Carbon Starved Anther Encodes a MYB Domain ProteinThat Regulates Sugar Partitioning Required for RicePollen Development W OA

Hui Zhang,a,b,1 Wanqi Liang,a,1 Xijia Yang,a Xue Luo,a Ning Jiang,c Hong Ma,d,e and Dabing Zhanga,b,2

a School of Life Science and Biotechnology, Shanghai Jiao Tong University, Shanghai 200240, Chinab Bio-X Research Center, Key Laboratory of Genetics and Development and Neuropsychiatric Diseases, Ministry of Education,

Shanghai Jiao Tong University, Shanghai 200240, Chinac Department of Horticulture, Michigan State University, East Lansing, Michigan 48824d State Key Laboratory of Genetic Engineering, Institute of Plant Biology, Center for Evolutionary Biology, School of Life

Sciences, Fudan University, Shanghai 200433, Chinae Department of Biology, Huck Institutes of the Life Sciences, Pennsylvania State University, University Park, Pennsylvania

16082

In flowering plants, sink tissues rely on transport of carbohydrates from photosynthetic tissues (sources) for nutrition and

energy. However, how sugar partitioning in plants is regulated at the molecular level during development remains unknown.

We have isolated and characterized a rice (Oryza sativa) mutant, carbon starved anther (csa), that showed increased sugar

contents in leaves and stems and reduced levels of sugars and starch in floral organs. In particular, the csa mutant had

reduced levels of carbohydrates in later anthers and was male sterile. The csa mutant had reduced accumulation of 14C-

labeled sugars in anther sink tissue. CSA was isolated by map-based cloning and was shown to encode an R2R3 MYB

transcription factor that was expressed preferentially in the anther tapetal cells and in the sugar-transporting vascular

tissues. In addition, the expression of MST8, encoding a monosaccharide transporter, was greatly reduced in csa anthers.

Furthermore, CSA was found to be associated in vivo and in vitro with the promoter of MST8. Our findings suggest that CSA

is a key transcriptional regulator for sugar partitioning in rice during male reproductive development. This study also

establishes a molecular model system for further elucidation of the genetic control of carbon partitioning in plants.

INTRODUCTION

Plants are highly specialized autotrophic organisms with distinct

tasks for various organs, such as photosynthesis and production

of sugars and other organic nutrients in leaves and uptake of

water and mineral nutrients in roots. To modulate the develop-

ment and nutrient exchange between the organs, plants have

evolved a vascular system composed of the xylem and the

phloem. The xylem is responsible for transporting water and

minerals from the root system to the shoot, and the phloem is

responsible for transporting organic nutrients from source tis-

sues, such as leaves, to sink tissues, such as roots, developing

organs from the shoot apex, and reproductive organs. Photo-

synthetic sugars are key substances in primarymetabolism; they

not only function as the major energy source and provide the

building blocks for macromolecules but also play crucial roles as

signaling molecules (Rolland et al., 2006). Plant cells have the

ability to take up sugars as carbon skeletons for production of

cellular components (i.e., cell wall) and othermetabolites, often in

response to plant hormones and external stresses (Lalonde

et al., 2004; Rolland et al., 2006).

Whereas glucose is the most important form of carbon for

energy and the form transported in animals, the disaccharide

sucrose is the main form of carbon for long-distance transport in

plants (Lemoine, 2000; Lalonde et al., 2004). Carbon partitioning

in plants between the source tissues and the various competing

sink tissues is a dynamic process that includes two key compo-

nents: the loading of photosynthetic assimilates from the source

into the phloem tissue and their unloading from the phloem into

the sink tissues (Lemoine, 2000). Several genes, such asSucrose

Transporters (SUTs), TIEDYED, andH+-ATPase, encoding trans-

membrane proteins have been shown to be important for phloem

loading of sucrose. Mutations in these genes cause excess

carbon accumulation in leaves and reduced or delayed growth

(Gottwald et al., 2000; Rolland et al., 2006; Buttner, 2007; Kocal

et al., 2008; Wang et al., 2008a; Slewinski et al., 2009). The

phloem unloading pathway is required for sink organs, such as

developing anthers, in which sucrose moves from phloem cells

to sink cells via plasmodesmata. Alternatively, sucrose can be

cleaved by cell wall invertases, forming glucose and fructose,

which can be taken up by sink tissues via monosaccharide

1 These authors contributed equally to this work.2 Address correspondence to zhangdb@sjtu.edu.cn.The author responsible for distribution of materials integral to thefindings presented in this article in accordance with the policy describedin the Instructions for Authors (www.plantcell.org) is: Dabing Zhang(zhangdb@sjtu.edu.cn).WOnline version contains Web-only data.OAOpen Access articles can be viewed online without a subscription.www.plantcell.org/cgi/doi/10.1105/tpc.109.073668

The Plant Cell, Vol. 22: 672–689, March 2010, www.plantcell.org ã 2010 American Society of Plant Biologists

transporters (MSTs) (Rolland et al., 2006; Buttner, 2007; Kocal

et al., 2008). However, the key genes responsible for regulating

the source–sink interaction for sugar transport remain elusive.

As a nonphotosynthetic male reproductive organ, the anther

obtains photosynthetic assimilates mainly from source organs to

support pollen development and maturation (Goetz et al., 2001).

Within the anther, the developing pollen is immersed in locular

fluid containing nutrients such as sugars and lipids from the

sporophytic (somatic) tissue tapetum (Pacini et al., 2006). The

early stages of pollen development are characterized by active

growth and high metabolic activity in the anther. Thus, anthers

have the highest sink strength in the developing flower, and large

amounts of sugars are mobilized to anthers to support their early

development (Oliver et al., 2007). At late stages, pollen matura-

tion requires the accumulation of starch, which functions as an

energy reserve for germination and thus serves as a marker of

pollen maturity (Datta et al., 2002). Disturbances in sugar

unloading and metabolism in the anther can significantly impair

pollen development and cause male sterility (Goetz et al., 2001;

Datta et al., 2002; Oliver et al., 2005; Mamun et al., 2006; Oliver

et al., 2007). Still, the regulatory mechanism underlying assim-

ilate partitioning remains poorly understood.

In this work, we report the identification of a key regulator gene

in rice (Oryza sativa), Carbon Starved Anther (CSA), encoding a

putative R2R3 MYB-type transcription factor that is involved in

regulating sugar partitioning during male reproductive develop-

ment. Results of sugar measurement and [14C]sucrose labeling

suggest that CSA may control assimilate partitioning in rice from

the topmost leaf (flag leaf) to the sink tissues in the flower,

particularly the anther. Consistent with this, the CSA gene is

preferentially expressed in the vascular tissue and the tapetumof

the anther, as well as in other sinks. Moreover, using chromatin

immunoprecipitation (ChIP) and electrophoretic mobility shift

assay (EMSA), we demonstrate that the CSA protein is able to

bind the promoter region of MST8, which encodes an MST.

RESULTS

Genetic Analysis of the csaMutant

Previously, we used g-ray radiation to generate a rice mutant

library in the 9522 background (O. sativa ssp Japonica) (Chen

et al., 2006a). The csamutant was isolated by its complete male

sterility under the growth condition with 30/24 6 18C day/night

temperature and 50 to 70% relative humidity (Figure 1).When the

csa plant was pollinated with wild-type pollen, all F1 progeny

displayed a normal phenotype, indicating that csa is a recessive

mutant. F2 progeny segregated for 419 normal and 126 mutant

plants (x2 = 1.028 for 3:1, P > 0.05), indicating monofactorial

recessive inheritance of the mutant characteristic.

Morphological Features of the csaMutant

Shorter Culm Length

During the seedling, tillering (formation of multiple shoots near

the base), and heading (formation of the reproductive shoots)

developmental stages, csa plants had no visible difference from

wild-type plants except that they were smaller in size (Figure 1A).

In mature plants, even though panicle (inflorescence) lengths of

wild-type and csa plants were similar (Figures 1C and 1M), the

culm (main stem) length of csa was slightly shorter than that of

the wild type (i.e., 75.26 2.3 cm for the wild type and 65.36 1.8

cm for csa; n = 25) (Figures 1A and 1L). The decreased culm

length of csa was mainly due to the reduced length of the

uppermost four internodes. Compared with the wild type, the

lengthsof internodes I to IV incsawere6.47, 3.27, 0.74, and0.74cm

shorter, respectively (Figures 1B, 1M, and 1N).

Abnormal Pollen Development and Maturation

Despite the reduced culm length, csa plants produced flowers

with apparently normal outer sterile organs called lemma and

palea (Figures 1C to 1E) but failed to generate normal anthers.

The csa anthers were white and smaller than those of the wild

type (Figures 1D to 1G). While examined using scanning electron

microscopy, the anther epidermal cells appeared to be smaller

than wild-type cells at stage 13 during anther development (see

Supplemental Figures 1A, 1B, 1E, and 1F online). Also, unlike

wild-type mature pollen, the csa pollen could not be deeply

stained by iodine–potassium iodide (I2-KI), and csa plants were

complete male sterile (Figures 1J and 1K). The csa pistils

appeared normal (Figures 1H and 1I), and we observed that

csa was able to produce normal seeds when backcrossed with

the wild-type pollen.

