Acetylesterase-Mediated Deacetylation of Pectin Impairs CellElongation, Pollen Germination, and Plant Reproduction C W

Jin-Ying Gou,a Lisa M. Miller,b Guichuan Hou,c Xiao-Hong Yu,a Xiao-Ya Chen,d and Chang-Jun Liua,1

a Biology Department, Brookhaven National Laboratory, Upton, New York 11973b National Synchrotron Light Source, Brookhaven National Laboratory, Upton, New York 11973c Appalachian State University, Boone, North Carolina 28608-2027d National Key Laboratory of Plant Molecular Genetics, Institute of Plant Physiology and Ecology, Shanghai Institutes for

Biological Sciences, Shanghai 200032, China

Pectin is a major component of the primary cell wall of higher plants. Some galacturonyl residues in the backbone of

pectinaceous polysaccharides are often O-acetylated at the C-2 or C-3 position, and the resulting acetylesters change

dynamically during the growth and development of plants. The processes involve both enzymatic acetylation and

deacetylation. Through genomic sequence analysis, we identified a pectin acetylesterase (PAE1) from black cotton-

wood (Populus trichocarpa). Recombinant Pt PAE1 exhibited preferential activity in releasing the acetate moiety from

sugar beet (Beta vulgaris) and potato (Solanum tuberosum) pectin in vitro. Overexpressing Pt PAE1 in tobacco

(Nicotiana tabacum) decreased the level of acetyl esters of pectin but not of xylan. Deacetylation engendered

differential changes in the composition and/or structure of cell wall polysaccharides that subsequently impaired the

cellular elongation of floral styles and filaments, the germination of pollen grains, and the growth of pollen tubes.

Consequently, plants overexpressing PAE1 exhibited severe male sterility. Furthermore, in contrast to the conventional

view, PAE1-mediated deacetylation substantially lowered the digestibility of pectin. Our data suggest that pectin

acetylesterase functions as an important structural regulator in planta by modulating the precise status of pectin

acetylation to affect the remodeling and physiochemical properties of the cell wall’s polysaccharides, thereby affecting

cell extensibility.

INTRODUCTION

Theprimary cell walls of plants are rigid yet dynamically organized

networks comprising a mixture of polysaccharides and structural

proteins, of which pectin is a major component (Carpita and

McCann, 2000; Ridley et al., 2001; Mohnen, 2008). The pectina-

ceous polysaccharides consist of different structural domains,

including the linear homogalacturonan (HG) regions of up to 200

residues of (1,4)-linkeda-D-galacturonic acid (GalA) and the highly

branched “hairy” regions, including rhamnogalacturonan I (RG-I),

which encompasses the repeating disaccharide units of a-(1,2)-L-

Rha-a-(1,4)-D-GalA, the highly substituted galacturonan RG-II

domain, and the xylogalacturonan domain (Carpita and McCann,

2000; Mohnen, 2008). Some galacturonyl residues in the back-

bone of pectinaceous polysaccharides are often O-acetylated at

the C-2 or C-3 hydroxyl, or methylesterified at the C-6 carboxyl,

although the distribution of methyl- and acetylesters between HG

and RG is still unclear (Kouwijzer et al., 1996; Perrone et al., 2002;

Ralet et al., 2008). In the pectins of bamboo shoot (Phyllostachys

edulis), flax (Linum usitatissimum), potato (Solanum tuberosum),

cotton (Gossypium hirsutum), carrot (Daucus carota), tobacco

(Nicotiana tabacum), and tomato (Solanum lycopersicum), acetyl

groups reportedly are highly abundant in the RG-I domain

(Komalavilas and Mort, 1989; Ishii, 1995, 1997; Schols and

Voragen, 1996), whereas in the pectin of sugar beet (Beta

vulgaris),;75%of the acetyl groups belong to the HGbackbone

(Kouwijzer et al., 1996; Ralet et al., 2005).

Presumably, pectins are synthesized in the cis-Golgi, methyl-

esterified in the medial- Golgi, substituted in the trans-Golgi, and

then secreted into the cell wall in a highly esterified state (Micheli,

2001; Sterling et al., 2001; Mohnen, 2008). The function of pectin

methylesterification has been explored extensively. Both its

extent and distribution are found to be important for determining

pectin’s physiochemical and biological properties (Peaucelle

et al., 2008; Wolf et al., 2009). The enzymes responsible for

regulatingpectinmethylesters are alsowell characterized (Tieman

et al., 1992; Wen et al., 1999; Bosch and Hepler, 2005; Pelloux

et al., 2007; Pelletier et al., 2010). By contrast, studies on the

occurrence and functions of pectin acetylation are scarce (Pauly

and Scheller, 2000; Manabe et al., 2011). Purportedly, pectin is

acetylated between the Golgi and the cell wall during its exocy-

tosis; then, the exported acetylated pectin is incorporated into cell

walls (Liners et al., 1994; Pauly and Scheller, 2000; Scheller and

Ulvskov, 2010). The extent and distribution of acetylesters vary

according to the source of pectin (Schols and Voragen, 1994).

1 Address correspondence to cliu@bnl.gov.The author responsible for distribution of materials integral to the findingspresented in this article in accordance with the policy described in theInstructions for Authors (www.plantcell.org) is: Chang-Jun Liu (cliu@bnl.gov).CSome figures in this article are displayed in color online but in blackand white in the print edition.WOnline version contains Web-only data.www.plantcell.org/cgi/doi/10.1105/tpc.111.092411

The Plant Cell, Vol. 24: 50–65, January 2012, www.plantcell.org ã 2012 American Society of Plant Biologists. All rights reserved.

Pectins from strawberry (Fragaria ananassa) and sugar beet

contain a sizeable portion of acetyl groups, viz., 1.2 and 2.5%

(w/w), respectively, equivalent to 5 and 10% acetylation of the

available galacturonyl residues, while citrus fruit and apple

(Malus domestica) pectins have fewer acetylesters, ;0.25 and

0.4%, respectively (Voragen et al., 1986). Moreover, the degree

of O-acetylation of pectin changes during the growth and differ-

entiation of plant tissues and in response to the environmental

conditions (Liners et al., 1994; Vercauteren et al., 2002;Gou et al.,

2008). The acetate content in the cell wall of juvenile leaves of

black cottonwood (Populus trichocarpa) is much higher than that

in mature and senescent leaves (Gou et al., 2008). The change in

acetylation during development points to an in muro dynamic

structural modification that might be important for the organiza-

tion, metabolism, and function of polysaccharides in planta.

However, there is limited informationon the enzymatic processes

of both the acetylation and deacetylation of polysaccharides and

their biological effects on plant growth and development.

Earlier investigations suggested pectin acetylesters might be

one crucial structural factor in regulating the polymer’s biophys-

ical properties (Liners et al., 1994). For example, like methylester

status, the degree of acetylation affected the gel formation of

extracted pectins. Partial hydrolysis of the acetyl esters in beet

pectins improved their gelation properties (Pippen et al., 1950;

Williamson et al., 1990; Ralet et al., 2003). Deacetylation also

increased pectin’s solubility in water by lowering the hydropho-

bicity of the polysaccharide backbone (Rombouts and Thibault,

1986). On the other hand, acetylesters demonstrably protect

polysaccharides against enzymatic digestion (Biely et al., 1986;

Schols and Voragen, 1994; Chen and Mart, 1996; Benen et al.,

1999; Bonnin et al., 2003). Thus, removing those moieties either

with digestive enzymes or by chemical saponification in vitro

seems to be aprerequisite for degrading the polysaccharide core

(Kauppinen et al., 1995).

