Structural basis for the interaction between pectin methylesterase and a specific inhibitor protein.
ABSTRACT Pectin, one of the main components of the plant cell wall, is secreted in a highly methyl-esterified form and subsequently deesterified in muro by pectin methylesterases (PMEs). In many developmental processes, PMEs are regulated by either differential expression or posttranslational control by protein inhibitors (PMEIs). PMEIs are typically active against plant PMEs and ineffective against microbial enzymes. Here, we describe the three-dimensional structure of the complex between the most abundant PME isoform from tomato fruit (Lycopersicon esculentum) and PMEI from kiwi (Actinidia deliciosa) at 1.9-A resolution. The enzyme folds into a right-handed parallel beta-helical structure typical of pectic enzymes. The inhibitor is almost all helical, with four long alpha-helices aligned in an antiparallel manner in a classical up-and-down four-helical bundle. The two proteins form a stoichiometric 1:1 complex in which the inhibitor covers the shallow cleft of the enzyme where the putative active site is located. The four-helix bundle of the inhibitor packs roughly perpendicular to the main axis of the parallel beta-helix of PME, and three helices of the bundle interact with the enzyme. The interaction interface displays a polar character, typical of nonobligate complexes formed by soluble proteins. The structure of the complex gives an insight into the specificity of the inhibitor toward plant PMEs and the mechanism of regulation of these enzymes.
- SourceAvailable from: Igor Vladimirovich Maksimov[Show abstract] [Hide abstract]
ABSTRACT: Functional traits, potentially associated with resistance to infection, were investigated in cultivars of bread wheat. Plant responses to infection by hemibiotrophic fungus Stagonospora (Septoria) nodorum were studied under laboratory conditions. Infection-induced up-regulation of genes coding for class III peroxidase, oxalate oxidase and protease inhibitor was greater in leaves with reduced disease symptoms (percentage of chlorotic or necrotic leaf area). Similar association was detected between activity of pectinase inhibitors and disease severity. Resistant cultivar also differed from the susceptible one by increased content of Н2О2 in infected tissues and more intensive deposition of lignin. We discuss possibility of using these functional traits in plant breeding for increased stress tolerance.Agricultural Sciences 07/2014; 5:722 - 729.
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ABSTRACT: BackgroundDUF642 proteins constitute a highly conserved family of proteins that are associated with the cell wall and are specific to spermatophytes. Transcriptome studies have suggested that members of this family are involved in seed development and germination processes. Previous in vitro studies have revealed that At4g32460- and At5g11420-encoded proteins interact with the catalytic domain of pectin methyl esterase 3 (AtPME3, which is encoded by At3g14310). PMEs play an important role in plant development, including seed germination. The aim of this study was to evaluate the function of the DUF642 gene At4g32460 during seed germination and plant development and to determine its relation to PME activity regulation.ResultsOur results indicated that the DUF642 proteins encoded by At4g32460 and At5g11420 could be positive regulators of PME activity during several developmental processes. Transgenic lines overexpressing these proteins showed increased PME activity during seed germination, and improved seed germination performance. In plants expressing At4g32460 antisense RNA, PME activity was decreased in the leaves, and the siliques were very short and contained no seeds. This phenotype was also present in the SALK_142260 and SALK_054867 lines for At4g32460.Conclusions Our results suggested that the DUF642 family contributes to the complexity of the methylesterification process by participating in the fine regulation of pectin status during plant development.BMC Plant Biology 12/2014; 14(1):338. · 3.94 Impact Factor
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ABSTRACT: Abstract After replication in the cytoplasm, viruses spread from the infected cell into the neighboring cells through plasmodesmata, membranous channels embedded by the cell wall. As obligate parasites, viruses have acquired the ability to utilize host factors that unwillingly cooperate for the viral infection process. For example, the viral movement proteins (MP) interacts with the host pectin methylesterase (PME) and both proteins cooperate to sustain the viral spread. However, how and where PMEs interact with MPs and how the PME/MP complexes favor the viral translocation is not well understood. Recently, we demonstrated that the overexpression of PME inhibitors (PMEIs) in tobacco and Arabidopsis plants limits the movement of Tobacco mosaic virus and Turnip vein clearing virus and reduces plant susceptibility to these viruses. Here we discuss how overexpression of PMEI may reduce tobamovirus spreading.Plant signaling & behavior 10/2014;
Structural Basis for the Interaction between Pectin
Methylesterase and a Specific Inhibitor Protein
Adele Di Matteo,a,bAlfonso Giovane,cAlessandro Raiola,b,1Laura Camardella,dDaniele Bonivento,a
Giulia De Lorenzo,bFelice Cervone,bDaniela Bellincampi,b,2and Demetrius Tsernogloua
aDepartment of Biochemical Sciences, University of Rome, 00185 Rome, Italy
bDepartment of Plant Biology, University of Rome, 00185 Rome, Italy
cDepartment of Biochemistry and Biophysics, Second University of Naples, I-80138, Naples, Italy
dInstitute of Protein Biochemistry, Consiglio Nazionale delle Ricerche, I-80125, Naples, Italy
Pectin, one of the main components of the plant cell wall, is secreted in a highly methyl-esterified form and subsequently
deesterified in muro by pectin methylesterases (PMEs). In many developmental processes, PMEs are regulated by either
differential expression or posttranslational control by protein inhibitors (PMEIs). PMEIs are typically active against plant
PMEs and ineffective against microbial enzymes. Here, we describe the three-dimensional structure of the complex
between the most abundant PME isoform from tomato fruit (Lycopersicon esculentum) and PMEI from kiwi (Actinidia
deliciosa) at 1.9-A˚resolution. The enzyme folds into a right-handed parallel b-helical structure typical of pectic enzymes.