To detect possible cellular morphological alteration in the csa

mutant, we examined the wild-type and mutant anther develop-

ment in detail using transverse sections. Based on morpholog-

ical landmarks or cellular events visible under the light

microscope and previous classification of anther development

(Feng et al., 2001; Li et al., 2006), we recently further divided rice

anther development into 14 stages (Zhang and Wilson, 2009).

From stages 1 to 5, anther primordia differentiate and form the

characteristic anther structure with microspore mother cells,

somatic cells, and connective and vascular tissues. During

stages 7 to 9, microspore mother cells undergo meiosis and

generate dyads and then tetrads of haploid microspores.

Morphological defects were not detected in csa anthers until

stage 10 (Figure 2). At this stage, the middle layer of wild-type

anthers was thin and band-like, the endothecium became

narrower, and the tapetum begun to degenerate; also, micro-

spores appeared round and vacuolated (Figure 2C). However,

the csa middle layer and endothecium became abnormally

expanded and thicker than normal at this stage, and the micro-

spore had irregular appearance (Figure 2D). At stage 11, thewild-

type middle layer and endothecium degenerated, and typical

falcate (sickle-like shape) pollen grains were formed (Figure 2E),

whereas csa showed delayed degradation of the middle layer

and endothecium and produced severely abnormal pollen (Fig-

ure 2F). At stage 13, during mature pollen formation, the wild-

type anther wall layers were nearly completely degraded and

invisible. Inside the anther, mature pollen grains were deeply

stained with 0.05% toluidine blue, indicating that the wild-type

microspore is full of starch, lipids, and other storage materials

(Figure 2G) that are important for pollen viability and function.

However, in the csa anther, anther wall layers persisted at stage

CSA Regulates Sugar Partitioning in Rice Anther 673

13. In particular, the endothecium near the connective tissues

expanded, and the developing pollen disintegrated into debris

(Figure 2H).

When pollen grains were stained with 4’,6-diamidino-2-

phenylindole (DAPI), which stains nucleic acids, it was obvious

that late pollen developmental stages were abnormal in csa

mutants (see Supplemental Figures 2A to 2H online). At stage 11,

both wild-type and csa pollen grains could undergo the first

mitosis (see Supplemental Figures 2C and 2G online). Later, the

generative cell in wild-type pollen divided to form two sperm

cells, and the mature pollen was formed containing three cells

(i.e., a larger vegetative cell that surrounded two smaller sperm

cells at stage 13) (see Supplemental Figure 2D online). By

contrast, the second mitosis seemed to be delayed in the csa

pollen, which was smaller than the wild-type pollen, and no

obvious formation of the pollen with two sperm nuclei was

observed at this stage (see Supplemental Figure 2H online).

To further understand csa anther defects, we examined male

reproductive organs using transmission electron microscopy.

Consistent with the above observations, there was no obvious

difference of anther wall layers and microspores between the

wild type and csa at stage 9 (see Supplemental Figures 3A and

3D online). However, developmental defects of csa anther wall

layers and pollen were observed at stage 10. The wild-type

middle layer and tapetal layer appeared condensed and less

visible, and a vacuolated pollen grain with a round shape formed

in the wild type (see Supplemental Figures 3B and 3C online). By

contrast, csa middle layer and tapetal cells seemed less con-

densed and degenerated (see Supplemental Figure 3E online),

and the csa microspore appeared to have uneven cytoplasm

(see Supplemental Figure 3F online). At stage 13, the wild-type

anther wall cell layers were largely degenerated, and the major

remaining structures were cell walls of the epidermis and endo-

thecium, with relatively few (compared with the csa mutant)

Figure 1. Comparison of the Wild Type and the csa Mutant.

(A) Comparison of a wild-type plant (left) and a csa mutant plant (right) after heading. Bar = 20 cm.

(B) Comparison of the internode elongation of the wild type (left) and csa (right) at the heading stage. Bar = 10 cm.

(C) Comparison of the seed setting of the wild type (left) and csa (right). Bar = 5 cm.

(D) and (E) The spikelet of the wild type (D) and csa (E) after removing the palea and half the lemma. Bars = 2 mm.

(F) and (G) The wild-type anther (F) and the csa anther (G). Bars = 2 mm.

(H) and (I) The wild-type pistil (H) and the csa pistil (I). Bars = 2 mm.

(J) and (K) The I2-KI staining pollen grains of the wild type (J) and csa (K). Bars = 100 mm.

(L) Comparison of plant height between a wild-type plant (black bars) and a csa mutant (white bars).

(M) Comparison of the length of panicle and top four internodes (I to IV, where I is the uppermost) between wild-type (black bars) and csa (white bars)

plants. Data presented are means of results from 25 plants. Error bars indicate SD.

(N) Comparison of relative length percentage of top four internodes between wild-type (left) and csa (right) plants.

674 The Plant Cell

hair-like cuticle structures on the anther surface (see Supple-

mental Figure 3G online). At this stage, the wild-type pollen was

full of storage materials, such as starch granules and lipids (see

Supplemental Figure 3H online), with a normal pollen wall con-

taining exine and intine layers (see Supplemental Figure 3I

online). However, the csa epidermis and endothecium seemed

to be less degenerated and abnormally persisted with irregular

cell shape at stage 13 (see Supplemental Figure 3J online). In

addition, at stage 13, scanning electron microscopy analysis

confirmed that the csa anther had an abnormal cuticle (see

Supplemental Figures 1C, 1D, 1G, and 1H online). Although the

csa pollen wall seemed to have normal exine, the intine was less

condensed, and pollen grains collapsed with reduced accumu-

lation of starch and other storage materials (see Supplemental

Figures 3K and 3L online). These observations suggest that CSA

plays a crucial role in anther and pollen development in rice.

Altered Assimilate Partitioning during Pollen Development

Morphological analyses indicated that the csa mutant had de-

fects during late anther development and pollen maturation,

especially the reduction in starch accumulation in the pollen

grain. Nutrients such as starch are preferentially accumulated in

the pollen to provide energy for pollination, and the starch level is

a metabolic marker of pollen maturity (Datta et al., 2002). To test

whether csa was abnormal in sugar partitioning from the source

tissue (the flag leaf) to the sink tissue (the developing anther), we

stained the flag leaf and internode I (the uppermost internode)

using I2-KI to observe starch distribution by the end of the day. At

stage 11, we did not observe obvious starch accumulation in

either wild-type or csa flag leaves (Figure 3A). Meanwhile, strong

starch staining was observed at the base region of the wild-type

internode I, as well as that of the csa internode I (Figures 3C, 3E,

and 3G), confirming the accumulation of starch within the stem

tissue, which acts as a temporary sink during rice reproductive

development (Scofield et al., 2007). At stage 13, starch deposi-

tion was detected at reduced levels in the flag leaf and the stem

of the wild type (Figures 3B, 3D, and 3F), suggesting that the

accumulated starch is converted and allocated for reproductive

development. However, at stage 13, abnormal starch accumu-

lation was observed in the csa internode I region and the flag leaf

(Figures 3B, 3D, and 3H). It appears that the photosynthetic

sugar in csawas not transported normally from the flag leaf to the

anther and other sink tissues during late pollen development.

To measure carbohydrate distribution, we employed gas

chromatography–mass spectrometry (GC-MS) analysis to test

the content of sugars in the anther, lemma/palea, and flag leaf of

the wild type and csa. Results of GC-MS analysis revealed that

the levels of sucrose, glucose, and fructose in the anther de-

creased gradually during wild-type anther development (Figure

4A). The starch level in the wild type showed a 25-fold increase

from stage 9 to stage 13, suggesting that the conversion from

sucrose, glucose, and fructose to starch occurs normally in the

wild-type anther during pollen formation (Figure 4A; see Supple-

mental Table 1 online). Compared with the wild type, the csa

anther had significantly reduced levels of glucose and fructose

from stage 9 to stage 13 (P < 0.05), and sucrose levels were

slightly lower (Figure 4A; see Supplemental Table 1 online),

suggesting that the csa anther likely has defects in importing

sugars from the source tissues. Furthermore, although starch

content in the csa anther seemed to be normal at stage 9, it

decreased to;22% (5.5mg/g freshweight [FW] for thewild type

and 1.2 mg/g FW for csa) and 49% (152.3 mg/g FW for the wild

Figure 2. Transverse Sections Showing Anther and Microspore Devel-

opment of the Wild Type and csa.

Four stages of anther development in the wild type and the correspond-

ing of the csa mutant were compared. Transverse sections were stained

with 0.05% toluidine blue O. Images from wild-type plants are shown in

(A), (C), (E), and (G); (B), (D), (F), and (H) are the csa mutant. (A) and (B),

stage 9; (C) and (D), stage 10; (E) and (F), stage 11; (G) and (H), stage 13.

E, epidermis; En, endothecium; ML, middle layer; T, tapetum; Msp,

microspore; MP, mature pollen; St, stomium. Bars = 15 mm.