The synthesis and dynamics of cell wall acetylesters involve at

least two types of enzymes, acetyltransferases, which transacy-

late the sugar residues of polymers (Pauly and Scheller, 2000;

Gille et al., 2011; Lee et al., 2011; Manabe et al., 2011), and

acetylesterases, which cleave the ester bond between a glycosyl

carbon and an acetyl group, thus releasing acetate frommodified

polysaccharides (Williamson, 1991; Bordenave et al., 1995). Pec-

tin acetylesterase (EC 3.1.1.6; PAE) belongs to CAZy class 12 and

13 of the CE family (http://www.cazy.org/). The enzymes were

isolated from bacteria (Shevchik and Hugouvieux-Cotte-Pattat,

1997, 2003), fungi (Kauppinen et al., 1995; Bonnin et al., 2008),

and plants (Williamson, 1991; Bordenave et al., 1995; Christensen

et al., 1996). Two different PAEs were purified from orange (Citrus

sinensis) peel, one with amolecular mass (MW) of 29 to 30 kD and

a pI of 5.1 (Williamson, 1991), the other with a molecular mass of

42 kDand a pI > 9 (Christensen et al., 1996). The PAE purified from

the cell walls of mung bean (Vigna radiata) hypocotyls had

physicochemical properties (molecular mass and pI) similar to

those of the higher molecular mass acetylesterase from orange

(Bordenave et al., 1995). The PAEs from both species exhibited

the highest activity on synthetic substrates, such as triacetin and

p-nitrophenyl acetate, and were active on the native pectin of

sugar beet and flax (Williamson, 1991; Bordenave et al., 1995;

Christensen et al., 1996). A cDNA encoding a putative PAE was

isolated from mung bean seedlings, based on the peptide se-

quences of the purified protein (Breton et al., 1996). However, no

further biochemical and biological analyses were made to ascer-

tain its functions.

Approximately 10 putative PAE genes were deduced in the

P. trichocarpa genome (Geisler-Lee et al., 2006); however, none

are functionally characterized. In this study, we characterized

one of those homologous genes. Expressing this gene in planta

reduced the content of acetylester primarily in pectinaceous

polysaccharides and, consequently, resulted in pleiotropic ef-

fects on the plant’s growth, development, and reproduction.

RESULTS

Gene Expression Analysis of Putative PAEs from

P. trichocarpa

Previous analyses of genomic sequences in Populus recognized

a set of putative carbohydrate esterase (CE) genes encoding

pectin acetyltransferases (Geisler-Lee et al., 2006). To explore

their functions, we retrieved the gene sequences from the

P. trichocarpa genome. Sequence alignment and phylogenetic

analysis showed that nine putative Populus CEs (excluding one

partial sequence), together with 12 putative acetylesterase ho-

mologs from Arabidopsis thaliana (http://www.cazy.org/), clus-

tered into three different clades. All plant esterase genes are

distinct from those of bacteria and fungi. Among them, CE13_4,

5, and 7 are grouped with the PAE cDNA cloned frommung bean

(Breton et al., 1996) and separate from the others (Figure 1; see

Supplemental Data Set 1 online).

These P. trichocarpa putative CE genes displayed distinct

tissue-specific (see Supplemental Figures 1A to 1H online) and

temporal (see Supplemental Figures 1I to 1P online) expression

patterns, suggesting their potential distinct physiological func-

tions. Among them, CE13_5 was highly expressed in young

leaves and in the early developing nodes (see Supplemental

Figures 1C and 1K online). Its homologous cDNA probe in

Populus balsamifera was also highly expressed in female and

male catkins, in addition to young leaves and seedlings, as found

in a developmental tissue transcriptomic analysis (Wilkins et al.,

2009; see Supplemental Figure 2 online). These expression

patterns imply that CE13_5may be involved inmodifying primary

cell walls in fast growing tissues. Accordingly, we cloned this

gene for further functional analysis.

Recombinant Pt CE13_5 Exhibits Acetylesterase Activity

in Vitro

Using RT-PCR, we isolated cDNA encoding CE13_5 from the

total RNAs of the leaves and stems of P. trichocarpa. The cDNA

clone (accession number HQ223420) contains 1185 nucleotides

and encodes a polypeptide of 394 amino acid residues, with a

deduced molecular mass of 42.8 kD and pI of 6.6.

The amino acid residues of CE13_5 share;62% identity with

the PAE from mung bean (Breton et al., 1996); both have a

conserved GXSXG motif, characteristic of the Ser hydrolase

superfamily. The encoded sequence of Pt CE13_5 differs from

the bacterial PAEs, PaeY, PaeX, and YxiM (;2 to ;10%

Acetylesterase Modulates Plant Growth 51

identities), and from the RG acetylesterase of Aspergillus acu-

leatus (;6% identity) (Figure 1), pointing to the distinct evolu-

tionary origins of the polysaccharide acetylesterases.

To characterize the biochemical function of Pt CE13_5, we

expressed the gene in Escherichia coli strain Rosetta (DE3)

pLysS, a heterologous expression system with a tightly con-

trolled level of protein expression. Although the recombinant

protein was severely aggregated and sequestered into inclusion

bodies, we recovered and purified a small portion of the soluble

recombinant protein from the transgenic E. coli.

To examine its potential esterase activity on polysaccharides,

we chemically derived the acetylated polysaccharides (Gou

et al., 2008). The recombinant protein showed high activity in

releasing acetyl moieties from acetylated pectate (polygalactur-

onan), xylan, and arabinogalactan, with the highest activity on the

first one (see Supplemental Figure 3 online). Like the acetyles-

terases purified from plant species, the recombinant enzyme

also displayed activity on the synthetic chemical 2-naphthol

acetate (see Supplemental Figure 3 online).

Since chemical derivatizationmay entail the hyperacetylation of

polysaccharides, andsomight not represent thenativeacetylation

Figure 2. Enzymatic Activity of Recombinant Pt PAE1.

(A) Acetylesterase activity of recombinant PAE1 with different natural

carbohydrate polymers.

(B)Methylesters released by 2 mg pectin esterase (as positive control), Pt

PAE1, and buffer (as negative control [Neg. Ctrl.]) within 15 min at 358C.

(C) Ferulic acid released by alkaline treatment (2 N NaOH as positive

control), 2 mg Pt PAE1, or buffer as negative control in 15 min at 358C.

Data represent mean of triplicate samples. The error bar represents SE.

Figure 1. Phylogenetic Analysis of Polysaccharide Acetylesterases.

The neighbor-joining tree was constructed from the aligned full-length

amino acid sequences of putative carbohydrate acetylesterases from P.

trichocarpa and Arabidopsis. The amino acid sequences of the following

enzymes were also included: CAA67728 (Vr PAE, from V. radiata),

AJ507215 (PaeX, from Erwinia chrysantemi), Y09828 (PaeY, from E.

chrysantemi), BAA11692 (YxiM, from Bacillus subtilis), and CAA61858

(RAGA1, from A. aculeatus). Bar represents the output distance as

number of substitutions per site (i.e., 0.2 substitutions per site).

52 The Plant Cell

pattern of cell wall polymers, we incubated Pt CE13_5 recombi-

nant proteins with polysaccharides prepared from natural re-

sources. These included the RG-I–enriched pectin from potato

(Ishii, 1997), the HG-enriched pectin from sugar beet (Quemener

et al., 2003; Ralet et al., 2005), the birch wood (Betula spp) xylan,

and a xylan/hemicellulose preparation from cell walls of tobacco

leaves. We employed a mild alkaline treatment to calibrate the

acetylester content in those polymers used for substrates (see

Supplemental Figure 4 online). As depicted in Figure 2A, the

recombinant protein showed high activity in releasing acetyl

moieties from sugar beet and potato pectins but substantially

lower activity with birch wood xylan and the tobacco xylan/

hemicellulosepreparation. Although the limited solubility of natural

polymers (pectin and xylan) prevented a detailed kinetic analysis,

determining the kinetic parameters using chemically acetylated

polymers revealed that the recombinant Pt CE13_5 preferentially

reacts with acetylated pectate over xylan (Table 1).