The inhibitor is almost all helical, with four long a-helices aligned in an antiparallel manner in a classical up-and-down four-
helical bundle. The two proteins form a stoichiometric 1:1 complex in which the inhibitor covers the shallow cleft of the
enzyme where the putative active site is located. The four-helix bundle of the inhibitor packs roughly perpendicular to the
main axis of the parallel b-helix of PME, and three helices of the bundle interact with the enzyme. The interaction interface
displays a polar character, typical of nonobligate complexes formed by soluble proteins. The structure of the complex gives
an insight into the specificity of the inhibitor toward plant PMEs and the mechanism of regulation of these enzymes.
Pectin, one of the main components of the plant cell wall, is
continually modified and remodeled during plant growth and
development (Ridley et al., 2001). For example, the pattern of
pectin esterification changes during cell expansion, growth, and
fruit ripening as well as during infection by phytopathogenic
microorganisms (Steele et al., 1997; Willats et al., 2001). After
secretion into the wall as a highly methylesterified form, pectin is
deesterified in muro by pectin methylesterases (PMEs) (E.C.
184.108.40.206) in a spatially regulated manner during development
(Knox et al., 1990). Demethylation leads to the formation of
polyuronides aggregating into calcium-linked gels that are im-
portant in controlling the porosity and mechanical properties
of the wall (Willats et al., 2001). By generating free carboxylic
2001). PMEs produced by plants take part in important physio-
logical processes, such as microsporogenesis, pollen growth,
elongation, fruit ripening, and loss of tissue integrity (Tieman and
Handa, 1994; Wen et al., 1999; Micheli et al., 2000; Pilling et al.,
2000; Micheli, 2001; Pilling et al., 2004). They have also been
reported to play a role in response to fungal pathogens
(Wietholter et al., 2003) and are required for the systemic spread
Chen et al., 2000; Chen and Citovsky, 2003). PMEs are not only
produced byplantsbutalso bymicrobial pathogens(DeLorenzo
et al., 1997) and by symbiotic microorganisms during their
interactions with plants (Lievens et al., 2002).
Isoforms of PME differing by molecular weight, pI, and bio-
chemical activity are encoded by large families of genes, either
constitutively expressed (Giovane et al., 1994; Gaffe et al., 1997;
Micheli, 2001) or differentially regulated in specific tissues and
developmental stages (Micheli et al., 2000; Micheli, 2001). In ad-
dition to the transcriptional control, a mechanism of regulation of
were discovered in kiwi (Actinidia deliciosa) (Balestrieri et al.,
thaliana (Wolf et al., 2003; Raiola et al., 2004). These inhibitors,
named PMEIs, typically inhibit PMEs of plant origin and do not
affect the activity of microbial enzymes (Giovane et al., 2004).
Although a role of PMEIs in regulating the activity of endog-
enous PMEs is most likely, a physiological action of these
inhibitors toward enzymes derived from different species can-
not be excluded. It is known that PMEs and PMEIs are both
expressed in flower tissues and pollen grains (Wolf et al., 2003;
Markovic and Janecek, 2004; Raiola et al., 2004; L. Camardella,
A. Giovane, and D. Bellincampi, unpublished results) and that
wind and animal visitations continually bring pollen onto flowers
1Current address: Dipartimento del Territorio e Sistemi Agro-Forestali,
Universita ` di Padova, Italy.
2To whom correspondence should be addressed. E-mail daniela.
email@example.com; fax 39-06-49912446.
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy described
in the Instructions for Authors (www.plantcell.org) is: Daniela Bellin-
Article, publication date, and citation information can be found at
The Plant Cell, Vol. 17, 849–858, March 2005, www.plantcell.org ª 2005 American Society of Plant Biologists
of heterologous species. The kiwi inhibitor (AcPMEI, SwissProt
accession number P83326) is very efficient against PME of
tomato fruit (Lycopersicon esculentum) (PME-1, SwissProt ac-
cession number P14280) and forms a noncovalent 1:1 complex
(Mattei et al., 2002).