CSA Regulates Sugar Partitioning in Rice Anther 675

type and 75.2 mg/g FW for csa) of normal levels at stages 11 and

13, respectively (Figure 4A; see Supplemental Table 1 online).

At early anther development, the rice outer floral organs lemma

and palea likely act as the sink tissue, assimilating carbohydrate

from source tissues; later during pollen starch synthesis, these

outer floral organs were proposed to function as the source

organs supplying carbohydrate for pollen maturation (Abebe

et al., 2004). Consistent with this hypothesis, we observed that

contents of glucose and fructose in the wild-type lemma/palea

decreased from stage 9 to stage 13. The levels of glucose and

fructose in the wild-type lemma/palea were slightly lower than

those of the csa mutant at stage 9 (Figure 4B). The sucrose

amounts in the wild-type and csa lemma/palea were very similar

at stage 9, but the csa lemma/palea at stages 11 and 13 had

notably lower contents of sucrose than those of the correspond-

ing wild-type lemma/palea, respectively. Also, we observed

;30% lower starch content in the csa lemma/palea compared

with the wild type at these stages (Figure 4B; see Supplemental

Table 1 online).

Accompanied by the reduced accumulation of starch in the

csa anther and lemma/palea, the levels of sucrose and starch in

the flag leaf were increased in the csa mutant compared with

the wild type (Figure 4C). In particular, the starch content in the

csa flag leaf increased to about twofold of that in the wild-type

flag leaf at stage 13 (Figure 4C; see Supplemental Table

1 online).

These results suggest that the csamutant likely has defects in

sugar partitioning from the flag leaf to the lemma/palea and

anther. The remarkable decrease of sucrose and starch levels in

the csa anther at the late pollen development stage might have

resulted from the disruption of carbohydrate uptake or utilization

in anther, causing male sterility.

CSA Regulates Carbon Accumulation in the Anthers

To further test the role of CSA in regulating sugar partitioning

during rice male reproductive development, we performed a

[14C]sucrose feeding assay using excised stems containing

leaves and the panicle from the wild type and csa to assess the

sugar distribution from the stem to the sink anther at stages 11

and 13. The excised stem containing internodes I to IV, as well as

the flag leaf and flowers, were placed and incubated in water

containing added [14C]sucrose, with internode IV being sub-

merged in the water directly. After 12 h of treatment, the amount

of isotope was tested using amiddle section of each of internode

I, II, and III; the sections were designated S1, S2, and S3 from the

bottom to the top. At stage 11, the isotope signal strengths in S1

segments were similar in the wild type and the csamutant, but in

the S2 and S3 segments, the csa stem had slightly more isotope

signals than those of the wild type (Figure 5A; see Supplemental

Table 2 online). At the stage 13, from S1 to S3, the levels of

accumulated isotope signal in csa were all higher than those of

the wild type (Figure 5B; see Supplemental Table 2 online).

Conversely, we observed the isotope signals in the wild-type

lemma/palea were higher than those of csa at stages 11 and 13.

This analysis suggested that csa was defective in sugar parti-

tioning from the leaf to flower via stem during rice reproductive

development.

Figure 3. I2-KI Staining the Flag Leaf and Stem in Wild Type and csa.

(A) I2-KI staining of flag leaves from the wild type (left) and csa (right) at stage 11.

(B) I2-KI staining of flag leaves from the wild type (left) and csa (right) at stage 13.

(C) I2-KI staining of stems from the wild type (top) and csa (bottom) at stage 11.

(D) I2-KI staining of stems from the wild type (top) and csa (bottom) at stage 13. Arrows in (C) and (D) indicate starch deposition.

(E) and (G) I2-KI–stained free-hand sections of stem cell division zones of the wild type (E) and csa (G) at stage 11.

(F) and (H) I2-KI–stained free-hand sections of stem cell division zones from the wild type (F) and csa (H) at stage 13.

Arrows indicate starch deposition in (C) and (D); arrows indicate the vascular tissue (VT) in (E) to (H). Bars = 1cm in (C) and (D) and 150 mm in (E) to (H).

676 The Plant Cell

In addition, the [14C]sucrose feeding assay was performed

using the excised panicles that included a portion of internode I

to detect sugar partitioning in the anther and lemma/palea of csa.

After a 12-h treatment, we observed accumulated isotope sig-

nals in thewild-type anther at stages 9, 11, and 13 (Figure 5C; see

Supplemental Table 2 online), indicating that source tissues

supply abundant sugars for pollen development. By contrast,

signals in the csa anthers were very low at stage 9. At stage 11,

the isotope level had an increase in the csa anther, but it was not

as great as that in the wild type (Figure 5C; see Supplemental

Table 2 online). At stage 13, the isotope signals in the csa anther

were clearly lower than that in the wild type (Figure 5C). Similar to

the distribution of isotope signals in the rice anther, we observed

higher isotope signals in the wild-type lemma/palea than in the

csa mutant from stage 9 to stage 13 (Figure 5D; see Supple-

mental Table 2 online). Consistently, the isotope signals in the

csa anther and lemma/palea were observed to be lower than

those of the wild type after 1- and 6-h treatments, respectively

(see Supplemental Figure 4 online), while the accumulated

isotope signals in both the wild type and the csa mutant

increased from 1 to 12 h after treatment. This suggests that the

redistribution of labeled sucrose occurred within a very short

period (an hour), and, not surprisingly, the total amount of

redistributed products increased as time elapsed.

To determine the chemical nature of the labeled molecules in

anthers, we separated the sugars (sucrose and hexoses) in the

soluble extract from anthers after a 1- and 12-h treatment with14C-labeled sucrose using thin layer chromatography. We ob-

served the signals of the labeled sucrose and hexose (glucose

and fructose) in both the wild type and csa (see Supplemental

Table 3 online). The level of both sucrose and hexose, indicated

by the radioactive signal, was lower in the mutant anthers than

thewild-type anthers at any timepoint (seeSupplemental Table 3

online). For wild-type plants, a large proportion of the labeled

products were hexoses an hour after the treatment. In csa

mutants, the level of labeled hexose was only one-third that of

the wild type, while the level of sucrose was only slightly lower

than that of the wild type (see Supplemental Table 3 online). By

12 h after treatment, the fraction of sucrose increased in wild-

type plants, whereas the relative ratio of sucrose and hexosewas

similar to that of the csa mutant (see Supplemental Table 3

online). As a result, the csa mutant anthers seemed to be more

deprived of sucrose compared with the wild-type anthers at this

time point. This observation suggests that the csa mutation has

influenced the redistribution of the radiolabeled sucrose and that

the most immediate and dramatic alteration is the reduced level

of hexose in the csa anthers.

These observations indicated that the csamutation caused the

defect of carbon accumulation in the anther; thus, we named this

gene Carbon Starved Anther.

Isolation of the CSA Gene

To isolate the CSA gene, we initially mapped the CSA locus

between two InDelmolecularmarkers, OS104 andOS106, on the

short arm of rice chromosome 1. Tomore precisely localizeCSA,

750 mutants from a F2 mapping population were identified and

analyzed using seven polymorphic InDel markers (see Supple-

mental Table 4 online). Finally, CSA was located between two

InDel markers Z134 and Z138, which define a region of 23 kb

(Figure 6A). By sequencing the mutant genomic DNA, we found

that both a single nucleotide deletion and a G-to-A transition had

occurred in a gene, with a gene ID Os01g16810 (The Institute for

Genomic Research), Os01g0274800 (National Center for Bio-

technology Information), or Os01t0274800-01 (Rice Annotation

Project Database) (Figure 6B), causing a frame shift and prema-

ture translational termination (see Supplemental Figure 5 online).

Those are the only mutations in the entire gene as well as in the

Figure 4. Sugar and Starch Levels in the Wild Type and csa.

Sugar and starch levels at stages 9, 11, and 13 in anther (A), lemma/

palea (B), and flag leaf (C). Data presented are means 6 SE (n = 3) with

units of mg/g FW. Fru, fructose; Glu, glucose; Suc, sucrose; S, starch.

CSA Regulates Sugar Partitioning in Rice Anther 677

2-kb upstream and 1-kb downstream regions. Furthermore, we

determined the intron-exon pattern of the Os01g0274800 gene

by comparing the genomic sequence with the obtained full-

length cDNA (AK107461) from the Rice Genome Resource

Center (RGRC-NIAS; http://www.rgrc.dna.affrc.go.jp/stock.

html) (Figure 6B).

To further verify the identity of this gene asCSA, we performed

a functional complementation experiment. A binary plasmid

carrying an ;4.3-kb wild-type genomic fragment containing

2157-bp upstream sequence, 1665-bp coding region of

Os01g0274800, and 525-bp downstream sequence from the

BAC clone AP000837 was able to rescue the male-sterile phe-

notype of the csa homozygous plants (Figures 7A to 7C). The

complemented lines displayed yellow anthers with starch-filled

pollen grains (Figure 7C) and high seed-setting rate (right of

Figure 7D), which were similar to those of the wild type (Figure 7A

and left of 7D). The carbohydrate accumulation within the flag

leaf and the internode I base was normal in the complemented

lines at anther stage 13 (Figures 7E and 7F). The reduced

accumulation of starch and increased sucrose level in the flag

leaf of the complemented plants were also observed by sugar

measurements (Figure 7G; see Supplemental Table 5 online).