To examine whether the recombinant enzyme has promiscu-

ous methyl- or feruloyl- esterase activity, we incubated the

enzymewith the highly methylesterified pectins from citrus fruits,

potato, and sugar beet. We found that the Pt CE13_5 recombi-

nant protein essentially showed no activity in releasing methyl-

esters from all of those natural polymers compared with the

pectin methylesterase (Figure 2B). Similarly, we did not detect

any activity in removing the feruloyl moiety from the incubated

sugar beet pectin, although a certain amount of ferulate is

released from this polymer by mild alkaline treatment (Figure

2C). We thus designated Pt CE13_5 as P. trichocarpa pectin

acetylesterase 1 (Pt PAE1).

Overexpression of PAE1 Reduces Acetyl Content in the

Pectin of Transgenic Plants

To investigate the biological function of Pt PAE1 in planta, we

expressed its full-length cDNA in tobacco (Figure 3A). Over-

expression of PAE altered the morphology of the tobacco plants

in all of the independent transgenic lines. During vegetative

growth, the primary apical buds of the Pt PAE1 transgenic plants

frequently wilted, followed by the emergence of new primary and

lateral buds (Figure 3B; see Supplemental Figure 5 online). The

expanded leaves of the transgenic plants exhibited a wavy

surface with curled edges, which differed from those of the wild

type (Figure 3C; see Supplemental Figure 5 online).

We fused Pt PAE1 at the C terminus of green fluorescent

protein (GFP) and stably expressed this construct in tobacco,

driven by the constitutive 35S cauliflower mosaic virus promoter.

The fluorescence signals appeared primarily in the contours of

the etiolated hypocotyl cells, in good agreement with propidium

iodide staining, a convenient histological marker for monitoring

the plant cell wall polysaccharides (McKenna et al., 2009) and in

sharp contrast with that of free GFP (Figures 3D to 3G), indicating

that PAE1 localized to cell walls.

Subsequently, we extracted individually the water- and acid-

soluble pectins from the expanded young leaves of both trans-

genic and control tobacco plants (see Supplemental Figure 6

online) and isolated xylan from the xylem tissues via xylanase

digestion. We readily detected acetyl esters in both the pectin

and xylan fractions from the transgenic and control tobacco

plants. However, their levels in both water- and acid-soluble

pectins of most transgenic plants were significantly lower than in

those of the control plants by about a 13 To ;42% and 8 to

;11%, respectively (Figures 4A and 4B). By contrast, the levels

of acetyl moieties in xylanase-digested fractions showed either

Figure 3. Overexpression of Pt PAE1 in Tobacco.

(A) RT-PCR examination of Pt PAE1 expression in transgenic tobacco

lines. Full-length PAE1 cDNA was detected in the transgenic lines but not

in the controls. V, transgenic vector (as positive control). The data

represent the result from one of three replicates.

(B) and (C) Vegetative development of transgenic (B) and control (C)

tobacco. Note that a couple of primary shoot apexes were apparent in

the transgenic line, Pt PAE1_6 (arrows in [B]) but that only one was

present in the control plant (arrow in [C]).

(D) to (F) Subcellular localization of Pt PAE1.

(D) Green fluorescence signal of GFP-PAE1 fusion monitored in the cells

of 5-d-old dark-grown transgenic seedling.

(E) The cells stained with propidium iodide.

(F) The merged image of (D) and (E).

(G) Green fluorescence signal of the free GFP. Bars = 2 mm.

Table 1. Kinetics of Recombinant Pt CE13-5

Substrate Vmax (mmole Acetate/mg/min) Km (mg/mL)a

Acetylated pectate 5.8 6 0.5 6.3 6 1.3

Acetylated xylan 5.1 6 1.3 25.3 6 8.6

aBecause the polymers lacked precise molecular mass, Km was not

expressed in units of molar concentration.

Acetylesterase Modulates Plant Growth 53

no change or even some increase in transgenic plants (Figure

4C). Meanwhile, the methylester content of pectin from trans-

genic tobacco leaves was notably increased, indicating potential

metabolic compensation or carbon flux redirection (see Supple-

mental Figure 7 online).

These data suggest that Pt PAE1 encodes an active esterase

that predominantly removes acetyl moieties from pectin in

vivo.

Overexpression of Pt PAE1 Affects the Development of the

Floral Tissues

As the transgenic tobacco plants approached reproductive

growth, we found a number of developing or open flowers that

had wilted or had fused petals (see Supplemental Figure 5

Figure 5. Development of Floral Tissues in Tobacco Plants Overex-

pressing Pt PAE1.

(A) Style and filaments in the developing flowers (at stage 12) of the

control and transgenic (Pt PAE1) plants.

(B) The lengths of epidermal cells of the style and filament at the apical,

middle, and basal parts of both tissues of independent transgenic lines.

Data represent the average of 40 cells. Error bar denotes the SD.

Asterisks indicate a statistically significant change (P < 0.05) under

Student’s t test.

(C) to (F) Positioning of stigma and anthers in the flower of the wild-type

control ([C] and [D]) and in the overexpression plants ([E] and [F]) at

flowering stages 11 ([C] and [E]) and 12 ([D] and [F]). Bar = 5 mm.

[See online article for color version of this figure.]

Figure 4. Cell Wall Acetylester Content of Pt PAE1 Overexpression

Plants.

(A) and (B) Acetylester content of the water- and acid-soluble pectins

from young leaves of the transgenic and control lines.

(C) Acetylester content of xylan from xylem cell walls. Data represent

mean of triplicate samples. Error bar represents SD. Asterisks indicate a

statistically significant change (P < 0.05) under Student’s t test.

54 The Plant Cell

online). The lengths of the styles and filaments in the transgenic

flowers were shorter than those in the flowers of the control

plants (Figure 5A; see Supplemental Figure 8A online), indicating

that the expression of Pt PAE1 retarded the elongation of both

pistil and stamen tissues. Calculating the lengths of the epider-

mal cells at different positions of the stylar tract and filaments in

flowers at flowering stage 12 (Drews et al., 1992) revealed that all

epidermal cells of the filament and the cells of the basal and

middle stylar tract of independent transgenic lines were signif-

icantly shorter than those of the control (Figure 5B), indicating

that the expression of PAE1 depresses cell elongation. Conse-

quently, the relative lengths of the stigma and anthers in

Figure 6. Cell Wall Esters of the Style and Filament.

(A) to (H) Optical and FTIR images of the basal tissues of the style and middle portion of the filament. The false-color images represent the ratio of ester

to polysaccharide in the style (A) and filament (E) of control plants and the Pt PAE1 overexpression ([C] and [G]) plants. The corresponding optical

images of the tissues are shown in (B), (D), (F), and (H).

(I) The representative FTIR spectra of the style, filament, and their corresponding control tissues. Each averaged spectrumwas normalized with the area

under the polysaccharide peaks so that the sizes of the ester peaks (at 1740 cm�1, arrow) were easily compared. Tg, transgenic.

(J) Content of pectin acetyl ester in the style and filament. Asterisks indicate a statistically significant change (P < 0.05) under Student’s t test. Data

represent mean of triplicate experiments. Error bar represents SE.

Acetylesterase Modulates Plant Growth 55

transgenic flower were greatly altered. In control flowers, four of

five anthers were at nearly the same height as the stigma or

slightly above it (Figures 5C and 5D), whereas, in transgenic

flowers, from flowering stage 11, the positions of all five anthers

were lower than the stigma (Figure 5E), and this morphological

change became more pronounced as flowering reached stage

12 (Figure 5F). The ratio of filament to pistil in transgenic flowers

was markedly less than that of the controls (see Supplemental

Figure 8Bonline), confirming the severe impairment of elongation

of the filament cells in the transgenic plants.