To date, the structures of only two PMEs, one from carrot
(Daucus carota) (PDB code 1GQ8) (Johansson et al., 2002) and
one from the bacterium Erwinia chrysanthemi (PDB code 1QJV)
(Jenkins et al., 2001), have been solved. Very recently, the
structure of the PMEI from Arabidopsis (At-PMEI1) has been
determined (Hothorn et al., 2004b), whereas structural informa-
crystal structure of the complex between a plant PME and its
specific inhibitor PMEI at 1.9-A˚resolution. This structure allows
a detailed analysis of the mode of interaction between the two
proteins in terms of specificity and sheds light into the regulation
of pectin deesterification in plants.
RESULTS AND DISCUSSION
easily separated by biochemical methods (Camardella et al.,
2000; Mattei et al., 2002). To obtain an amount of homogeneous
PMEI suitable for structural characterization, a synthetic gene
was generated on the basis of the amino acid sequence of the
prevalent PMEI isoform from kiwifruit (Camardella et al., 2000)
and expressed in Pichia pastoris. The protein, purified to homo-
geneity, displayed chemical, physical, and spectral properties
identical to those of the prevalent natural isoform from kiwi
(Scognamiglio et al., 2003). The enzyme was mixed with a molar
excess of inhibitor, and the resulting PME/PMEI complex was
purified by ion exchange chromatography.
The three-dimensional structure of the complex was deter-
mined at 1.9-A˚resolution using a combination of single iso-
morphous replacement and molecular replacement methods.
Details about data collection, phasing, and refinement statistics
are summarized in Table 1. The model, comprising 317 residues
for PME, 151 for PMEI, and 462 water molecules, has been
refined to an R factor of 20.0% and an Rfreeof 23.1% and has
the most favored or in the additional allowed regions of the
Ramachandran plot (Table 1).
The Structure of Tomato PME Exhibits the Typical Fold
of Pectic Enzymes
PME-1from tomato belongsto family CE8 of the sequence-based
classification of carbohydrate esterases (http://afmb.cnrs-mrs.fr/
Table 1. Data Collection, Phasing, and Refinement Statistics
Data CollectionForm A Form BK2OsO4
Unit cell dimensions
a ¼ b ¼ 120.26 A˚;
c ¼ 97.29 A˚; a ¼ b ¼ g ¼ 908
a ¼ b ¼ 90.38 A˚;
c ¼ 149.1 A˚; a ¼ b ¼ 90.08;
g ¼ 120.08
50.0 to 1.9
a ¼ b ¼ 120.53 A˚;
c ¼ 97.40 A˚;
a ¼ b ¼ g ¼ 908
25.0 to 3.2
Resolution limits (A˚)
30.0 to 2.8
Completeness (last shell)
Average I/s (last shell)
Soaking time (h)
FOM (before DM)a
FOM (after DM)
Percentage of residues in allowed
Percentage of residues in additional
Percentage of residues in generously
Percentage of residues in not allowed
25.0 to 1.991.9
Amino acid residues
Amino acid residues
aDM, density modification; FOM, figure of merit.
850The Plant Cell
observed in pectate lyase C (Yoder et al., 1993) and typical of
pectic enzymes (Jenkins and Pickersgill, 2001) (Figure 1). The
b-helix consists of seven complete coils, which have different
lengths because the number of amino acids located in the loops
connecting the b-strands is variable. Each coil consists of three
b-strands that line up to form three extended parallel b-sheets
called PB1, PB2, and PB3. T1 identifies the stack of turns
between PB1 and PB2, T2 the stack between PB2 and PB3, and
T3 those between PB3 and PB1. Letters following T identify the
position of each turn with respect to the coil of the b-helix,
whereas A corresponds to the first turn in the N-terminal region.
Turns T1 (except for TB1) are short and mainly composed of
residues in aL-conformation and are responsible for the sharp
bend between the sheets as observed in other parallel b-helix
structures (Federici et al., 2001; Jenkins and Pickersgill, 2001).
Turns T2 and T3 are generally longer and more variable; in
particular, TF3 and most of T2 turns protrude from the central
parallel b-helix to form the shallow cleft where the putative ac-
tive site is located. In contrast with what was reported (Markovic
and Jornvall, 1992), no electron density corresponding to the
disulphide bridges Cys98-Cys125 and Cys166-Cys200 was
observed. The absence of these bridges was confirmed by
upon titration with the Ellman’s reagent in denaturing conditions
(data not shown). The N-terminal region of PME is composed by
a short a-helix followed by a b-strand that lines up with PB1. The
C-terminal region has an extended conformation in which a long
tail and four short and distorted a-helices protrude out of the
parallel b-helix flanking PB1.