Those results confirm that the csa mutant phenotype is caused

by Os01g0274800 dysfunction.

The CSA open reading frame encoded a putative R2R3-type

MYB transcription factor of 268 amino acids with two MYB

domains (Figure 6B; see Supplemental Figure 5 online). Phylo-

genetic analysis between CSA and its closest 14 homologs

indicated that CSA is closely related to the R2R3 MYB proteins

MYB56 from Arabidopsis and LOC_ Os08g33800 from rice

(Figure 6I; see Supplemental Figure 6 online). Also, we observed

two putative nuclear localization signal sequences in CSA using

the P-sort program (http://psort.ims.u-tokyo.ac.jp/form.html)

analysis (see Supplemental Figure 5 online). To confirm the

CSA nuclear localization, we constructed a translation fusion

between the full-length CSA coding region and the cDNA for the

green fluorescent protein. The CSA-GFP fusion construct and

the GFP alone control, both driven by the cauliflower mosaic

virus 35S promoter, were introduced into onion epidermal cells

by particle bombardment. As expected, the CSA-GFP fusion

protein was observed exclusively in the nucleus (Figures 6C to

6E). By contrast, the free GFP was found in the nucleoplasm, as

well as in the cytoplasm (Figures 6F to 6H). This result suggests

that CSA is localized to the nucleus.

CSA Expression Is Mainly in Vascular Tissues and

the Tapetum

The main morphological defects of csa occurred in anther

development due to the biochemical abnormality in sugar

partitioning into flower/anther, whereas there was no dramatic

phenotype for vegetative development. To test how CSA acts in

the affected mutant tissues to regulate sugar partitioning, we

analyzed the CSA expression pattern using RT-PCR, promoter-

b-glucuronidase (GUS) fusions, and in situ hybridization.

RT-PCR analysis using total RNA prepared from rice vegeta-

tive and reproductive organs showed that the CSA transcripts

were undetectable in stem and leaf, but detectable in root. In the

sterile empty glume, which surrounds the rice flower, no CSA

expression signal was observed. Strong expression of CSA was

detected in the lemma and palea and weaker expression in the

pistil and seed. As expected, the CSA transcript was clearly

detected in the anther from stage 9 to stage 13 (Figure 8A).

Analysis of transgenic rice lines with the GUS reporter gene

driven by the CSA promoter (;2.3 kb) indicated that in the

germinating seedlings, GUS expression was mainly detected in

coleoptile and root vascular tissue, as well as the primordia of

lateral root (Figures 8B and 8C; see Supplemental Figures 7A and

Figure 5. 14C-Signal Accumulation in the Flower/Anther and Stem of the Wild Type and csa after 12-h Treatment.

(A) 14C-signal accumulation in the stems of wild-type and csa plants at stage 11.

(B) 14C-signal accumulation in the stems of the wild type and csa at stage 13.

(C) 14C-signal accumulation in the anther of the wild type and csa at stages 9, 11, and 13.

(D) 14C-signal accumulation in the lemma/palea of the wild type and csa at stages 9, 11, and 13.

S1 to S3, stem segments from the base to the top. The data are given as means 6 SE (n = 3). The unit is expressed as cpm/mg, FW.

678 The Plant Cell

Figure 6. Molecular Identification of CSA.

CSA Regulates Sugar Partitioning in Rice Anther 679

7B online). We did not observe GUS staining in stem and leaf

blades, but the staining was visible in the leaf collar (Figure 8D).

GUS expression was enhanced in the region of wounding and

callus (Figure 8E; see Supplemental Figure 7C online). In flowers,

theGUSexpression could be observed in the veins of the lemma/

palea and pistil (Figures 8F and 8G; see Supplemental Figures 7D

and 7H online). In addition to the expression of GUS in anther

vascular tissue from stage 9 to stage 13 (Figures 8H to 8K; see

Supplemental Figures 7E to 7G online), we found GUS expres-

sion in anther wall layers at stage 9 (Figures 8H and 8I). This

suggests that CSA is likely expressed in the anther wall layers at

the early stage when the tapetum is present. During later stages,

as the tapetum degenerated, the expression of CSA is likely

restricted in the anther vascular tissue. Through the observation

of autofluorescence triggered by UV light, we detected the xylem

cells among the anther vascular tissue where no GUS staining

was detected (see Supplemental Figure 7G online), probably

because these cells are not viable. Also, GUS activity was

observed in the embryo and the dorsal vascular tissues of seeds

(see Supplemental Figures 7H to 7J online).

To further confirm the CSA expression pattern, we performed

RNA in situ hybridization with wild-type floral and root sections.

Figure 6. (continued).

(A) Fine mapping of the CSA gene on chromosome 1. Names and positions of the molecular markers are indicated on the vertical line. AP000837 is the

accession number of the relevant genomic sequence. cM is the unit of genetic distance (centimorgans). Numbers in parentheses represent

recombination events in the appropriate interval. The CSA locus was mapped to a 23-kb region between molecular markers Z134 and Z138.

(B) A schematic representation of the exon and intron organization ofCSA. The mutant sequence has a nucleotide deletion and a G-to-A transition in the

first exon. +1 Indicates the starting nucleotide of translation, and the stop codon (TAG) is +1098. Black boxes indicate exons; intervening lines indicate

introns; gray boxes indicate untranslated regions.

(C) to (E) The onion epidermal cell that expressed CSA-GFP.

(F) to (H) The onion epidermal cell that expressed GFP as control. Bars = 50 mm, all six panels are at the same magnification.

(I) Phylogenetic analysis of CSA and its 14 close homologs. The proteins were named according to their gene names from Arabidopsis thaliana and rice,

and others were according to their National Center for Biotechnology Information accession numbers followed by their species names (abbreviation).

Os GAMYB is defined as an outgroup. The scale bar indicates the number of amino acid substitutions per site. The alignment for the constructed tree is

shown in Supplemental Figure 6 online, with sequences listed in Supplemental Data Set 1 online.

Figure 7. Complementation of the csa Mutant and Phenotype Analysis.

(A) to (C) Anthers and I2-KI–stained pollen grains of the wild type (A), the csa mutant (B), and the complemented line ([C]; CL).

(D) Comparison of the seed setting of the wild type, csa, and the complemented line.

(E) I2-KI–stained flag leaves of the wild type, csa, and the complemented line at stage 13.

(F) I2-KI–stained internode I stems of the wild type, csa, and the complemented line at stage 13.

(G) Sucrose and starch levels in flag leaves of the wild type, csa, and the complemented line at stage 13.

Bars = 2 mm in (A) to (C), 5 cm in (D), and 1 cm in (E) and (F). Suc, sucrose; S, starch.

680 The Plant Cell

Consistent with the GUS staining results, the CSA expression

signals were detected in the vascular tissues of the root and

anther (Figures 8L to 8S). From stage 9 to stage 13, the CSA

expression signals were observed in the tapetum and vascular

tissue of anther connective tissue compared with the control

signals observedwith the sense probe (Figures 8L to 8Q). At stage

9, we also observed CSA expression signals at the microspore

surface (Figure 8L). In addition, detectableCSAexpression signals

were observed in root vascular tissues (Figures 8R and 8S).

Therefore, the location of CSA expression is consistent with

the hypothesis that this gene is associated with the sugar

partitioning into the anther, a major sink organ in rice.

Figure 8. CSA Expression Pattern.

(A) Spatial and temporal expression analyses of CSA by RT-PCR. RNAs were extracted from the root of 15-d-old seedlings, the shoot, leaf, glume, and

lemma/palea from the plants at heading stage. L/P, lemma and palea.

(B) to (K) GUS activity in the pCSA-GUS line.

(B) CSA expression in the root vascular tissue.

(C) GUS activity in the region of lateral root initiation.

(D) GUS activity in the leaf collar.

(E) GUS activity in the wounding tissue.

(F) GUS activity in the lemma/palea.

(G) GUS activity in the pistil.

(H) to (K) GUS activities in anther of stage 9 (H), stage 10 (I), stage 11 (J), and stage 13 (K).

(L) to (S) In situ analyses of the CSA expression in anther at stage 9 ([L] and [M]), stage 11 ([N] and [O]), and stage 13 ([P] and [Q]).

(R) and (S) In situ analyses of CSA expression in the root; pink color in (R) indicates the CSA expression.

(L), (N), (P), and (R) Probed with the CSA antisense probe.

(M), (O), (Q), and (S) Probed with the CSA sense probe.

Arrows indicate theCSA expression positions. T, tapetum; Msp, microspore; VT, vascular tissue. Bars = 1mm in (B) and (C), 1 cm in (D) and (E), 2 mm in

(F) to (K), 30 mm in (L) to (Q), and 60 mm in (R) and (S).