To examine the potential changes of the cell wall composition,

particularly of the acetylesters of the style and filaments, we

collected these tissues from the transgenic and control tobacco

plants at flowering stage 12 and imaged them via high-resolution

Fourier transform infrared microspectroscopy (FTIR) (McCann

et al., 1992). We calculated the ratio of the detected ester signal

(integrated area from 1725 to 1775 cm21; linear baseline from

900 to 1775 cm21) to that of the polysaccharides (900 to 1180

cm21; linear baseline from 900 to 1180 cm21) to generate a false-

color image on transverse sections of these organs. As depicted

in Figures 6A and 6B, in a transverse section of the style of control

plants, the epidermal cells of the transmitting tract exhibited a

high ratio of esters to polysaccharides. By contrast, the ratio in

the transgenic style was much decreased (Figures 6C and 6D).

We observed similar patterns in the cell walls of filament cells of

the control and transgenic plants (Figures 6E to 6G), denoting a

diminution of total esters in the cell walls of those transgenic floral

tissues.

To further ascertain the changes in esters in those transgenic

tissues, we extracted pectins from styles and filaments and

measured the contents of both acetyl- and methylester. We

detected a significant reduction in acetylesters, calculated on the

basis of equal amounts of pectin (measured for uronic acid), in

the pectinaceous fractions of both the filaments and styles in

transgenic plants compared with the controls (Figure 6J). How-

ever, there was no statistically significant difference in their

methylester content, although the amount varied considerably in

different lines (see Supplemental Figure 9 online). These data

show that expressing PAE1 reduced pectin acetylation in fast-

growing floral tissues.

To further analyze the effects of PAE1 expression on the com-

positionof thecellwalls of the style andfilament,wedepolymerized

their cell wall polysaccharides andmeasured the content of neutral

and acidic sugars, respectively, by gas chromatography–mass

spectrometry and a colorimetric method. Uronic acid decreased

significantly in the cell walls of both tissues in transgenic plants,

indicating a change in thepectinmatrix polymeror its extractability;

meanwhile, we observed substantial increases in the hemicellulo-

sic and cellulosic sugars, such as Gal, Xyl, andGlc in the filaments,

and of Man and Gal in the style cell walls in three of four examined

transgenic lines (Table 2). These data imply that PAE1-mediated

deacetylation differentially affects the biogenesis and remodeling

of the entire spectrum of cell wall polysaccharides in those fast-

growing tissues.

Overexpression of Pt PAE1 Affects the Development of

Pollen Grains

The expression of PAE1 also impaired the formation and devel-

opment of pollen grains within the anthers. At flowering stage 12,

the freshly dehiscent anthers of the control plants were fully filled

with dehydrated mature pollen grains (Figure 7A); by contrast,

there were fewer grains within the pollen sacs of transgenic

plants (Figures 7B to 7D).

We then chemically fixed grains from both control and trans-

genic plants and observed them under a scanning electron

microscope. The pollen grains from the former had a uniform

spherical shape and a regular exine pattern (Figures 7E and 7G),

and the germination apertures were readily apparent (Figure 7G).

By contrast, many pollen grains from transgenic plants had

collapsed; the exine of cell walls displayed an irregular pattern,

and the germination pores on the surface of the pollen grains had

essentially disappeared (Figures 7F and 7H). We then observed

these pollen grains by transmission electron microscopy. While

Table 2. Cell Wall Composition of Filament and Style

Filament Pt PAE1_2 Pt PAE1_4 Pt PAE1_5 Pt PAE1_6 Control

Rha 1.79 6 0.33 2.37 6 0.16 2.88 6 0.03 1.90 6 0.82 1.93 6 0.03

Ara 2.24 6 0.39 2.93 6 0.24 3.78 6 0.05 3.85 6 0.76 2.27 6 0.08

Xyl 8.87 6 1.97 14.05 6 1.17** 14.85 6 0.67** 19.33 6 3.39** 9.79 6 0.19

Man 1.17 6 0.06 1.36 6 0.03 1.85 6 0.05 1.55 6 0.10 1.32 6 0.13

Glc 94.93 6 8.27** 96.81 6 3.65** 105.34 6 0.14** 101.19 6 6.30** 78.78 6 3.47

Gal 12.82 6 1.19* 13.45 6 3.65** 16.71 6 0.14** 17.42 6 1.63** 10.15 6 3.47

Uronic acid 68.61 6 20.06* 119.60 6 12.00* 110.54 6 8.91* 122.91 6 27.14 127.49 6 14.20

Style Pt PAE1_2 Pt PAE1_4 Pt PAE1_5 Pt PAE1_6 Control

Rha 1.05 6 0.06 1.45 6 0.09 1.97 6 0.11 1. 56 6 0.11 1.39 6 0.12

Fuc 4.32 6 0.35 5.84 6 0.21 8.30 6 1.18 6.94 6 0.92 6.33 6 0.06

Xyl 9.21 6 1.17* 10.15 6 0.43** 15.44 6 2.18 14.06 6 2.04 12.17 6 0.10

Man 1.46 6 0.06** 1.19 6 0.08* 1.56 6 0.08** 1.22 6 0.06* 1.04 6 0.03

Glc 90.61 6 3.95** 109.54 6 3.01** 130.04 6 10.09 134.73 6 15.23 129.34 6 3.37

Gal 15.06 6 0.79** 21.27 6 0.90** 27.2 6 2.71** 24.84 6 2.96** 17.83 6 0.45

Uronic acid 54.60 6 7.63* 59.89 6 1.80* 60.52 6 2.66* 52.36 6 3.22* 75.72 6 2.27

The data represent mean (mg/g dry cell wall) of triplicate samples6 SE. *t test, P value < 0.05; **t test, P value < 0.01. The t tests were the comparisons

between individual transformants and the control line.

56 The Plant Cell

the exine structure essentially remained in the normal shaped

pollen grains of transgenic plants, the rod-like bacula (columella),

tectum, and triphine of the exine had disappeared from the

surface of the collapsed grains. Moreover, the intine and under-

lying electron-translucent layer, likely representing the cellulose

microfibrils, of transgenic pollen grains appeared to be disorga-

nized and looser than those of the controls (Figures 7I to 7K).

These images suggested that expressing PAE1 altered struc-

tures of the pollen wall.

To examine the potential compositional changes in the cell walls

of pollen grains, we analyzed transgenic and controlmature pollen

grains at flowering stage 12 via FTIR microspectroscopy after

ethanol extraction. As Figure 8A depicts, the characteristic ester

bond vibration at 1740 cm21 of FTIR spectra (McCann et al., 2007)

is less in the pollen from the transgenic lines. The ratio of the ester-

bond signal to the total signals of polysaccharides ranged from

Figure 7. Overexpression of Pt PAE1 Affects Pollen Grain Development

in Tobacco.

(A) to (D) Mature pollen grains in the pollen sacs of the control (A) and

transgenic ([B] to [D]) plants at flowering stage 12.

(E) to (H) Scanning electron microscopic images of the pollen grains of

the control ([E] and [G]) and transgenic ([F] and [H]) plants.

(I) to (K) Transmission electron microscope images of the control (I),

transgenic normal (J), and transgenic collapsed (K) pollen grains. Arrows

point to the changes of the intine layer and the putative cellulose

microfibrils.

Bars = 2.5 mm in (A) to (D) 50 mm in (E) and (F), 20 mm in (G) and (H), and

1 mm in (I) to (K).

[See online article for color version of this figure.]

Figure 8. Chemical Composition of Mature Pollen Grains.

(A) Average FTIR spectra of pollen grains from the transgenic (dotted

lines) and control (solid line) plants. Each spectrum represents the

average of 20 samples. The arrow indicates the peak of the acylester

group.

(B) Acetic acid content of pollen grains isolated from transgenic and

control plants. Asterisks indicate statistically significant changes under

Student’s t test (P < 0.01). Data represent the mean of triplicate samples.

Error bar represents SE.

Acetylesterase Modulates Plant Growth 57

0.05 to 0.06 in transgenic plants, values significantly lower (t test,

P < 0.001) than those in the wild type (0.14).