The putative active site of PME is located on the PB3 sheet in
a cleft shaped by TB2, TC2, TF2, and TF3. Many aromatic
residues (Phe80, Tyr135, Phe156, Tyr218, Trp223, and Trp248)
putatively involved in substrate binding arelocated in thispocket
(Johansson et al., 2002). These residues are well conserved in
plant PMEs (Markovic and Janecek, 2004). Tyr135, Phe156, and
Trp223 are also conserved in PME of E. chrysanthemi (Jenkins
et al., 2001). Asp132, Asp153, and Arg 221, located inside the
crevice, have been hypothesized to be the catalytic residues
is located 2.82 A˚from and interacts with the NE of Arg221,
NH2 of Arg221. Moreover, OD2 of Asp153 is at H-bonding
distance (2.63 A˚) from a water residue (W227) that also forms an
proposed mechanism of action of PME from carrot (Johansson
et al., 2002), we can infer a mechanism of catalysis in which
Asp153, polarized by the proximity with Arg221, performs
a nucleophilic attack on the carboxymethyl group of the sub-
strate. The tetrahedral anionic intermediate formed is stabilized
by the interaction with two conserved Gln residues (Gln109 and
Gln131). Afterwards, Asp132 likely acts as a proton donor in the
cleavage step where methanol is released. The resulting car-
boxylate group of Asp132 then behaves as a base and receives
by Johansson (Johansson et al., 2002) foresees a primary
nucleophilic attack performed by the water molecule deproto-
nated both by Asp132 and Asp153.
Superimposition of the known PME structures of carrot, E.
chrysanthemi, and tomato reveals the similarity of the overall
folding topologies. The similarity of tomato and carrot PMEs is
more extensive with a root mean square deviation (RMSD) value
of 0.7 A˚calculated for all Ca atoms (Figure 3A), whereas the
bacterial enzyme can be well superimposed to tomato PME only
for 284 Ca atoms out of 317 and with a higher RMSD value of
1.8 A˚(Figure 3B). The main differences between the plant and
the bacterial enzymes are located on TB2, TC2, TF2, TG3, and
TH3; these turns protrude out of the b-helix and are much longer
in the bacterial enzyme, making its putative active site cleft
deeper and narrower than that of plant PMEs.
The Inhibitor Folds in an Up-and-Down
PMEI is almost all helical, with four long helices (a1, a2, a3, and
a4) aligned in an antiparallel manner in a classical up-and-down
four-helical bundle (Figure 1). The interior of the bundle is
stabilized by hydrophobic interactions and by a disulphide
bridge between Cys74 and Cys114, which connects helices a2
and a3. The N-terminal region, composed of three short and
distorted helices (aa, ab, and ac), extends outside the central
domain and lines roughly parallel to the plane defined by the
helices a1 and a4. A disulphide bridge between Cys9 and Cys18
connects aa and ab.
Figure 1. Structure of the PME-PMEI complex.
Ribbon representation illustrating the relative positions of PMEI and PME
in the complex. The enzyme is shown in green–blue on the left side. The
inhibitor is represented in yellow–red on the right side; the a-helices of
the four-helix bundle are indicated as a1 to a4, whereas helices of the
N-terminal region are named aa, ab, and ac. The inhibitor binds the
active site region of the enzyme, hampering its access to the substrate.
3D Structure of the PME-PMEI Complex 851
According to sequence-based classification, PMEIs belong
to the family PF04043 (Pfam database, http://pfam.wustl.edu/)
of invertase inhibitor (INH)/PMEIs and share with INH several
structural properties (Scognamiglio et al., 2003). Recently, the
structure of an invertase inhibitor from tobacco (Nicotiana
tabacum) (Nt-CIF) has been elucidated (PDB code 1RJ1)
(Hothorn et al., 2004a) as well as the structure of a PMEI
from Arabidopsis (At-PMEI1) (Hothorn et al., 2004b). Structural
superimposition of PMEI from kiwi and Nt-CIF reveals a striking
similarity between the two proteins, although their sequence
identity is rather low (29.2%) (Figure 4). An RMSD value of 1.7 A˚
calculated on 144 Ca out of 151 confirms that the overall fold is
very similar in both inhibitors. Main differences are located in
the N-terminal region and in the loops connecting the helices of
the bundle; notably an amino acid insertion, located in helix a2
of Nt-CIF partially distorts the helix (Figure 4). Structural
superimposition between At-PMEI1 and AcPMEI is not possible
because coordinates of At-PMEI1 are not yet available. How-
ever, considering the superimposition of At-PMEI1 and Nt-CIF
(Hothorn et al., 2004b), the two inhibitors are quite similar in
the bundle, whereas significant differences are located in the
N-terminal extension. It is puzzling that the N-terminal region of
AcPMEI folds back and packs with the bundle through hydro-
phobic interaction (Figure 1), whereas the N-terminal extension
of At-PMEI1, which crystallizes in a dimeric form, packs against
the bundle of another molecule (Hothorn et al., 2004b). In-
terestingly Pro-28 of At-PMEI1, which is located in the linker
between the N-terminal region and the four-helix bundle and
is responsible for the orientation of the N-terminal region, is
replaced by a Lys in AcPMEI, suggesting that the different
topology of the two inhibitors is due to the presence of different
residues at the same position.