CSA Regulates Sugar Partitioning in Rice Anther 681

CSA Directly RegulatesMST8 in Rice

MSTs have the ability to transport a variable range of monosac-

charides across membrane barriers and have been shown to

play an important role in assimilate supply for sink tissue devel-

opment (Buttner, 2007; Wang et al., 2008b). In rice, one MST

member, MST8 (Os01g38670), is expressed in the tapetum,

microspore, and anther vascular bundle, which has been shown

to be a key component of the anther apoplastic sugar transport

pathway (Oliver et al., 2005, 2007; Mamun et al., 2006). As

expected, MST8 was expressed from stage 9 to stage 13 in the

anthers of wild-type plants (Figure 9A; seeSupplemental Figure 8

online). In the csa mutant, very low MST8 expression was

detectable in the anther from the early stage to the late stage

(Figure 9A; see Supplemental Figure 8 online). In the wild-type

lemma/palea, MST8 had higher expression at stages 11 and 13

(Figure 9B; see Supplemental Figure 8 online). By contrast, only

weak expression was detected in the csa lemma and palea at

these stages (Figure 9B).

CSA is a putative R2R3 MYB transcription factor, which is

expected to regulate gene expression by binding to the pro-

moters of the target genes. Putative MYB binding sequences

(MBSs) were identified using the tools described in plant

CARE (for cis-acting regulatory element) (http://bioinformatics.

psb.ugent.be/webtools/plantcare/html/; Lescot et al., 2002):

CCAAT-box and (pyAAC(G/T)G) in the promoter region ofMST8.

To test whether CSA has the ability to bind the promoter region

of MST8, we developed rabbit polyclonal antibodies against a

bacterially expressed recombinant CSA fragment. The specific-

ity of the CSA antibody was confirmed using protein gel blot

analysis, which detected a product of 30 kD, the expected size of

CSA in the wild-type anther. By contrast, no CSA signal was

observed in the csa mutant (see Supplemental Figure 9 online).

Our quantitative ChIP-PCR results indicated that two DNA frag-

ments, MST8-1 (208 bp) and MST8-2 (191 bp), of the upstream

MST8 region containing the predicted MYB transcription factor

binding sites (CAACGG) were enriched when the affinity-purified

CSA antibodies were used (Figures 9C and 9D). Meanwhile, no

enrichment of either a 191-bpMST8-3 DNA fragment (the 39 endwas 302 bp from the 59 end of the predicted MBS in MST8-2) of

MST8 upstream region without the predicted CCAAT-box or a

318-bp upstream region of ACTIN1 was observed using the

affinity-purified CSA antibodies (Figure 9D).

Furthermore, an EMSA revealed that the recombinant CSA

protein is able to bind a 114-bp DNA fragment containing two

CCAAT-boxes of the MST8 upstream region (Figure 9E). When

unlabeled DNA fragments were present as competitors, the

excess MST8 competitor DNAs reduced the complex formation

in a concentration-dependent manner (Figure 9E). These results

support the hypothesis that CSA directly regulates MST8.

CSA Affects the Expression of Genes Involved in Sugar

Partitioning in Flower/Anther

Because sugar levels were greatly altered during csa anther

development, we examined the mRNA levels of several key

genes involved in the cleavage, transport, and utilization of

sucrose in rice anther and lemma/palea. We compared the

expression of rice INV4, SUT3, UGP2, and GBSS1 at stages 9,

11, and 13 of anther development in the wild type and the csa

mutant using quantitative RT-PCR analysis. The expression

levels of INV4, SUT3, UGP2, and GBSS1 in the anthers and

lemma/palea were comparable between the wild type and the

complemented lines at stages 11 and 13 (see Supplemental

Figure 10 online) but were reduced in the csa mutant (see

Supplemental Figure 10 online). Therefore, we speculated that

either those genes are regulated by CSA or their expression is

influenced by the carbon starvation in csa mutants.

INV4 encodes an extracellular cell wall–bound invertase

(CW-INV, CIN, or INV), which is a key component of the sucrose

phloem unloading pathway, and its activity for cleaving sucrose

is a biochemical marker of sink strength for carbohydrate

partitioning (Ranwala and Miller, 1998; Oliver et al., 2005,

2007). SUT3 is one of five SUTs identified rice (Aoki et al.,

2003). UDP-glucose pyrophosphorylase (UGPase) has the ability

to reversibly convert glucose-1-phosphate and UTP into UDP-

glucose and pyrophosphate. In the anther, UDP-glucose is

primarily involved in the synthesis of starch, and two homologous

UGPase genes,UGP1 andUGP2, are present in the rice genome

(Chen et al., 2007; Woo et al., 2008; Mu et al., 2009). Unlike the

expression alteration of UGP2 in the csa mutant, UGP1 was

observed with no obvious expression change in the csa mutant

during the anther development (see Supplemental Figures 10E

and 10J online). GBSSI encodes a starch synthase (GBSS) that

constitutes the final step in which the glucose moiety of ADP-

glucose is transferred to the nonreducing end of the starch

molecule in the starch granule (Ohdan et al., 2005).

Even though the INV4 promoter contains one CAACTG motif

and the GBSSI promoter contains one CAACGG motif, no

association of CSA with these motifs was detected using ChIP-

PCR assay. Moreover, no predicted MBS was observed in the

promoter regions of SUT3 and UGP2. These results implied that

CSA may indirectly regulate the expression of INV4, SUT3,

UGP2, and GBSS1 during anther development. However, we

cannot exclude the possibility that CSA, in concert with another

unknown factor(s), coregulates the expression of these genes

that have no apparent MYB binding sites in their promoters.

DISCUSSION

CSA Is Required for Anther Development and

Pollen Maturation

Rice is one of the most important crops in the world, and in rice

breeding, the fertility of pollen grains is critical for rice yield. Many

cytoplasmic and nuclear mutations leading to male sterility lines,

which are of agricultural importance for the production of hybrids

to improve rice yield. Production of functional pollen grains in

flowering plants relies on cooperative functional interactions

between gametophytic and sporophytic tissues within the anther

(McCormick, 2004; Scott et al., 2004; Ma, 2005; Zhang and

Wilson, 2009). In this study, we report a novel rice gene, CSA,

which is critical for male gametophyte development. The csa

mutant shows delayed degradation of anther wall cell layers and

aborted pollen maturation during late pollen development.

682 The Plant Cell

The presence of sufficient levels of sucrose is of vital impor-

tance for the growth of the male reproductive cells in plants. In

Lilium, it has been demonstrated that the cells of the outer anther

wall cell layers and the connective tissue are interconnected by

cytoplasmic bridges called plasmodesmata, allowing assimi-

lates to pass, via the symplastic pathway, from cells of the

vascular bundle to the most internal portions of the middle layer

(Clement and Audran, 1995). Because plasmodesmata are not

detected between the middle layer and tapetum, sugars trans-

ported to pollen in the locule have to cross the middle layer and

tapetum by the apoplastic pathway, which requires transport

across the plasma membrane. The mechanism underlying the

apoplastic pathway is complex, involving many enzymatic sys-

tems to enhance the control of sugar transport (Clement and

Audran, 1995). INV4 and MST8 are proposed to be key compo-

nents of apoplastic sugar transport pathway from tapetal cells to

locular fluid because of their similar expression patterns during

anther development (Oliver et al., 2005, 2007; Mamun et al.,

2006). In this study, we show that control of sugar partitioning by

CSA greatly affects male reproductive tissue development in

Figure 9. Regulation of Rice MST8 by CSA.

(A) and (B) Relative mRNA levels of MST8 in the anther (A) and the lemma/palea (B) of the wild type (black), csa (white), and complemented line (CL)

(gray) analyzed by real-time PCR. Error bars indicate SD; each reaction has four quantitative PCR biological replicates.

(C) Predicted CCAAT-boxes of rice MST8 and ACTIN1 upstream regions. Black boxes indicate canonical binding sites for plant R2R3-MYB proteins

of the form pyAAC(G/T)G (CCAAT-box); numbers indicate the position of these motifs relative to the putative transcriptional start site; the bent

arrow denotes the translational start site. The gray fragments (MST8-1,MST8-2,MST8-3, and Actin1) indicate the position used in ChIP-qPCR assays.

MST8-1 and MST8-2 contain the predicted CCAAT-motif, and MST8-3 has no predicted CCAAT-motif as the control. The black fragment (MST8-4)

with two predicted CCAAT-motifs was used in gel shift assays.

(D) ChIP enrichment test by PCR shows the binding of CSA to the regulatory region ofMST8-1 andMST8-2. The fold enrichments in the IP sample over

the minus antibody control are shown. Error bars indicate SD; each reaction has four quantitative PCR biological replicates.

(E) Recombinant CSA binding to the promoter region of MST8-4 with containing two CCAAT-boxes was determined by EMSA. The binding complex

could be outcompeted with increasing quantities of unlabeled MST8-4 DNA fragments (325, 350, and 3100 of unlabeled MST8-4 DNA fragments).

CSA Regulates Sugar Partitioning in Rice Anther 683

rice. The expression levels of the INV4 andMST8 aswell asSUT3

were highly reduced in the csa anther (Figure 9; see Supplemen-

tal Figures 8 and 10 online). Thus, it is likely that the down-

regulation of these genes in csa causes an abnormal sugar

supply of the apoplastic sugar partitioning pathway from the

tapetum to locular fluid during anther development.