We then released and quantified the wall-bound esters from

the pollen grains. Acetylesters were readily detected in the cell

walls of pollen grains of both the control and transgenic lines; we

measured a significant reduction (up to 35%) of acetylester

content in transgenic pollen grains compared with the control

pollen (Figure 8B). However, the level of methylesters of both the

control and transgenic pollen grains was below the detection

limit, indicating that the cell walls of themature pollen grainswere

less methylesterified than the leaves and floral tissues. In addi-

tion, we monitored the wall-bound ferulate ester in both groups

of pollen grains and found no quantitative difference in ferulate

ester contents between these groups. These data further confirm

that the reduction in ester signal observed in the FTIR micro-

spectroscopy analysis was mainly the result of an alteration in

acetylester content of the pollen cell wall.

Overexpression of Pt PAE Hinders Pollen Germination and

Pollen Tube Elongation

Subsequently, we examined the ability of pollen grains isolated

from PAE1-overexpressing lines to germinate. We collected

pollen from freshly dehisced flowers and incubated it for 4 h in

growth medium. No pollen tubes emerged from the pollen grains

of the transgenic lines (Figure 9A), but the majority of control

grains germinated, and their emerging pollen tubes elongated

(Figure 9B).

To verify these in vitro observations, we used the grains from

both the transgenic and the control lines to pollinate thewild-type

stigma. Twenty-four hours later, few pollen tubes had emerged

and grown from the transgenic pollen grains (Figure 9C, indi-

cated with the blue fluorescence resulting from aniline blue

staining) in contrast with the numerous tubes that emerged from

the control lines (Figure 9D).

To confirm the effects of PAE1-mediated deacetylation on the

germination and growth of pollen tubes, we incubated the pollen

grains from the wild-type tobacco plants with purified recombi-

nant PAE1 in growth medium. After 4 h, many pollen grains had

germinated, and their pollen tubes grew in the medium supplied

onlywith BSA as the control (Figures 9F and 9H). However, pollen

grains incubated in medium with exogenous PAE swelled and

most failed to produce tubes (Figures 9E and 9G). Moreover, a

few emerged tubes were shorter and bulkier than those of the

control (Figures 9G and 9H).

Overexpression of Pt PAE1 Depresses the Development of

the Capsule

The pollen tube delivers sperm cells to the female gametophyte

in the ovule. Failure of the pollen grain to germinate and of the

tube to elongate in plants overexpressing Pt PAE1 severely

inhibited fertilization and seed development.

Two weeks after anthesis, only a few seeds were set in the

developing seedpods of transgenic plants (Figure 10A). Eventually,

the transgenic lines yielded much smaller mature capsules (Figure

10C),with an averageweight of >70% less than that of thewild type

(Figure 10E), and the former contained few or no mature seeds.

The Overexpression of Pt PAE1 Lowers Pectin Digestibility

Reportedly, the acetylation of polysaccharides decreases their

digestibility (Schols and Voragen, 1994; Chen and Mart, 1996;

Figure 9. Pollen Grain Germination and Pollen Tube Elongation.

(A) and (B) Germination of mature pollen from Pt PAE1 overexpression (A) and control (B) plants in solid germination medium.

(C) and (D) Pollen germination and tube growth of Pt PAE1 overexpression (C) and the control (D) plants in the transmitting tract of the wild-type style.

(E) to (H) In vitro pollen germination and pollen tube growth when wild-type pollen was treated with exogenous Pt PAE1 recombinant protein ([E] and

[G]) or with BSA as the control ([F] and [H]).

Bars= 20 mm in (A) to (D) and 50 mm in (E) to (H).

58 The Plant Cell

Benen et al., 1999; Bonnin et al., 2003). To clarify the effect of

overexpression of PtPAE1 on pectin digestibility, we isolated the

acid-soluble pectins from the young leaves of the transgenic and

the control plants and treated equal amounts of these pectins

(based on quantification of uronic acids) with pectinase from

Aspergillus that randomly hydrolyzes a-(1-4)-D-galactosiduronic

linkages in pectin and other galacturonans (Pereira et al., 1992).

We labeled the digested products with 2-anthranilic acid, a

fluorophore that reacts specifically with the free reducing sugar

of glycan to form a stable Schiff’s base, therefore enabling us to

detect the labeled products via UV spectra (Sato et al., 1997,

2005). We then resolved the labeled hydrolysates by HPLC on a

silica-based size exclusion column and monitored them with a

UV diode array detector. Two predominant peaks representing

the mono- and oligo-sugars were detected in all of the treated

samples. In addition, a third peak characterized as the labeled

citric acids from the digestive buffer was also resolved (Figure

11A). Quantification of the resulting hydrolysates revealed that

the amount of released monomeric sugars in the pectin of the

transgenic plants was ;60% less than that from the control

plants (Figure 11B), indicating the lower digestibility of pectina-

ceous polysaccharides of the transgenic lines. To further clarify

whether the reduced digestibility was caused directly by the

deacetylation or was due to the indirect effect of the potential

alteration of pectin structure or composition, following the ex-

pression of Pt PAE1, we incubated sugar beet pectin with or

without recombinant Pt PAE1 and then treated the samples with

pectinase. After resolving the samples by HPLC, monomeric

sugars were readily observed in both samples (Figure 11C), but

the level of this digestive product in the sample pretreatedwith Pt

PAE1 was significantly lower than that of untreated sample

(Figures 11C and 11D). The acetic acid released from sugar beet

pectins by PAE1 was also labeled; its peak overlapped with that

of citric acid from the buffer (Figures 11C and 11D).

These data demonstrate that PAE1-mediated deacetylation

reduces the digestibility of pectin and hinders the release of small

molecule sugars, a finding that sharply contrasts with previous

studies on the acetylated hemicelluloses.

DISCUSSION

Poplar PAE1 Functions as a PAE

Continuous modification and remodeling of the composition and

structure of cell walls is a prominent feature of the growth and

development of plants (Ridley et al., 2001). Modulating the status

of acetylesters is a ubiquitous process in the cell wall polysac-

charides of higher plants, which necessitates the activity of

polysaccharide acetylesterases (Williamson, 1991; Bordenave

et al., 1995; Christensen et al., 1996). The distribution and

preferential expression of a set of putative Populus carbohydrate

acetylesterases in different tissues and at certain developmental

stages in poplar stems (see Supplemental Figure 1 online) high-

lights their potential diverse and distinct functions in remodeling

the complex configurations of the cell wall polysaccharides. Like

the previously isolated PAE gene from mung bean (Breton et al.,

1996), Pt PAE1 belongs to the Ser hydrolase superfamily,

possessing a conserved lipase GXSXG motif, a characteristic

of hydrolytic enzymes. Purified recombinant Pt PAE1 effectively

releases acetate from different natural and synthetically acety-

lated carbohydrates, among which the protein showed a cata-

lytic preference for acetylated pectin (Figure 2, Table 1).

However, the enzyme did not remove methyl- or feruloylesters

Figure 10. Development of the Seeds and Capsules of Pt PAE1 Transgenic Plants.

(A) to (D) Developing seeds in the capsules of the transgenic (A) and control (B) tobacco plants at 14 d postanthesis (the walls of capsules were peeled

off for clearance) and the mature capsules of the transgenic (C) and the control (D) plants. Bar = 0.5 cm.

(E) Average weight of capsules and produced seeds (per capsule) of transgenic (Pt PAE1_2, 4, 5, and 6) and four independent control ([A] to [D]) plants.

Data represent the mean of 10 capsules from each line. Error bar represents the SD. Asterisks indicate significant difference under pairwise comparison

to the control line (A); P < 0.01, Student’s t test.

[See online article for color version of this figure.]