The PME-PMEI Complex
PME and PMEI form a stoichiometric 1:1 complex in which the
inhibitor covers the shallow cleft of the enzyme where the
putative active site is located. The four-helix bundle of PMEI
packs roughly perpendicular to the parallel b-helix of PME, and
the three helices a2, a3, and a4, but not a1, interact with the
enzyme in proximity of the putative active site (Figures 1 and 5)
Figure 2. Close-Up View of the Tomato PME Active Site.
(A) Structure of tomato PME in which residues involved in catalysis (violet), in stabilization of the catalytic intermediate (orange), and in substrate binding
(blue) are shown in ball and stick representation.
(B) Further close-up view representation of amino acid residues and a water molecule (blue ball) putatively involved in catalysis; H-bond pattern is
Figure 3. Comparison of the Known Structures of PMEs.
(A) Overlay of the Ca trace of PME from tomato (green) and PME from
carrot (orange). Structures are almost completely superimposable, with
a RMSD value of 0.7 A˚, calculated on all Ca.
(B) Superimposition of PME from tomato (green) and PME from E.
chrysanthemi (violet). The RMSD value, calculated on 284 out of 317 Ca,
is 1.8 A˚. Although the b-helices are completely superimposable, main
differences are located in the length of the turns protruding out from the
b-helix in proximity of the putative active site cleft.
852The Plant Cell
The relative position of a2 and a3 helices is maintained by
a disulphide bridge between Cys74 and Cys114. The N-terminal
region of PMEI is poorly involved in the formation of the complex
and may play a role in the structural stability of the inhibitor, as
proposed for Nt-CIF (Hothorn et al., 2004a). Superimposition of
the free Nt-CIF with the complexed PMEI shows a similar fold
and orientation of helices in the bundle (Figure 4). Similarly, the
structure of the free carrot PME is almost completely superim-
posable to the structure of tomato PME engaged in the complex
with PMEI (Figure 3A). These features suggest that PME and
PMEI do not undergo dramatic conformational changes upon
formation of the complex.
Upon interaction, PME and PMEI bury 1148 A˚ 2and 1060 A˚ 2,
respectively, of their accessible surface area (ASA). The total of
2208 A˚2buried surface is somewhat larger than the average
interface area reported for noncovalent protein complexes
(DASA 1600 6 400 A˚2) (LoConte et al., 1999). The interaction
interface displays a surprisingly high polar character. No ex-
tended zones of hydrophobic interactions are present, whereas
more than half (55%) of buried surface arises from polar and
charged atoms (Figure 6). A large number of water molecules is
present at the interface and 17 of them mediate intermolecular
H-bonds (Figure 6). Such a polar character of the interface is
typical of nonobligate complexes formed by soluble proteins,
form (Jones and Thornton, 1996). Fifty residues (23 on PME and
27 on PMEI) establish contacts (Table 2). Twenty-two of these
residues are engaged in H-bonds, and four form salt bridges
(Table 3). Residues of PME forming contacts are mostly located
in the proximity of the putative pectin binding site, particularly on
PB3 and on T3 (Figure 2A). A large cluster of interacting residues
is present on turns TD3 and TF3 and on the fourth strand of PB3.
whereas only one contact is present in the C-terminal tail of the
enzyme. The contact residues of PMEI are mainly located on
helices a2 and a3, with a continuous surface that extends all
along. Ten residues are located on a2, 11 on a3, and four on a4;
two residues reside on the N-terminal region of the inhibitor.
In the article by Hothorn et al. (2004b), the N-terminal region of
At-PMEI1 has been proposed to be crucial for the interaction
with PMEs. This model does not fit with our crystallographic
structure of the PME-PMEI complex, and we cannot exclude
that the mode of interaction of At-PMEI1 to PMEs is some-
what peculiar. Experimental data using chimeras between the
N-terminal region of At-PMEI1 and the four-helix bundle of
Nt-CIF indicate that ;100 times more quantity of the chimera
is needed to obtain the same inhibition played by the natural
At-PMEI1. This suggests that the four-helix bundle is also
important for the interaction of At-PMEI1 and PME.
Figure 4. Structural Superimposition between PMEI and Nt-CIF.
PMEI (red) and Nt-CIF (blue) are superimposable, with a RMSD of 1.7 A˚,
calculated on 144 out of 151 Ca. Main differences are located in the
loops connecting the helices of the bundle and in the N-terminal region;
a distortion in the a2 helix of Nt-CIF is indicated by the arrow. Conserved
disulphide bridges are represented in yellow.
Figure 5. Molecular Surface of the PME-PMEI Complex.
(A) Representation of the molecular surface of the enzyme (violet) and the
inhibitor (yellow) in the complex.
(B) Same view of the complex as in (A), showing the molecular surface of
PME and a ribbon diagram of PMEI. The a-helices a2, a3, and a4 of the
inhibitor fit into the substrate binding cleft of the enzyme.
Figure 6. Representation of the Interacting Surface of PME and PMEI.
