-scribed in Liscum (2002). The BTB (broad complex, tramtrack, bric abrac)/POZ (pox virus and zinc finger) domain and the coiled-coil do-main are shown in the dark gray block and black block, respectively.The full length and fragments of NPH3 proteins were fused in-frameto the N-terminal end of sGFP and were expressed transiently by par-ticle bombardment under the control of the CaMV 35S promoter.Full length, full-length NPH3 protein fused to GFP; NPH3-201,NPH3 fragment of the N-terminus fused to GFP on Met680;Coiled-coil-C, NPH3 fragment of Phe645 to the C-terminus fusedto GFP; Coiled-coil, NPH3 fragment from Phe645 to Ser696 fusedto GFP; C-terminus, NPH3 fragment from Thr693 to the C-terminusfused to GFP; sGFP, GFP protein. Epidermal cells and guard cellsexpressing these proteins were inspected by GFP fluorescence usinga confocal laser microscope. All pictures are cross-sectional.




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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 flat-

tening in response to very weak blue light (Figures 3A and 6A),

but does not mediate chloroplast movement or stomatal open-

ing (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 flatten-

ing 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 lmol m�2 s�1 blue light to the red light, but no actual

growth enhancement was found in the phot1-5 mutant. Inter-

estingly, 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 sto-

matal 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.

Figure 7. Growth Enhancement, Chloroplast Accumulation, andStomatal Opening in Response to Low Intensity of Blue Light.

Wild-type (gl1), phot1-5, nph3-201, and nph3-6 plants were grownfor 5 weeks under red light (25 lmol m�2 s�1) with or without bluelight (0.1 lmol m�2 s�1). The growth was determined as freshweight of green tissues.(A) Growth enhancement by blue light in wild-type and mutantplants. Plants grown under red light (left) and red light with bluelight (right).(B) Fresh weights of green tissues of plants. Bars represent means 6SE (n = 25). Asterisks show significant statistical differences by t-test(P ,0.05) in fresh weights.(C) Distribution of chloroplasts in mesophyll cells of wild-type andmutant leaves under our growth conditions.(D) Stomatal aperture in leaves of the wild type and mutants underour growth conditions. Apertures are expressed as the ratio ofwidth to length of the guard cell pair, as described in Takemiyaet al. (2005). Bars represent means 6 SE (n = 25).

Figure 6. Leaf Flattening in Wild Type and Various Mutants in Re-sponse to Low and High Intensities of Blue Light.

Plants of the wild types (gl1, Col-0, and WS), phot1-5, phot2-1,phot1-5 phot2-1, nph3-201, and nph3-202 were initially grown un-der white light at 50 lmol m�2 s�1 from fluorescent lamps for 7 d.The plants were then transferred under red light (25 lmol m�2 s�1)with blue light of two different intensities and allowed to grow foran additional 10 d to determine the leaf flattening.(A) Leaf flattening of the wild types and mutants with blue light at0.1 lmol m�2 s�1.(B) Leaf flattening of wild-types and mutants with blue light at5 lmol m�2 s�1. White bars represent 1 cm.




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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 to-

ward the source of blue light (Figure 8A, 0 h). Such leaf orien-

tation in response to blue light was not found in the mutants

of phot1-5or nph3-201 (data not shown). Then, we transferred

these plants to both red (25 lmol m�2 s�1) and blue (0.1 lmol

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; hL), and began a rel-

atively slower phase after about 2 h (Figure 8A and B, 2 h).

Then, the leaf surface gradually approached the maximum an-

gle within 8 h (Figure 8B, leaf angle in left graph), and main-

tained this position thereafter with a very slight change

Figure 8. Changes in Leaf Position in Re-sponse to Blue Light.

Wild-type (gl1) plants were grown underwhite light (50 lmol m�2 s�1) for 7 dand then transferred to red light (25lmol m�2 s�1) from above with blue light(0.1 lmol m�2 s�1) from the plant side, andwere grown for 5 d, as indicated in Figure1E. The plants were then transferred againand irradiated with blue light (0.1 lmolm�2 s�1) from above under the red light,and growth was allowed for an additional5 d.(A) Side view of the plants after the secondtransfer. Pictures were taken at the indi-cated times from the perpendicular tothe direction of the first applied blue light,which had been derived from the left (up-per panels), and taken from the same di-rection of the blue light (lower panels).White solid arrowheads show the firsttrue leaves. White open arrowheads showcotyledons. The black arrow indicates thedirection of the first blue light treatment.The white arrow shows the direction ofthe second blue light treatment.(B) Angle of the first leaf from the vertical(hL) and that of the first leaf petiole fromthe vertical (hP). Typical changes in theseangles in response to blue light are shown.The left illustration indicates the changeof angles during 8 h with high time reso-lution. The right illustration shows thechange of angles during 5 d. Gray ovalsrepresent the first leaves. White ovalsshow the cotyledons.(C) Rotation of the first leaves which oc-curred after the initial leaf orientation.Pictures were taken at the indicated timesfrom above. White solid arrowheads showthe first true leaves. Black arrows indi-cate the direction of blue light appliedpreviously.(D) Petiole rotation. Typical changes inthe angles of petioles (hR) in response toblue light are shown. Gray ovals repre-sent the first leaves. White ovals showthe cotyledons.




