Ester Cross-Link Profiling of the Cutin Polymer of
Wild-Type and Cutin Synthase Tomato Mutants
Highlights Different Mechanisms of Polymerization1
Glenn Philippe, Cédric Gaillard, Johann Petit, Nathalie Geneix, Michèle Dalgalarrondo, Cécile Bres,
Jean-Philippe Mauxion, Rochus Franke, Christophe Rothan, Lukas Schreiber, Didier Marion, and
Bénédicte Bakan*
Institut National de la Recherche Agronomique, Research Unit Biopolymers Interactions Assemblies, BP71627
44316, Nantes cedex 3, France (G.P., C.G., N.G., M.D., D.M., B.B.); Institut National de la Recherche
Agronomique, University of Bordeaux, Unité Mixte de Recherche 1332 Fruit Biology and Pathology, 33140
Villenave d’Ornon, France (J.P., C.B., J.-P.M., C.R.); and Institute of Cellular and Molecular Botany, University
of Bonn, D–53115 Bonn, Germany (R.F., L.S.)
ORCID ID: 0000-0001-7003-9929 (L.S.).
Cuticle function is closely related to the structure of the cutin polymer. However, the structure and formation of this hydrophobic
polyester of glycerol and hydroxy/epoxy fatty acids has not been fully resolved. An apoplastic GDSL-lipase known as CUTIN
SYNTHASE1 (CUS1) is required for cutin deposition in tomato (Solanum lycopersicum) fruit exocarp. In vitro, CUS1 catalyzes the
self-transesterification of 2-monoacylglycerol of 9(10),16-dihydroxyhexadecanoic acid, the major tomato cutin monomer. This
reaction releases glycerol and leads to the formation of oligomers with the secondary hydroxyl group remaining nonesterified.
To check this mechanism in planta, a benzyl etherification of nonesterified hydroxyl groups of glycerol and hydroxy fatty acids was
performed within cutin. Remarkably, in addition to a significant decrease in cutin deposition, mid-chain hydroxyl esterification of
the dihydroxyhexadecanoic acid was affected in tomato RNA interference and ethyl methanesulfonate-cus1 mutants. Furthermore,
in these mutants, the esterification of both sn-1,3 and sn-2 positions of glycerol was impacted, and their cutin contained a higher
molar glycerol-to-dihydroxyhexadecanoic acid ratio. Therefore, in planta, CUS1 can catalyze the esterification of both primary and
secondary alcohol groups of cutin monomers, and another enzymatic or nonenzymatic mechanism of polymerization may coexist
with CUS1-catalyzed polymerization. This mechanism is poorly efficient with secondary alcohol groups and produces polyesters
with lower molecular size. Confocal Raman imaging of benzyl etherified cutins showed that the polymerization is heterogenous at
the fruit surface. Finally, by comparing tomato mutants either affected or not in cutin polymerization, we concluded that the level of
cutin cross-linking had no significant impact on water permeance.
Cuticles are ubiquitous hydrophobic barriers at the
surfaces of aerial plant organs. These complex hydrophobic assemblies consist of a biopolymer, cutin, coated
and filled with waxes and can also comprise embedded
1
This work was supported by the Institut National de la Recherche Agronomique and the Région Pays de la Loire to G.P.
* Address correspondence to [email protected].
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.plantphysiol.org) is:
Bénédicte Bakan ([email protected]).
G.P. designed the experiments with inputs from B.B., D.M., and
C.R., performed the experiments with support from C.G., N.G., M.D.,
and J.P., and analyzed the data with extensive inputs from B.B., D.M.,
and C.R.; J.P., C.B., and J.-P.M. performed screening of the EMS
Micro-Tom mutant collection and generated RNAi VWA106 mutants; C.G. carried out Raman microspectroscopy analyses and interpreted the data with inputs from D.M. and B.B.; B.B. performed water
permeance experiments with inputs from R.F. and L.S.; N.G. and
M.D. were involved in immunoblotting and lipid analyses; B.B. conceived the original research plan, supervised the experiments, and
wrote the article with the contribution of all the authors.
www.plantphysiol.org/cgi/doi/10.1104/pp.15.01620
cell wall polysaccharides. Waxes comprise solventsoluble aliphatic molecules with long hydrocarbon
chains, terpenes, and steroids (Kunst and Samuels,
2003; Nawrath, 2006; Pollard et al., 2008; Samuels et al.,
2008; Schreiber, 2010; Lee and Suh, 2015). Cutin is an
insoluble polyester of v- and mid-chain hydroxy C16 and
C18 fatty acids. Glycerol has also been described as a
ubiquitous cutin monomer (Graça et al., 2002; Pollard
et al., 2008). In some cuticles, a hydrophobic polymer
that is resistant to alkaline hydrolysis (i.e. cutan) has
been observed (Gupta et al., 2006; Li-Beisson et al., 2010).