CSA Is a Member of the Family of R2R3 MYB

Transcription Factors

MYB domain proteins in plants form a superfamily. In the rice

genome, 183 putative MYB genes have been identified (Chen

et al., 2006b). In rice, GAMYB is an MYB transcription factor

controlling rice pollen development (Kaneko et al., 2004). The

gamyb-2 mutant microspore mother cells were abnormal, and

themutant microspores could not adhere to tapetal cells starting

from the tetrad stage (Kaneko et al., 2004). AID1 encodes a

protein with a single MYB domain that controls anther dehis-

cence and pollen development (Zhu et al., 2004). In addition,

three MYB proteins (MYBS1, MYBS2, and MYBS3), each with a

single MYB domain, can regulate the expression of a-amylase

during rice seed development (Lu et al., 2002). Compared with

these previously studied MYB proteins, the CSA protein likely

has a unique function in the MYB family in controlling carbon

partitioning during the anther development and pollen matura-

tion. Phylogenetic analysis indicates that CSA is most similar to

the rice protein LOC_Os08g33800 and Arabidopsis protein

MYB56. The function of theArabidopsis gene is unknown. Future

experiments are required to test whether those genes are in-

volved in the transport of carbohydrates.

CSA Controls Sugar Partitioning into the Anther

Sucrose is the main metabolic substrate for starch synthesis in

nonphotosynthetic sink tissues (Ohdan et al., 2005). Upon arrival

in sink tissues, sucrose may be cleaved by invertase and

unloaded by hexose transporters. The importance of cleavage

of sucrose by invertase in the apoplastic pathway has been

supported by the identification of hexose transporters (Sherson

et al., 2003; Buttner, 2007). The expression pattern of MSTs

suggests that they act mainly in hexose uptake into the sink. In

Arabidopsis, STP2 encodes a high-affinity, low-specificity MST

with the ability to transport various hexoses and pentoses. This

gene is specifically expressed during pollen maturation and

germination, and it plays a role in callose degradation for pollen

maturation (Truernit et al., 1999). Similarly, Arabidopsis STP6

(Scholz-Starke et al., 2003), STP9 (Schneidereit et al., 2003), and

STP11 (Schneidereit et al., 2005) have been shown to be ex-

pressed during anther/pollen development. Also, several MSTs

from rice have been demonstrated to play a crucial role in sugar

distribution (Toyofuku et al., 2000; Ngampanya et al., 2003;

Mamun et al., 2006; Wang et al., 2007, 2008b).

RiceMST8 has a spatial and temporal expression pattern that

strongly resembles that of INV4, which is expressed in rice

anthers (Oliver et al., 2005, 2007;Mamun et al., 2006), suggesting

thatMST8may function in the same pathway as INV4 controlling

photosynthetic carbon allocation. More intriguingly, expression

analyses revealed that CSA transcripts accumulated in vascular

tissues of the anther, as well as in the tapetum, which is similar to

the pattern seen with MST8. In csa flowers, expression levels of

INV4 and MST8 are dramatically reduced. Therefore, we spec-

ulate that, in the csa mutant due to the downregulated expres-

sion of INV4 and MST8, the sugar unloading may be severely

blocked, impairing the sugar concentration gradients between

sources and organs driving source-to-sink transport (stems/

flowers). In other words, the lack of CSA function may cause

reduced unloading, leading to the accumulation of sugars in the

leaf/stem. This is consistent with results of our feeding experi-

ment with [14C]sucrose. If we consider that, within only 1 h, the

accumulation of labeled products in anthers should reflect the

immediate impact of transport or unloading of sugars, this may

well explain why the hexose level in csa is dramatically lower than

in wild-type anthers (see Supplemental Table 3 online). Further-

more, genes related to sucrose utilization, such as UGP2 and

GBSS1, were downregulated by theCSAmutation during anther

development, resulting in the decreased metabolic capacity to

convert sucrose to starch in the csa mutant (see Supplemental

Figure 10 online). Consequently, sugar distribution indicated by

the [14C]sucrose feeding analysis in the csa mutant is abnormal

during anther development.

More importantly, the fact that CSA could directly bind to the

promoter region of MST8 suggests that CSA is likely a key

transcriptional regulator for photosynthate partitioning from

leaves to anthers. For other genes, such as INV4, it is possible

that they are indirectly regulated by CSA. One way this might

occur, for example, is that the absence of CSA function leads to a

reduction in sugar concentration, which serves as signal to

regulate the expression of INV4 and other genes. It is worth

mentioning that although the uptake of sugars in lemma/palea

seems to be affected in the csamutant, they do not demonstrate

any visible phenotype. It is possible that the morphological

development of lemma/palea largely precedes the time of func-

tional CSA.

In summary, this work characterizes the key role of CSA in

regulating sugar partitioning required for rice anther develop-

ment and pollen maturation. One downstream target gene,

MST8, has been shown to be a likely direct target gene of

CSA. According to our model, the reduced expression level of

MST8 in the csa mutant causes specific defect in sugar uptake

(or unloading) into anthers, which results in carbon starvation and

male sterility. Moreover, the reduced uptake of sugars in anthers

alters the sink-source relationship, and it further influences the

partitioning of assimilates and the expression of other genes in

the pathway. The characterization of the csa mutant provides

new insight into the genetic and transcriptional control of assim-

ilates partitioning in plants.

METHODS

Mutant Material and Growth Conditions

The F2mapping population was generated from a cross between the rice

(Oryza sativa) csamutant (ssp japonica) andGuangLuAi (ssp indica). In the

F2 population, male-sterile plants were selected in the winter season

(short-day light, ;12-h light, in Hainan province, China) for gene map-

ping. Other plants were grown in a greenhouse with a 30/24 6 18C day/

684 The Plant Cell

night temperature, 50 to 70% relative humidity, and a light/dark period of

13 h/11 h.

Characterization of Mutant Phenotype

Plantsmaterials were photographedwith aNikon E995 digital camera and

a Motic K400 dissecting microscope. For cross section observation, the

materials were collected and fixed as described (Li et al., 2006). Floral

tissues were embedded in Spurr’s resin (Sigma-Aldrich); semithin (4 mm)

sections were made using an Ultracut E ultramicrotome (Leica Micro-

systems) and stainedwith 0.05% toluidine blue (Li et al., 2006). Transverse

sectionswere photographed using a Nikon E600microscope and a Nikon

DXM1200 digital camera. For transmission electron microscopy obser-

vation, anthers were fixed, washed, embedded, and stained as described

previously (Li et al., 2006) and examined with a JEM-1230 transmission

electron microscope (JEOL). For scanning electron microscopy observa-

tion, anthers were collected and processed essentially as described by

Keijzer et al. (1996) and observed with a JSM-6360LV scanning electron

microscope (JEOL). For preparing the free-hand sections, the materials

were fixed into a hole of a radish block. Transverse sections were then

madeby hand-sectioning. The sectionswere photographedusing aNikon

E600 microscope and a Nikon DXM1200 digital camera.

For DAPI staining to stain nuclei, samples were fixed overnight in FAA

(50% ethanol, 10% formalin, and 5% acetic acid). Then, the microspores

were squeezed out to the slide and stained with two to three drops of

DAPI stain solution (1.25 mg mL21 in 0.1*PBS). After being covered with a

cover glass, the nuclei were examined under the fluorescence micro-

scope immediately.

Soluble Sugar Assays by GC-MS and Starch Measurement

Metabolites were analyzed essentially as previously described (Lisec

et al., 2006). About fifty milligrams (fresh weight) of anther, lemma/palea,

or flag leaf were harvested and ground into a fine powder in liquid

nitrogen. Seven hundred microliters of methanol was immediately added

to the powder to stop enzymatic activity, and 50 mL of 0.2 ngmL21 rabitol

(Sigma-Aldrich) was then added. After centrifugation at 10,000g for 2min,

the supernatant was transferred to a new tube and dried for sugar assay,

and the remaining pellet was used to assay starch content using a starch

assay kit (product number SA20-1KT; Sigma-Aldrich). For methoxima-

tion, 40 mL of methoxyamine hydrochloride in pyridine (20 mgmL21) was

used at 308C for 90 min. Afterward, 40 mL of N-methyl-N-trimethylsilyl-

trifluoroacetamide was added, and themixture was incubated at 378C for

30 min. GC-MS analysis was performed using an Agilent 6890 series gas

chromatograph fitted with a capillary column (0.25 mm3 30 m, 0.25-mm

film thickness [HP-5MS]). The gas chromatograph was combined with a

quadrupole mass selective detector (Agilent). Samples (2 mL) were

injected at a 1:25 split ratio into a GC-MS system. The detector and

injector temperature were maintained at 2808C. The temperature pro-

gram was as follows: isothermal for 2 min at 708C, followed by a 58C per

min ramp to 1308C, then 108C per min ramp to 1808C, then 58C per min

ramp to 2858C and holding at this temperature for 10 min. Nitrogen was

used as the carrier gas at a flow rate of 20 mL min21.