To open up the complex, PMEI has been shifted along its major axis and
rotated by 1808 around the vertical axis indicated. The molecular
surfaces contributed by carbon atoms are in green, and those contrib-
uted by oxygen and nitrogen are in red and blue, respectively. Water
molecules involved in water-mediated hydrogen bonds are represented
as violet spheres.
3D Structure of the PME-PMEI Complex853
Whereas the electrostatic potential surface of PMEI shows an
acidicpatchformed byGlu76,Asp80,andAsp83on a2helix and
Asp 96, Asp109, and Asp116 on a3 helix, a positive counterpart
is not found on the potential surface of PME. However, Asp116
(OD1 and OD2) and Glu76 (OE1) of PMEI are involved in salt
bridges with Lys224 (NZ) and Arg81 (NE) of PME, respectively.
PMEI complex at physiological pH (Kd¼ 5 nM, pH 5.5) (Mattei
et al., 2002).
The stability of the complex is pH dependent, being higher in
acidic conditions, typical of the apoplastic environment, and
decreasing drastically by raising the pH from 6.5 to 7.5; no
formation of the complex occurs at pH 8.5 (D’Avino et al., 2003).
The contact between the NE2 of His137 in PME and OG1 of
Thr113 in the inhibitor (2.92 A˚) may be crucial for determining the
strength of the interaction in this pH range. The contribution of
a high number of ionizable groups, the pK value of which is
affected by their chemical environment, could also be important.
Interestingly, at pH 6.0, the condition at which structure of the
complex has been solved, OD1 of Asp140 in PMEI is located 2.4
and 3.21 A˚from OD2 and OD1, respectively, of Asp188 in PME
them is protonated and acts as a hydrogen bond donor. Asp188
is located on the TF3 loop, where a wide patch of interacting
residues that forms a network of intermolecular H-bonds
(Asp188, Asn190, Gln191, and Ala192) is located (Table 2). We
hypothesize that at higher pH values the deprotonation of an
aspartic residue generates an electrostatic repulsion that loos-
ens the H-bond network and destabilizes the complex. Interest-
ingly, both PMEIs from Arabidopsis exhibit a Gly residue instead
of Asp140 and form a complex with a lower pH dependence with
tomato PME (Raiola et al., 2004).
Detailed analysis of residues involved in forming the complex
reveals thattheputative catalyticresidues(Asp132,Asp153, and
Arg221) do not establish contacts with the inhibitor, neither do
Gln109 and Gln131, which are thought to stabilize the anionic
intermediate formed after the first nucleophilic attack. Instead,
three aromatic residues (Phe80, Tyr135, and Trp223), likely
responsible for substrate binding, interact with the inhibitor.
Remarkably, Phe80 is one of the residues mostly involved in the
interaction, burying an area of 81 A˚2upon formation of the
complex. This residue establishes 17 contacts with four different
residues of the inhibitor (Thr73, Glu76, Asn77, and Thr113) and
a water-mediated hydrogen bond. Trp223 of PME forms three
contacts with its interacting counterpart, whereas Tyr135 forms
only one contact; moreover, each of them forms a water-
mediated hydrogen bond. Upon formation of the complex with
the inhibitor, Trp223 buries almost half of its solvent-exposed
in PME upon addition of the inhibitor (D’Avino et al., 2003). We
can infer the mode of action of the inhibitor: on one hand, the
inhibitor covers the active site cleft preventing the access of
the substrate, and on the other hand, it prevents the interactions
of Phe80, Tyr135, and Trp223 with the substrate. These obser-
vations are consistent with the observed competitive mode of
inhibition. Notably, a similar mode of inhibition has been pro-
posed for PGIP, a protein inhibitor of fungal polygalacturonases
(De Lorenzo et al., 2001; Di Matteo et al., 2003), suggesting that
Table 2. PME-PMEI Intermolecular Contacts
Enzyme Residues Inhibitor ResiduesNumber of Contacts
Thr73 Arg70 Gly69
Thr73 Glu76 Asn77 Thr113
Gly69 Glu72 Thr73 Glu76
Ser84 Ile102 Tyr103 Ala106
Ser99 Ile102 Tyr103
Phe108 Asp140 Leu143
Lys11 Asn101 Asp140
Leu143 Val144 Asn147
Asn101 Ser105 Phe108
Asn101 Ile102 Ser105
The number of atom–atom contacts at a distance of below 4.1 A˚have
been identified using the program Contact (CCP4).
Table 3. Hydrogen Bonding and Salt Link Interaction between PME
and PMEI in the Complex
Ala 192 N
(*), Salt link interaction.
854 The Plant Cell
this may represent a general strategy evolved by plants for
controlling pectic enzymes. Because mutations in residues in-
volved in substrate binding affect enzyme activity, they are likely
to be counter-selected during evolution. The involvement of
these residues in the interaction with PMEI therefore minimizes
the possibility for plant PMEs to escape recognition.