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(Figure 8A and B, right graph). The petiole angle in the pro-

jected 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; hP). The rate of the rapid leaflet orienta-

tion 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 so-

lar-tracking response inArabidopsis, and is mediated by phot1.

During the leaf repositioning responses, the petiole was

arch-shaped from 4 to 24 h, a conformation that facilitated ori-

enting 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 rel-

atively 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

Blue Light-Mediated Leaf Positioning Promotes

Light-Capturing Efficiency

Plants control leaf position in response to environmental stim-

uli, 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 opti-

mal 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 phototro-

pins (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 uti-

lize 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 di-

rection after 24 h. These results suggest that the plants pref-

erentially 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 in-

creased 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 al-

though the responses are mediated by the same photorecep-

tors, the mechanisms between leaf positioning and leaf

movement may differ, since the complete Arabidopsis leaf po-

sitioning 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 sug-

gested 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 grav-

itropism (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 Ara-

bidopsis and Oryza, respectively (Motchoulski and Liscum,

1999; Haga et al., 2005). Another example of NPH3 involve-

ment is phot1-mediated destabilization of Lhcb and rbcL tran-

scripts (Folta and Kaufman, 2003). In the present study, we




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found for the first time that NPH3 mediated both leaf position-

ing and leaf flattening in the phot1-dependent pathway (Fig-

ures 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 mem-

brane may facilitate phot1–NPH3 complex formation and sig-

ing (Motchoulski and Liscum, 1999; Lariguet et al., 2006;

Figure 3A).

NPH3 is suggested to function as a common signal com-

ponent 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 (Fig-

ures 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 compo-

nent 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 mor-

phological changes in photosynthetic tissues under various

light environments (Niklas and Owens, 1989; Ballare and

Scopel, 1997).

In a previous work we demonstrated that phot1 dramati-

cally enhances plant growth in response to a very low intensity

of blue light, and that the enhancement is achieved by inte-

grating phot1-mediated responses, including those of chloro-

plast 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 en-

hancement, 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 open-

ing. Taking advantage of this property of the nph3 mutant, we

showed that this mutant slightly enhanced plant growth un-

der 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 move-

ment and stomatal opening contribute only slightly to the en-

hancement 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 sim-

ilar 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 phe-

notype of the phot1 phot2 mutant (Sakai et al., 2001; Saka-

moto 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 partici-

pation of auxin in these responses using transgenic plants in

which auxin distribution can be visualized (Friml et al., 2002).

METHODS

Plant Materials and Growth Conditions

Arabidopsis thaliana wild-type and mutants plants were

grown under white fluorescent lamps at 50 lmol m�2 s�1

for 7 d under a 14/10 h light–dark cycle. The plants were then

transferred to red light (25 lmol m�2 s�1) with or without blue

light (0.1 or 5 lmol 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 lmol m�2 s�1) with or without blue light

(0.1 lmol 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 mu-

tant instead of the WS background nph3-202 mutant to com-

pare 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




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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 pheno-

types 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, meso-

phyll cell protoplasts, leaves, stems, and roots of 4-week-old

plants with ISOGEN (Nippon Gene, Tokyo, Japan). First-strand

cDNAs were synthesized from 5 lg of each total RNA by Super-

Script III reverse transcriptase using oligo(dT)12–18 primer (Invi-

trogen, Carlsbad, CA, USA). A 500 bp fragment of NPH3 cDNA

was amplified with the primers 5#-GGTTGGAGTTGGAGGTG-

GAG-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#-TTCCCTTGGTCCTTTCT-

TGCTTCC-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#-CCTA-

TAAATACGACGGATCG-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#-NTC-

GASTWTSGWGTT-3#, 5#-NGTCGASWGANAWGAA-3#, 5#-WGT-

GNAGWANCANAGA-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 geno-

micNPH3 gene under the control of the nativeNPH3 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 sub-

cloned 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 con-

taining 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 trans-

formed 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 lg 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#-CCGGCCATGGCCTTTCAGGAAGGATGGGCT-

GCAG-3’ and 5#- CCGGCCATGGCTGAAATTGAGTTCCTCCATC-

GTCTTG-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




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coiled-coil-C by inverse PCR with oligonucleotide primers 5#-

GCCATGGTGAGCAAGGGC-3’ and 5#-AGAAGATGGCGTGTTCTT-

CACTTTCC-3’ (for coiled-coil), and 5#-ACGCCATCTTCTTCGGC-

TTGGACC-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 in-

tensity at 660 nm; and Stick-B-32, maximum intensity at 470

nm; Eyela, Tokyo, Japan). Photon flux densities were deter-

mined with a quantum meter (LI-250; Li-Cor, Lincoln, NE,

USA) equipped with a light sensor (LI-190 SA; Li-Cor).

ACKNOWLEDGMENTS

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 Cul-

ture of Japan (grant Nos 16207003, 17084005 to K.S. and

14704003 to T.K.).

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