Radiolabeling

According to the definition by Felker et al. (1984), the wild-type and csa

stems containing internodes I to IV, the flag leaf, and flowers were placed

and incubated in water containing 1 mCi [fructose-U-14C]sucrose

(21.8 GBq mmol21 in 9:1 ethanol:water; MP Biochemicals), and the

internode IV was submerged in the water directly. We detected the

amount of isotope using the middle regions of internodes I, II, and III.

The wild-type and csa panicles with;100 flowers were excised at stages

9, 11, and 13, and each panicle was transferred into 0.1 mL water

containing 1 mCi [fructose-U-14C]sucrose for 1, 6, and 12 h at room

temperature, using the panicle in water without 14C-labeled sucrose as

control. We then collected the lemma, palea, and anther for analysis. The

materials were incubated with 300 mL of 60% HClO4 and 600 mL of H2O2

at 558C for 15 h until the samples appeared clear. After cooling to room

temperature, 3 mL of scintillation fluid [6 g 2,5-diphenyloxazole (PPO),

0.075 g p-bis-2-(5-phenyloxazolyl)-benzene (POPPO), 250 mL Triton

X-100, and toluene to 1 liter] and 5 mL of 2-methoxyethanol were added

and mixed. The radioactivity (cpm) was measured by liquid scintillation

counting (Beckman LS650) (Eksittikul et al., 2001). Counting of isotope

signals was performed with at least three biological replicates.

Thin Layer Chromatography of 14C-Sugars

The 14C-sugars in the anther after a 1- and 12-h treatment were extracted

using 80% ethanol at 808C for 10 min and applied to silica gel 60 F254

plates (20 3 20 cm in size, 0.25 cm in layer thickness; Merck). The

chromatograms were performed using an ascending solvent system of

ethyl acetate:acetic acid:methanol:water (60:15:15:10 by volume). Sug-

ars were detected using 1,3-dihydroxynaphthalene (CAS #132-86-5; Alfa

Aesar) visualizing reagent (Ruan and Patrick, 1995). Standards of su-

crose, glucose, and fructose (Sigma-Aldrich) were used as the reference

control. The corresponding bands were scraped into the isotope tube,

and 2 mL ethanol and 3 mL of scintillation fluid were added and mixed.

The radioactivity (cpm) was measured using the similar conditions as

described above, and three biological replicates were tested.

Molecular Cloning of CSA

For mapping of the CSA locus, total DNA was isolated using the CTAB

(cetyl trimethyl ammonium bromide) method from rice leaves of each

selected plant. InDel (insertion-deletion) markers were developed ac-

cording to the sequence difference between the genome sequence of

Japonica Nipponpare and Indica 9311 (Li et al., 2006). Polymorphism

regions between the two rice subspecies were identified by aligning the

BAC clones sequences of Japonica and Indica, and the primers used for

gene mapping were designed based on the polymorphism regions. The

primers for molecular cloning of CSA are listed in Supplemental Table 4

online. The PCR products were separated on 6% polyacrylamide dena-

turing gels, and bands were visualized by silver staining (Xu et al., 2002).

Complementation of the csaMutant

For complementation, ;4.3-kb genomic DNA fragment containing the

entire CSA coding region, a 2151-bp upstream sequence, and a 525-bp

downstream sequence was amplified from BAC clone OSJNBb0058A16

(kindly provided by B. Han, National Center for Gene Research, Shanghai

Institutes for Biological Sciences, Chinese Academy of Sciences) with

primer MYBPF 59-GGATCCGCTATGCACCTAGACGAGTGTTGTC-39

and MYBR 59-GAATTCGTGACCACTGAGCAAGGAGTAGCTC-39; at-

tached restriction enzyme sites BamHI and EcoRI are underlined. The

amplified fragment was cloned into pMD18-T (TaKaRa), released by

BamHI-EcoRI digestion, and subcloned into BamHI-EcoRI–digested

binary vector pCAMBIA1301 (CAMBIA; hygromycin resistance). Then,

the calli induced using the homogenous csa young panicles, whichmainly

included palea and lemma, were used for transformation with Agro-

bacterium tumefaciens EHA105 carrying the p1301CSA plasmid as

previously described by Hiei et al. (1997).

CSA Nuclear Localization Analysis

The GFP cDNA was amplified from pBSK-GFP vector with the following

primers: 59-CCCGGGATGGGTAAAGGAGAAGAACTTTTCACTG-39 and

59-GAGCTCTTATTTGTATAGTTCATCCATGCCATGTG-39 (attached

CSA Regulates Sugar Partitioning in Rice Anther 685

restriction enzyme sites SacI and SmaI are underlined). The PCR product

was cloned into pMD18-T vector (TaKaRa) and was released by SacI-

SmaI digestion, then was subcloned into the SacI-PmacI–digested

pBI121 vector containing the cauliflower mosaic virus 35S promoter to

generate p121-GFP. The CSA cDNA (AK107461 provided by RGRC) was

amplified from the cDNA clone vector pCMVFL3 (RGRC-NIAS; http://

www.rgrc.dna.affrc.go.jp/stock.html) with primers (59-TCTAGAATGG-

CTCACGAGATGATGGGTG-39 and 59-CCCGGGTGTCGCGCCGACG-

CCGAGGAAG-39, attached restriction site is underlined). The amplified

fragment was digested with XbaI-SmaI and ligated with the same enzyme

digested p121-GFP to create p121-CSA-GFP. Transient expression of

the p121-CSA-GFP fusion and p121-GFP alone (as a control) in the onion

epidermis was performed as previously described (Collings et al., 2000)

using a helium biolistic device (Bio-Rad PDS-1000). The samples were

observed with a confocal laser microscope (Zeiss LSM510).

Phylogenetic Analysis

We constructed neighbor-joining (NJ) trees using the MEGA software

(version 3.1) (http://www.megasoftware.net/index.html) (Kumar et al.,

2004) with the following parameters: Poisson correction, pairwise dele-

tion, and bootstrap (1000 replicates; random seed). The MrBayes soft-

ware (Huelsenbeck and Ronquist, 2001; Ronquist and Huelsenbeck,

2003) was used to construct Bayesian trees after running for 106 gener-

ations, with four Markov chains, and sampled every 1000 generations.

The WAG model was used for amino acid substitutions (Whelan et al.,

2001), and invariable plus eight categories of g-distributed substitution

rateswere used to correct the among-site substitution rate heterogeneity.

NJ tree was shown with bootstrap values fromNJ and Bayesian posterior

probability, unless otherwise indicated. When only one value is shown, it

is the NJ bootstrap values.

RT-PCR and Quantitative PCR Assay

Total RNA was isolated from rice tissues (root shoot, leaf, lemma/palea,

and anthers) at different stages with the Trizol Reagent kit (Invitrogen)

according to themanufacturer’s protocol. The stages of rice anthers were

classified according to Zhang and Wilson (2009). After treatment with

DNase (Promega), the isolated 0.3 mg RNA was reverse transcribed to

synthesize first-strand cDNA using the ReverTra Ace-a-First-Strand

cDNA synthesis kit (TOYOBO). Three microliters of the reverse transcrip-

tion products were used as template in the following PCR reaction. PCR

was performed with TaKaRa Ex Taq DNA polymerase for 34 cycles of

denaturation for 40 s at 948C, annealing for 40 s at 588C, and extension for

1 min at 728C, followed by a final extension for 5 min. Quantitative PCR

analysis was performed using SYBR Premix EX Taq (TaKaRa) on a Rotor-

Gene RG3000A detection system (Corbett Research). All PCR experi-

ments were conducted in a reaction mixture containing 10 pmole each

primer and 3 mMmagnesium chloride, and 2 mL of the reverse transcrip-

tion products were used as template. Samples were denatured for 5 min

at 948C; followed by 40 cycles of 20 s of denaturation at 958C, 30 s of

annealing at 608C, and 30 s of elongation at 728C; followed lastly by one

cycle of 1 s of denaturation at 958C, 30 s of annealing at 658C, and 30 s of

denaturation at 958C. After the renaturation, the melting parameters were

assessed. Each experiment was repeated six times. Data acquisition and

analyses were performed using the method described by Roter-Gene

version 6.0 (Build 38) software. Samples were normalized using ACTIN1

expression; the relative expression levels weremeasured using the 2(2DCt)

analysis method.

In Situ Hybridization

Tissues of wild-type flowers at various developmental stages and roots

were fixed in 5% acetic acid, 50% ethanol, and 3.7% formaldehyde in

water for 16 h at 48C. After dehydration through an ethanol series, tissues

were embedded in Paraplast Plus (Oxford Labware) and sectioned at

8-mm thickness using an YL3-A rotary microtome (Shanghai Instrument

Factory). After sequence analysis, a 389-bp CSA cDNA fragment (577 to

965) and a 263-bpMST8 fragment (1384 to 1646) with less similarity with

other rice genes were amplified using RT-PCR as described above,

respectively. The PCR products were confirmed by sequencing and

cloned into pBluescript II KS+ phagemid vector (Stratagene) at the

BamHI-HindIII sites. Then, these segments were transcribed in vitro

under SP6 or T7 promoter with RNA polymerase using the DIG RNA

labeling kit (Roche). The mixture was prepared for the DIG-labeled RNA

antisense or sense probe. RNA hybridization and immunological detec-

tion of the hybridized probeswere performed according to the protocol of

Kouchi and Hata (1993). Images were obtained using the Olympus Nikon

E600 microscope.