Acetylesterase Modulates Plant Growth 59

frompolysaccharides that contain considerable amounts of such

esterifications, suggesting it is specific in modulating polysac-

charide acetylesters. Overexpressing this gene in tobacco sig-

nificantly reduced the content of acetylesters of pectin in young

leaves, the fast-growing floral tissues, and the pollen grains, in

which pectin is the predominant cell wall component. Although

the recombinant protein also showed low activity toward the

acetylated nascent xylan in vitro, constitutive expression of Pt

PAE1 in tobacco did not reduce the acetylation of xylan. In fact, in

some examined transgenic plants, the acetylester content of

xylan even increased slightly (Figure 4). This finding seems to

implicate a complex yet largely unknown spatial and metabolic

organization of polysaccharidemodification. It also highlights the

caution needed in interpreting the biological significance of in

vitro enzyme assays. Overall, our in vitro and in vivo data suggest

that Pt PAE1 primarily functions as a PAE.

PAEModulates Extensibility of Cell Walls in

Reproductive Tissues

Cell growth is intimately linkedwith the expansion of the cell walls

(Parre and Geitmann, 2005). Previous studies demonstrated the

dramatic consequences of the methylesterification status of

pectin on cell wall’s texture and mechanical properties and,

thereby, on cellular growth and cell shape (Bosch et al., 2005;

Bosch and Hepler, 2005; Wolf et al., 2009). In this study, we

found that Pt PAE1–mediated deacetylation greatly impeded

pollen grain germination and tube elongation (Figure 9). In the

pollen apertures and the apex of pollen tubes, the cell walls lack

the callosic inner lining, and pectin is their major component

(Geitmann and Steer, 2006). Although constitutive overexpres-

sion of PAE1 in planta caused complicated changes in cell wall

composition and structure, the impairments on the germination

of pollen grains and the extension of the pollen tube likely are

dominated by the reduction in acetylester in the cell wall pectin.

When exposing wild-type pollen grains to exogenous Pt PAE1

enzyme, the rehydrated pollen grains exhibited similar behavior

as those transgenic pollen grains expressing Pt PAE1 (Figure 9).

Although the inherent mechanisms underlying the inhibition of the

pollen tube’s germination andelongation in transgenic plantsmay

be complex, from a structural perspective, Pt PAE1–mediated

pectin deacetylation might change the physiochemical proper-

ties of the cell wall’s polysaccharide networks, thereby lowering

its extensibility. Kohn and Furda (1968) proposed that acetylation

of the hydroxyl at C-2 and/or C-3 of galacturonic acid residues

may impose steric hindrance, thus lowering the ability of pectic

polysaccharides to associate intermolecularly through calcium

ions. Conversely, deacetylation of pectin in transgenic plants

Figure 11. Effects of Pt PAE1 Overexpression on Pectin Digestibility.

(A) The digestion of leaf acid-soluble pectin from control (solid line) or transgenic (dotted lines) plants by pectinase.

(B) Quantification of monomeric and oligomeric sugars released in the reactions of (A).

(C) Pectinase digestion of sugar beet pectin with (dotted line) or without (solid line) recombinant PAE1 pretreatment.

(D) Quantification of the released monomeric and oligomeric sugars and acetic acid/citric acid in the reactions of (C). Data represent the mean

of triplicate samples. Error bar represents SD. A/C, acetic acid and citric acid overlapping peak; C, citric acid; M, galacturonic acid; O, oligomeric sugar.

*P < 0.05, Student’s t test; **P < 0.01, Student’s t test.

60 The Plant Cell

might promote such associations, thus altering the mechanical

properties and the configuration of the pectin network and con-

sequently disturbing the programmed remodeling and structure

of the primary cell walls, so affecting the cell walls’ asymmetric

extension/elongation.

Expressing Pt PAE1 also disturbs the elongation of the style

and filament tissues of flowers (Figure 5). The shorter epidermal

cells of the staminal filaments and of the basal style in transgenic

plants point to the limited expansion of their cell walls. The cells

of these organs usually contain high levels of pectinaceous

polysaccharides, although levels vary by species. By contrast,

the rigidwalls of the apical stylar cells have a lower pectin content

and more cellulose and hemicelluloses (Marga et al., 2003).

Expressing Pt PAE1 reduced the amount of acetylesters of the

stylar and filament pectins (Figure 6). Along with deacetylation of

the stylar and filament pectins, the extractable acidic polysac-

charideswere reduced;meanwhile, cellulosic and hemicellulosic

components were increased in the cell walls of those reproduc-

tive tissues of transgenic plants (Table 2). These findings explain

the retardation of cell elongation. Together, these results indicate

that expansion of the plant cell wall requires a precise acetylation

status of the polysaccharides; genetically perturbing such struc-

tural modification would affect the biogenesis, deposition, and

remodeling of the entire polysaccharides in the cell wall.

Deacetylation of Pectin Reduces Its Digestibility

The digestibility of pectin is another factor that likely influences

the cell wall’s dynamic remodeling and, thereby, cellular growth

and shape (Wolf et al., 2009). One of the proposed functions of

pectin acetylation is to provide a steric barrier that prevents the

pectinolytic enzymes from depolymerizing cell wall polysacchar-

ides (Solms and Deuel, 1951; Rexova-Benkova et al., 1997;

Andre-Leroux et al., 2009). Therefore, we anticipated that the

deacetylation of pectin by Pt PAE1 would enhance its degrada-

tion. Surprisingly, we found that deacetylation considerably

slowed the breakdown of pectin by microbial pectinase (Figure

11). The reduction in the digestibility of pectin by deacetylation

was circumstantially confirmed in vitro using sugar beet pectin

pretreated with recombinant Pt PAE1 (Figure 11). This experi-

ment excluded the potential effects of the other composition and

structure changes of pectin, caused by PAE1 overexpression in

transgenic plants, on its digestibility. This finding implies that

when hydrolyzing pectin polysaccharide, certain structural con-

figurations of the polymer, including the precise pattern of

acetylation, are a prerequisite for the action of pectolytic en-

zymes. The modulation of the digestibility of polysaccharides by

PAE1 might be an additional or alternative factor that regulates

cell wall extension in planta. Consistent with this assumption,

global transcriptomic analyses of pollen germination and tubular

growth in Arabidopsis revealed that the expression of putative

PAEs is reciprocally coordinated with that of the cell wall–

degrading enzymes (Wang et al., 2008; Qin et al., 2009). Com-

pared with their expression levels in pollen grains, six putative

PAE genes exhibited lower expression in the emerging tubes;

this downregulation was accompanied by an increase in the

gene expression of glycosyl hydrolase and pectin lyase (Wang

et al., 2008).

Together, these data suggest that, like pectin methylestera-

ses, PAE precisely regulates the acetylation status of polymers,

which modulates the biophysical and physiological properties of

the cell wall polysaccharides and thereby affects the plant’s

growth, development, and reproduction.

METHODS

Plant Materials and Growth Conditions

Poplar ecotype Populus trichocarpa and tobacco (Nicotiana tabacum cv

Xanthi) were used in this study. Poplar plants were grown in a greenhouse

under conditions described previously (Yu et al., 2009). Tobacco plants

were grown in a growth chamber at 228C with a 16/8-h light/dark regime.

All chemicals were from Sigma-Aldrich unless otherwise stated.

Phylogenetic and Sequence Analysis

Phylogenetic analyses of putative polysaccharide acetylesterases were

conducted on the full-length sequences of Populus and Arabidopsis

thaliana putative homologs, together with those of the functionally char-

acterized bacterial, fungal, and plant PAEs. Full-length amino acid se-

quences were first aligned by ClustalW version 1.83 (Thompson et al.,

1994) with default parameters (http://www.ebi.ac.uk/Tools/clustalw/) and

imported into the Molecular Evolutionary Genetics Analysis (MEGA)

package, version 3.0 (Kumar et al., 2004). Phylogenetic analyses were

conducted using the neighbor-joining method (Saitou and Nei, 1987)

implemented in MEGA, with the option for pairwise deletion for handling

alignment gaps and with the Poisson correction model for computing

distance. For statistic reliability, bootstrap tests were undertaken with

1000 replicates. The final tree graphics were generated via the TreeView

program (Page, 1996).