The structure of the PME-PMEI complex provides a possible
explanation for the lack of inhibition of PMEIs on PMEs from the
bacterium E. crysanthemi and the fungus Aspergillus aculeatus
(Giovane et al., 2004; Raiola et al., 2004). In E. crysanthemi, the
putative binding site cleft is much deeper than in plant-derived
PMEs (Figure 3B). It is likely that the external loops of the
bacterial enzyme create a steric hindrance that prevents the
interaction with the inhibitor. On the other hand, the amino acid
sequence alignment among PME from tomato, carrot, and A.
residues important for the interaction of tomato PME with the
inhibitor are conserved in plant PMEs but not in the fungal
enzyme, thus providing a reason for the observed lack of
interaction (Figure 7).
PMEI and Nt-CIF exhibit an almost identical fold (Figure 4) but
recognize different target enzymes. The structural view of the
PME-PMEI complex also provides a possible explanation for the
absence of interaction of Nt-CIF with PME. Sequence compar-
ison between the PMEIs characterized so far and Nt-CIF shows
that the subset of residues of the kiwi inhibitor, Asn101, Asp109,
Thr113 (located on a3), and Asn147 (located on a4), which form
intermolecular H-bonds with the enzyme, are conserved only in
PMEIs. In addition, an amino acid insertion that produces
a distortion of the a2 helix of Nt-CIF is close to residues cor-
in H-bonds with PME (Figures 4 and 8). We speculate that the
lack of residues important for the formation of the complex as
well as the distortion of a2 helix of Nt-CIF compared with that of
PMEI are responsible for the lack of interaction between Nt-CIF
Figure 7. Sequence Comparison of PMEs from Tomato (PME_LYCES), Carrot (PME_DAUCA), and A. aculeatus (PME_ASPAC).
Residues involved in H-bonds (violet), Van der Waals contacts (green), and water-mediated H-bonds (yellow) with the inhibitor are shown. Conserved
residues are in blue.
Figure 8. Sequence Comparison of PMEIs form Kiwi (AcPMEI), PMEIs from Arabidopsis (AthPMEI-1, Accession Number NP_175236; AthPMEI-2,
Accession Number NP_188348), and Invertase Inhibitor from Tobacco (NtCIF).
Residues of the kiwi inhibitor involved in H-bonds (violet), Van der Waals contacts (green), and water-mediated H-bonds (yellow) with tomato PME are
shown. Conserved residues are in blue.
3D Structure of the PME-PMEI Complex855
Expression and Purification of PMEI and PME
A synthetic AcPMEI gene was designed on the basis of the amino acid
sequence of the mature PMEI from kiwifruit (Actinidia deliciosa) (AcPMEI
accession number P83326 NCBI database) and expressed in Pichia
pastoris. The synthesis of three AcPMEI DNA gene fragments was
performed by PCR using PWO DNA Polymerase (Roche, Penzberg,
a pUC19 plasmid vector using two internal restriction sites designed to
facilitate cloning. AcPMEI was amplified from pUC19 and cloned into the
pPICZaA expression vector, used to transform P. pastoris strain X-33
(Invitrogen, Carlsbad, CA). The transformed cells were grown 3 d after
induction with 0.5% (v/v) methanol; the supernatant of the culture was
collected and total proteins precipitated with 80% (w/v) ammonium
sulfate. The precipitated fraction was dissolved in 10 mM Tris-HCl, pH
column (Amersham Pharmacia Biosciences). AcPMEI was eluted by
applying a linear gradient of 0 to 0.5 M NaCl in 10 mM Tris-HCl, pH 7.5.
The eluted inhibitor was concentrated and loaded onto a HiLoad 16/60
Superdex 75 column (Amersham Pharmacia Biosciences) equilibrated in
10 mM Tris-HCl, pH 7.5, and 0.25 M NaCl. The purified inhibitor,
from kiwi, exhibited a single band by SDS-PAGE and showed a single
peak upon reverse-phase HPLC on a Vydac C4 column. The N-terminal
sequence of the protein confirmed its identity and indicated the presence
of four additional amino acid residues, a remnant of the yeast signal
sequence. These additional amino acids did not impair the inhibitory
nM) (data not shown). The isoform PME-1 of tomato (Lycopersicon
esculentum) (SwissProt accession number P14280) was purified as
reported (Giovane et al., 1994). The inhibitory activity of AcPMEI against
PME-1 was determined by automatic titration as previously described
(Giovane et al., 1995). The inhibitory constant was calculated by Dixon
plot using 0.3% and 0.05% citrus pectin (63 to 66% methylation degree
from Sigma-Aldrich, St. Louis, MO).
Purification of the Complex
The PME-1/AcPMEI complex was obtained upon mixing PME-1 with
was purified by fast protein liquid chromatography on a Mono S HR 5/5
column (Amersham Pharmacia Biosciences) by applying a linear gradient
of 0 to 0.5 M NaCl and concentrated through Centriplus YM-3 filters
(Millipore, Bedford, MA).