Protein Expression and CSA Antibody Production

To produce the specific antibody for theChIP experiment, aCSA-specific

fragment (the coding sequence, from 487 to 795) was synthesized based

on the bacterial preferred codon usage (Qian et al., 2006) and cloned into

pET-32a vector (Novagen) to produce pET32a-CSA. The fusion protein

expression and purification were performed according to the manufac-

turer’s instructions, and antibody preparation in rabbit was performed as

described by Huang et al. (2003).

For the EMSA experiment, recombinant CSA protein was produced in

Escherichia coli using the full-length coding sequence of CSA synthe-

sized based on the bacterial preferred codon usage (Qian et al., 2006).

The synthesized CSA was cloned into pET30a vector (Novagen). Re-

combinant CSA protein was induced and affinity purified as above.

Protein Gel Blotting

Nuclei extracts were produced following the protocol that was used for

the ChIP experiments, except that the flower material was not fixed.

Proteins were separated on 12%SDS-PAGE gels and electroblotted onto

Hybond-C nitrocellulose membrane (RPN 303C; Amersham). Mem-

branes were blocked for 1 h with 5% BSA in PBS-Tween buffer (137

mM NaCl, 268 mM KCl, 47 mM KH2PO4, 8.1 mM Na2HPO4, and 0.05%

Tween, pH 7.4). Immunoprobing of CSA was conducted with the anti-

CSA polyclonal antibody for 2 h at room temperature at a dilution of 1:500

in PBS. Three washes of 5min eachwere performedwith PBS-Tween. An

anti-rabbit IgG conjugated with alkaline phosphatase was used as the

secondary antibody at 1:3000 dilution for 1 h at room temperature. Three

washes of 5 min each were performed with PBS-Tween, and target

proteins were visualized using 4-nitro blue tetrazoliu chloride (Roche) and

5-bromo-4-chloro-3-indolyl phosphate (Roche), according to the SABC

(streptavidin-biotin-peroxidase complex) method (Xing et al., 2009).

Histone H3 was used as a loading control for protein levels with Anti-

Histone H3 monoclonal antibody (Millipore) at a dilution of 1:1000.

ChIP and Quantitative PCR Analysis

The procedure for ChIP of CSA-DNA complexes in rice wild type was

modified from Haring et al. (2007). Rice spikelets at stages 9 and 11 were

fixed with formaldehyde under vacuum. Chromatin was isolated and

sonicated to produce DNA fragments shorter than 500 bp. Some

untreated sonicated chromatin was reversely cross-linked and used as

the total input DNA control. Immunoprecipitation with CSA-specific

immune antiserum and without any serum was performed as the refer-

ence above.

The amounts of genomic DNA immunoprecipitated were assayed by

real-time quantitative PCR using the same conditions as for quantitative

RT-PCR. For a PCR reaction, 0.5 mL of recovered DNA from ChIP or

686 The Plant Cell

controls or 1 mL of input DNA diluted 50-fold was added as template.

Each reaction was repeated four times. Quantification involved normal-

ization of each immune precipitation (IP) or control sample Ct to the input

DNA sample Ct to obtain a DCt (DCt IP or DCt Control), and then the

relative enrichment of each fragment was calculated using the following

equation: 22(DCt IP 2 DCt control). An unrelated DNA sequence from the rice

ACTIN1 gene was used as an internal control (Li et al., 2006). Primer

sequences used for the ChIP enrichment test are listed in Supplemental

Table 6 online.

EMSA

The DNA fragments containing the two CCAAT-boxes of the rice MST8

regulatory region, AGAAGC—CAACGG—CAACGG—TGGTC, were gen-

erated using PCR amplification with the following specific primers:

EMSAF (59-AGAAGCCAGCCTTGCGTCCAT-39) and EMSAR (59-GAC-

CAACTAATTATTTATCCG-39). The DNA fragment was cloned into

pMD18-T vector (TaKaRa) for sequence confirmation. Then, the fragment

was labeled with DIG-labeled kit (DDLK-010) using the specific primers.

The DNA binding reactions were performed according to Wang et al.

(2002) with the following modifications. Reaction components were

incubated in binding buffer [10 mM Tris-HCl, pH 7.5, 50 mM NaCl,

1 mM EDTA, 5% glycerol, 0.05 mg mL21 poly(dI-dC), and 0.1 mg mL21

BSA] at room temperature for 20 min. The entire reaction mixture was

analyzed on a 5% PAGE gel. After drying the gel, DIG-labeled DNA

fragments were detected.

Accession Numbers

Sequence data from this article for the mRNA and genomic DNA of CSA

can be found in the GenBank/EMBL data libraries under accession

numbers NM_001049255 and NC_008394, respectively. GenBank

accession numbers of all genes used in this study are AY822464

(MST8), AY220486 (INV4), AF419298 (SUT3), AF249880 (UGP2),

DQ395328 (UGP1), X62134 (GBSS1), X16280 (ACTIN1), NP197282 (At

MYB56), NP177115 (At MYB105), NP564261 (At MYB117), BAF23796

(Os08g33800), and CAA67000 (Os GAMYB).

Supplemental Data

The following materials are available in the online version of this article.

Supplemental Figure 1. Scanning Electron Microscopy Observations

of the Wild-Type and csa Anthers at Stage 13.

Supplemental Figure 2. DAPI Staining Showing Microspore Devel-

opment of the Wild Type and csa.

Supplemental Figure 3. Transmission Electron Micrographs of the

Anthers from the Wild Type and csa.

Supplemental Figure 4. 14C-Signal Accumulation in the Anther and

Lemma/Palea of the Wild Type and csa after 1, 6, and 12 h of

Treatment of [14C]Sucrose.

Supplemental Figure 5. Nucleotide and Amino Acid Sequences

of CSA.

Supplemental Figure 6. Sequence Alignment of CSA and Its Close

Homologs.

Supplemental Figure 7. Analysis of GUS Activity in the pCSA-GUS

Line.

Supplemental Figure 8. In Situ Analysis of the MST8 Expression in

the Anther.

Supplemental Figure 9. Protein Gel Blot Analysis of CSA in the

Nuclear Protein Extracts of Wild-Type and csa Flowers.

Supplemental Figure 10. Quantitative PCR Analyses of Relative

mRNA Levels of Genes Involved in Sugar Partitioning in the Anther

and Lemma/Palea.

Supplemental Table 1. Sugar and Starch Level Profiles in the Wild

Type and the csa Mutant.

Supplemental Table 2. 14C-Signal Accumulation in the Wild-Type

and csa Mutant Lemma/Palea, Anther, and Stem.

Supplemental Table 3. Accumulation of [14C]Sucrose and [14C]

Hexose in the Wild-Type and csa Anthers at Stage 13 after 1- and

12-h Treatment.

Supplemental Table 4. List of the Primers Used for Mapping and RT-

qPCR Analyses.

Supplemental Table 5. Sucrose and Starch Levels in the Flag Leaf of

Complemented csa Lines at Stage 13 of Anther Development.

Supplemental Table 6. List of the Primers Used for ChIP Enrichment

Assay.

Supplemental Data Set 1. Text File of the Alignment Used for the

Phylogenetic Analysis Shown in Figure 6I.

ACKOWLEDGEMENTS

We thank the anonymous reviewers for very helpful comments and B.

Han and the RGRC for providing BAC clone and cDNA clone, respec-

tively. We thank Z.-J. Luo and M.-J. Chen for mutant screening and

generation of F2 populations, X.-Y. Gao for plastic sections, scanning

electron microscopy, and transmission electron microscopy and X.-Y.

Chen for isotope detection. We also thank Y.-M. Liu, W. Jia and Y.-P.

Qiu for GC-MS assay. D.-M. Braun, Y. Zhang, and D. Werck-Reichhart

are gratefully acknowledged for their valuable suggestions and editing

on this manuscript. This work was supported by the Funds from the

National Basic Research Program of China (2009CB941500), the Na-

tional “863” High-Tech Project (2006AA10A102 and 2007AA10Z112),

the National Natural Science Foundation of China (30725022, 30830014,

and 90717109), the Chinese Transgenic Project (2009ZX08009-108B),

and the Shanghai Leading Academic Discipline Project (B205).

Received December 21, 2009; revised February 14, 2010; accepted

March 2, 2010; published March 19, 2010.

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DOI 10.1105/tpc.109.073668; originally published online March 19, 2010; 2010;22;672-689Plant Cell

Hui Zhang, Wanqi Liang, Xijia Yang, Xue Luo, Ning Jiang, Hong Ma and Dabing ZhangRequired for Rice Pollen Development

Encodes a MYB Domain Protein That Regulates Sugar PartitioningCarbon Starved Anther

 This information is current as of July 26, 2018

 

Supplemental Data /content/suppl/2010/03/15/tpc.109.073668.DC1.html

References /content/22/3/672.full.html#ref-list-1

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