RNA Extraction and Quantitative RT-PCR

We collected tissues frompoplar leaves, stems, roots, and apical buds as

previously described (Yu et al., 2009). Total RNAwas extracted by the hot

borate method following Gou et al. (2008). Reverse transcription was

performed with the SuperScript III first-strand synthesis system (Invitro-

gen) according to the manufacturer’s user manual. We performed real-

time RT-PCR with iQ SYBR Green Supermix (Bio-Rad). The poplar

ATPase1 gene served as the calibration internal standard (Yu et al., 2009).

The normalized expression level in the leaf and first node was set at 100;

the expression values of individual genes in different tissues were

compared with those of the leaf and first node (set at 100) to give relative

expression levels. The data are the mean of triplicate samples.

Protein and Plant Expression Vector Constructs, Tobacco

Transformation, and Subcellular Localization Analysis

Full-length Pt CE13_5 cDNA was amplified from the stem’s reverse

transcription product and subcloned into a Gateway cloning vector

pDONR207 (Invitrogen). The gene was then transferred into the Gateway

destination vector pHis9, derived from pET-28a, for protein expression,

and both the Gateway binary vectors pMDC32 and pMDC43 (Curtis and

Grossniklaus, 2003) for expressing this gene alone and for creating a

CE13_5-GFP fusion in plant. The resulted pMDC32 construct constituted

an expression cassette of Pt CE13_5, driven by a double 35S promoter. It

was transferred first into the Agrobacterium tumefaciens GV3101strain

that was then used for tobacco transformation through leaf disc infection

(Horsch et al., 1985). Four independent transgenic lines, PtPAE1_2,

PtPAE1_4, PtPAE1_5, and PtPAE1_6, were selected by means of

hygromycin resistance (50 mg/mL) and planted in soil after rooting. The

Acetylesterase Modulates Plant Growth 61

plants were maintained in a growth chamber under the conditions de-

scribed. The resulted pMDC43 construct constituted a chimeric gene of Pt

CE13_5 linked to the 39 end of GFP under the 23 35S promoter. The

construct was transformed into tobacco via the same leaf disc infection

procedure (Horsch et al., 1985) for subcellular localization analysis. The

fluorescence signals of the expressed GFP-Pt CE13_5 fusion protein in

dark-grown tobaccoseedlingswere observedunder aZeiss LSM510META

NLO two-photon confocal laser scanning microscope (Carl Zeiss Micro-

Imaging). The same materials were then stained with 10 mg/mL propidium

iodide in water for 5 min and then observed by confocal microscopy.

pCAMBIA1302 that harbors the freeGFPgene, theoriginal construct used to

generate Gateway vector pMDC43 (Curtis and Grossniklaus, 2003), was

expressed in tobacco and used as the control.

Enzymatic Assays

Recombinant protein was produced in Escherichia coli strain Rosetta

(DE3) pLysS by inductionwith 0.5mM isopropyl b-D-1-thiogalactopyrano-

side, overnight, at 188C. The protein was purified with nickel-nitriloacetic

acid resin from crude cell lysate. Sugar beet (Beta vulgaris) pectin was

kindly provided by CP Kelco, birch wood (Betula spp) xylan was ordered

from Sigma-Aldrich, and potato (Solanum tuberosum) pectin was pre-

pared from the cell walls of potatoes as described (Melton and Smith,

2001b). The sugar beet pectin was further purified by precipitation with

70% ethanol. For substrate specificity assays, 2 mg recombinant proteins

was incubated with 10 mg/mL pectin or 50 mg/mL xylan, in 100 mM Tris-

HCl buffer, pH 7.0, at 358C for 30 min. Acetate released from the reaction

was immediately quantified using the Acetate Kinase Format Kit

(K-ACETAK) fromMegazyme (Megazyme International Ireland) according

to the user’s manual.

To determine pectin methylesterase activity, the assay was performed

using 1 mg/mL esterified pectin (>80% methyl esterification) from citrus

fruit as the substrate and incubating with 2mg recombinant protein, or the

commercial pectin methylesterase (Sigma-Aldrich) as the control, in 100

mM Tris-HCl buffer, pH 7.0, at 358C for 15 min. Methanol released was

subsequently converted by alcohol oxidase (Sigma-Aldrich) and formal-

dehyde dehydrogenase (Sigma-Aldrich) coupling with the conversion of

cofactor NAD to NADH. The produced NADH was measured at OD340

with methanol as standard (Grsic-Rausch and Rausch, 2004).

For assaying feruloylesterase activity, sugar beet pectin was incubated

with 2 mg recombinant protein in 50 mM Tris-HCl buffer, pH 7, at 358C, or

treated with 2 N NaOH (as the control) for 15 min; then, the reaction

mixtures were acidified to pH 4.0. The ferulic acid was extracted three

times with 0.5 mL water-saturated ethyl acetate. The extracts were

combined, dried under an N2 stream, dissolved in methanol, and quan-

tified by HPLC according to the description by Gou et al. (2008).

Polysaccharide Acetylester Content Analyses

To quantify the content of acetylester bound on polysaccharides, we

prepared cell wall materials from the young leaves, styles, filaments, and

stems of transgenic and the wild-type control plants on the day of

flowering. Water-soluble pectin was extracted with hot water (808C),

adjusted to pH4, and then pelleted overnight in 70%ethanol at2208C. To

prepare acid-soluble pectin, after hot water extraction, the residuals were

extracted with 0.04 M HCl (808C), and the acid extracts were pelleted in

70% ethanol as mentioned before.

Two methods were adopted to prepare xylan/hemicellulose. The leaf’s

cell wall was first treated with 1% ammonium oxalate at 378C overnight to

remove pectin. The residuals were then treated with 1% amylase over-

night. Hemicellulose was extracted with DMSO at 708 overnight, and

pelleted with a threefold volume of ethanol according to the description

provided by Lee et al. (2011). Alternatively, xylan was isolated from xylem

cell wall preparations by treating the samples with xylanase (Sigma-

Aldrich) according to the methods described (Gunl et al., 2010). The

acetates bound on different polymers were released by incubation with

2 N NaOH at 378C overnight. The hydrolysates were neutralized with HCl

and analyzedwith the Acetate Kinase Format Kit (Megazyme International

Ireland).

Cell Wall Sugar Composition Analysis

Cell walls from the styles and filaments of transgenic and control plants on

the day of flowering were prepared according to Gunl et al. (2010). Pectin

content was measured colorimetrically using galacturonic acid as the

standard (Melton and Smith, 2001a). Neutral sugars were released from

the prepared cell walls through sulfuric acid depolymerization and then

converted to their alditol acetates (Melton and Smith, 2001b). These

alditol acetates were separated on an Agilent gas chromatography–mass

spectrometer (Agilent Technologies) equipped with a 30 mm3 0.25-mm

(internal diameter) Agilent JandW HP-5MS capillary column; the initial

oven temperature was maintained at 388C for 30 s, increased to 1708C at

508C/min, and then increased to 2308C at 28C/min and held at 2308C for

5 min. The individual sugars were identified by comparison with authentic

standard compounds; their quantification was based on the standard

curves of each derivatized individual sugar made from the same gas

chromatography–mass spectrometry run.