Crystallization, X-Ray Data Collection, Structure Determination,
The PME-PMEI complex crystallizes in two different crystal forms (A and
B), both grown using the vapor-diffusion technique at 218C. Form A was
with a reservoir solution containing 2.5 M NaCl, 0.1 M sodium acetate,
pH 4.5, and 0.2 M Li2SO4. Crystals appeared after 3 d and reached
final dimensions of ;0.4 3 0.5 3 0.5 mm after 1 week. Crystals were
transferred to a cryoprotectant solution consisting of 2.7 M NaCl, 0.1 M
sodium acetate, pH 4.5, 0.2 M Li2SO4, and 22% polyethylene glycol 200
for 1 min, then looped from the drop and flash-frozen in a nitrogen stream
at 100 K (Oxford Cryosystems, Oxford, UK). Diffraction data were
collected to 2.8-A˚resolution at the XRD Beamline of the ELETTRA
Synchrotron (Trieste, Italy). Crystals of form A belong to the space group
P41212, with unit cell dimensions a ¼ b ¼ 120.26 A˚, c ¼ 97.29 A˚, and a ¼
b ¼ g ¼ 90.08. Single crystals of form B were obtained in drops made up
by 1 mL of 11.0 mg/mL of purified complex and 1 mL of well solution con-
sisting of 1.6 M MgSO4, and 0.1 M Mes, pH 6.0. After 1 week, crystals
grew to ;0.5 3 0.6 3 0.3 mm. These crystals were flash-frozen in the
presence of 25% (v/v) glycerol in mother liquor, and diffraction data were
collected to 1.9-A˚resolution at the XRD Beamline of the ELETTRA Syn-
chrotron. Crystals of form B belong to the space group P3221, with unit
cell dimension of a ¼ b ¼ 90.38 A˚, c ¼ 149.1 A˚, a ¼ b ¼ 908, and g ¼ 1208.
anomalous scattering methods. The crystal of form A was soaked in
a solution containing 2.5 M NaCl, 0.1 M sodium acetate, pH 4.5, 0.2 M
flash-frozenin liquid nitrogen.Data were collected at the BW7A Beamline
of the Deutsches Elektronen Synchrotron (Hamburg, Germany). Oscilla-
tion images were integrated, scaled, and merged using DENZO and
SCALEPACK (Otwinowski and Minor, 1996). The program SOLVE
(Terwilliger, 1999) was used to perform heavy metal Patterson search
with the derivatized and native crystal of form A, leading to an overall
figure of merit of 0.45 at 30- to 3.5-A˚resolution. Solvent flattening as
implemented in the program RESOLVE (Terwilliger, 2000) was performed
resolution. The resulting electron density map was sufficiently connected
to build a partial model of the main chain of PMEI using QUANTA
(Molecular Structure, Woodlands, TX). The structure of PME from carrot
(Daucus carota) (PDB code 1GQ8) was used as a guide to locate the
b-helix of the tomato PME into the density. No interpretable density for
the N terminus and C terminus of PMEI as well as for all side chains of the
inhibitor was available at this stage. The partially built model was then
used as a template to perform a molecular replacement with native data
of form B with the program AMORE (Navaza, 1994), obtaining a clear and
interpretable electron density map at 1.9-A˚resolution that was used to
complete the tracing. The model was iteratively refined using REFMAC5
(Murshudov et al., 1997), subjected to a simulated annealing procedure
as implemented in CNS (Adams et al., 1997), visually inspected, and
manually rebuilt. Several water residues were added into the Fo-Fc
density map, countered at 4s, with the X-SOLVATE tool of QUANTA.
Only solvent residues with a Bfactor# 45 A˚ 2and hydrogen bonded to the
protein molecule were kept in the structure refinement. Further refine-
ment and rebuilding led to a crystallographic R factor of 20% and an Rfree
150 of chain B (PMEI), and 462 water residues. A total of 91.9% of
residues are in the most favored region of the Ramachandran space, with
7.9% in additional allowed regions, 0.2% in generously allowed regions,
and none in disallowed regions. Secondary structure assignment and the
check of the geometrical quality were performed using PROCHECK
(Laskowski et al., 1993). The atomic coordinates have been deposited in
the Protein Data Bank (www.pdb.org; PDB ID code 1XG2).
We are grateful to Maurizio Brunori for is encouragement and valuable
Deutsches Elektronen Synchrotron (Hamburg, Germany), and the Euro-
pean Synchrotron Radiation Facility (Grenoble, France) for beam time
allocation and technical support. This research was supported by the
European Union Gemini project (contract QLK1-2000-00911), the In-
stitute Pasteur-Fondazione Cenci Bolognetti, the Giovanni Armenise-
Harvard Foundation, and the Ministero dell’Universita ` e della Ricerca
Scientifica (PRIN 2002).
Received October 27, 2004; accepted December 28, 2004.
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