FTIR Imaging Analysis on Floral Tissues and Pollen Grains

The styles and filaments of the transgenic and wild-type flowers were

collected on the day of flowering, frozen in liquid nitrogen, and kept at

2808C until use. The samples were embedded quickly in FSC22 Frozen

Section Compound (Surgipath Medical Industries) and sectioned at 10-mm

slices with a Leica CM1950 Cryo-microtome (Leica Microsystems). The

sections were mounted on a barium-fluoride slide, washed with 70%

ethanol thoroughly to remove the remaining embedding compound, and

left to dry. We imaged the sections in the transmission mode with a Perkin-

Elmer Spectrum Spotlight infrared microscope coupled to a Spectrum

One FTIR Spectrometer (Perkin-Elmer). The images were collected with a

6.25-mm pixel resolution, a spectral resolution of 8 cm21, and an inte-

gration time of 16 scans per pixel.

The wild-type and transgenic pollen grains, collected on the day of

flowering, were kept in ethanol until use. For FTIR microspectroscopy

analysis, we first stained the pollen grains with 1% ruthenium red for 5min

to monitor the methylester content and to facilitate their focus under the

FTIR microscope. The grains were then flattened with a diamond com-

pression cell (Spectra-Tech), washed with 70% ethanol, and scanned in

transmissionmodewith a 12-mmsquare aperture, a spectral resolution of

4 cm21, and integration time of 128 scans with a Perkin-Elmer Spectrum

Spotlight IRmicroscope in “point”mode. Ten pollen grainswere analyzed

for each condition. The data sets from each sample were averaged in

OMMIC software. To determine the relative acetylester levels, we deter-

mined the peak area of the carbonyl ester (1725 to 1775 cm21) and the

total polysaccharide (900 to 1180 cm21) and then calculated the ratio of

wall-bound esters to total polysaccharides.

The Pistil-to-Stamen Cell Length Ratio and Microscopy Analysis of

the Pollen Grain

Petals were removed from flowers at stage 12. The length of the style and

filament was measured from the receptacles of more than 20 flowers.

After peeling off the epidermis from the basal, middle, and apical parts of

the style and filament, we stained the material with 10 mg/mL propidium

iodide for 10min and observed it under an optical microscope (Leica). Cell

length was measured for an average of 40 cells with an E-Ruler (http://

www.mycnknow.com/). Dehiscent pollen grains at the same stage were

observed under an optical microscope. More grains at the same

62 The Plant Cell

developmental stage were fixed in 2.5% glutaraldehyde diluted with 0.1

M sodium phosphate buffer and then dehydrated in a graded ethanol

series and dried with a Polaron critical-point drying apparatus (Polaron

Instruments). Theyweremounted on aluminumstubs, sputter coatedwith

gold, and imaged with a Quanta 200 environmental scanning electron

microscope (FEI Company).

Pollen Tube Germination

Pollen grains collected on the day of flowering were germinated in growth

medium containing 2% agarose according to Palanivelu and Preuss

(2006). They were kept in a closed environment at room temperature for

4 h and then observed under a Leica DM 5500B microscope.

For in vivo germination, the anthers were removed at development

stage 10 from wild-type flowers. Stage 12 pollen grains from these

anthers and from overexpression plants were spread on wild-type stig-

mas on the day of flowering (i.e., stage 12). Twenty-four hours later, the

stylar tissues were fixed in acetic acid, softened in 2 N NaOH for 24 h,

stained with aniline blue for 24 h at 48C, and then washed three times with

water. The UV fluorescence of these tissues was then examined under a

Zeiss LSM 510 META NLO two-photon confocal laser scanning micro-

scope (Carl Zeiss MicroImaging).

Pectin Digestion and Anthranilamide Labeling

Eighty microliters of 0.5% (w/v) sugar beet pectin was incubated with 10

mg recombinant Pt PAE1 in 50 mM Tris-HCl buffer, pH 7.0, at 378C

overnight and then digestedwith 5mL (25 units) pectinase (Sigma-Aldrich)

in 200mMcitric acid buffer, pH 4.0, for 10min at 378C; alternatively, 50mL

of 0.5% acid-soluble pectin from the PAE1 transgenic and control plants

was directly treated with 5 mL pectinase (Sigma-Aldrich). The products

were pelleted by adding a sixfold volume of acetonitrile and sedimented

at 2208C for 1 h. The recovered pellets were dissolved in 0.2 M

2-anthranilic acid (Sigma-Aldrich) in 1 M sodium cyanoborohydride

(Sigma-Aldrich) to label the carbonyl carbon of the acyclic-reducing

sugars. The labeled glycan was held at 658C for 2 h. The products were

then cooled to room temperature and pelleted again with sixfold aceto-

nitrile, air-dried, and redissolved in 100 mL water. Twenty-microliter

samples were injected into a BioSep-SEC-2000S column (Phenomenex)

and separated in 0.2% acetic acid or formic acid buffer at a flow rate of

0.25 mL per minute with an Agilent 1100 HPLC system equipped with

a UV diode-array detector (Agilent). UV absorbance was monitored at

254 nm and quantified using a standard curve of 2-anthranilic acid.

Accession Number

Sequence data from this article can be found in the Arabidopsis Genome

Initiative or GenBank/EMBLdatabases under accession numberHQ223420

for the identified PAE1.

Supplemental Data

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

Supplemental Figure 1. Expression of Populus Putative Polysac-

charide Acetylesterase Genes.

Supplemental Figure 2. In Silico Expression of the Pt PAE1 Homo-

logs in Populus balsamifera.

Supplemental Figure 3. Acetylesterase Activity of Recombinant Pt

PAE1 with Different Chemically Acetylated Substrates.

Supplemental Figure 4. Acetylester Content of Natural Substrates

Released by 2 N NaOH for 15 min at 358C.

Supplemental Figure 5. Growth Phenotype of Independent Pt PAE1

Overexpression Lines.

Supplemental Figure 6. The Amount of Extracted Pectin from Leaf

Cell Walls of Transgenic and Control Plants.

Supplemental Figure 7. Content of Methylester of Water-Soluble

Pectin from Leaves of Transgenic and Control Plants.

Supplemental Figure 8. The Length of Style and Filaments.

Supplemental Figure 9. Content of Methylester in Water-Soluble

Pectins Extracted from Style and Filament of Transgenic and Control

Plants.

Supplemental Data Set 1. Multisequence Alignment of Polysacchar-

ide Acetyltransferases.

ACKNOWLEDGMENTS

We thank Simon Park, William Willis, and Randy Smith at the National

Synchrotron Light Source for their help with FTIR microspectroscopy.

Sugar beet pectin was kindly provided by CP Kelco U.S. This work was

supported by the Division of Chemical Sciences, Geosciences, and

Biosciences, Office of Basic Energy Sciences of the U.S. Department of

Energy through Grant DEAC0298CH10886 and by the National Science

Foundation through Grant MCB-1051675 (to C.-J.L.), the Chinese

Academy of Sciences/State Administration of Foreign Experts Affairs

International Partnership Program for Creative Research Teams in Plant

Metabolisms (to X.-Y.C), and the scholarship for distinguished overseas

researcher from the National Science Foundation of China (31028003; to

C.-J.L.). Use of the National Synchrotron light and confocal microscope

at the Center of Functional Nanomaterials was supported by the Office

of Basic Energy Sciences, U.S. Department of Energy, under Contract

DEAC02-98CH10886.

AUTHOR CONTRIBUTIONS

C.-J.L. and J.-Y.G. designed experiments. J.-Y.G, L.M.M., G.H., and

X.-H.Y. performed experiments. C.-J.L., J.-Y.G., and X.-Y.C. analyzed

data. C.-J.L. wrote the article.

Received October 7, 2011; revised December 2, 2011; accepted

December 22, 2011; published January 13, 2012.

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Acetylesterase Modulates Plant Growth 65

DOI 10.1105/tpc.111.092411; originally published online January 13, 2012; 2012;24;50-65Plant Cell

Jin-Ying Gou, Lisa M. Miller, Guichuan Hou, Xiao-Hong Yu, Xiao-Ya Chen and Chang-Jun LiuPlant Reproduction

Acetylesterase-Mediated Deacetylation of Pectin Impairs Cell Elongation, Pollen Germination, and

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