Cutin fulfills multiple functions in plants, such as the
control of nonstomatal water loss (Sieber et al., 2000)
and the permeation of gases and solutes (Kersteins,
1996; Schreiber, 2010). Cutin also plays an essential role
in the regulation of cell adhesion during plant development by preventing organ fusion, as observed in
Arabidopsis (Arabidopsis thaliana) mutants with cuticle
defects (Sieber et al., 2000; Nawrath, 2006), or by participating in hull adhesion in grains (Taketa et al., 2008).
Finally, it is generally accepted that plant cuticle and
its polymeric skeleton, cutin, are primary barriers to
pathogens and that cutin monomers released by fungal
Plant PhysiologyÒ, February 2016, Vol. 170, pp. 807–820, www.plantphysiol.org Ó 2016 American Society of Plant Biologists. All Rights Reserved.
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807
Philippe et al.
cutinase are signaling molecules for both the pathogen
and plants (Gilbert et al., 1996; Schweizer et al., 1996;
Iwamoto et al., 2002; Yeats and Rose, 2013).
The biological functions of cutin are closely controlled by its structure, which is determined by its
monomer composition and by the number and position
of its ester bonds. Cutin monomer composition can
vary according to plant species, developmental stage
(Baker et al., 1982; Peschel et al., 2007; Mintz-Oron et al.,
2008), organs, and environmental stress (Espelie et al.,
1979; Li-Beisson et al., 2009; Panikashvili et al., 2009;
Bessire et al., 2011). Actually, cutin monomer composition determines the total number of hydroxyl (OH)
groups that are potentially available for the formation
of ester bonds and, therefore, the cross-linking of the
polyester (Bonaventure et al., 2004; Franke et al., 2005;
Peschel et al., 2007). The nonesterified OH groups enhance the hydrophilic character of the cutin polymer,
increasing its elasticity (Bargel and Neinhuis, 2005).
Whereas cutin monomer composition has been described extensively for different plant species, organs,
and development stages, the macromolecular structure
of the cutin polyester has been much less thoroughly
investigated. In particular, the connectivity between the
monomers is a key point for understanding the threedimensional expansion of the polyester in relation to the
polymerization process. Different approaches have been
proposed to delineate the polymeric architecture of cutin.
Linear dimers were identified after partial alkaline hydrolysis of tomato (Solanum lycopersicum) cutin (Osman
et al., 1995). NMR and mass spectrometry analyses
of oligomers released after partial depolymerization
revealed primary and secondary ester linkages between
cutin monomers (Graça et al., 2002; Stark and Tian, 2006)
as well as covalent linkages between some cutin OH fatty
acids and oligosaccharides (Tian et al., 2008). However, it
has been shown that partial hydrolysis does not necessarily release all of the representative building blocks of
the entire polymer (Deshmukh et al., 2003). Spectrometric analyses have also been developed for the polymer.
Attenuated total reflectance (ATR)-Fourier transform
infrared (FTIR) spectroscopy analyses of the methylene
and carbonyl stretching vibrations allowed the estimation of an ester cross-linking index for cutin but could not
differentiate the primary from the secondary ester linkages (Girard et al., 2012; Heredia-Guerrero et al., 2014).
NMR studies have provided evidence of both v- and
mid-chain esters in tomato (Deshmukh et al., 2003).
In this regard, tomato fruit has proved to be an interesting model for structural studies of the cutin polymer. Indeed, its astomatous cuticle can be easily
isolated and is devoid of cutan. Moreover, tomato cutin
composition is dominated by a monomer with two OH
groups, 9(10),16-dihydroxyhexadecanoic acid (Baker
et al., 1982; Osman et al., 1999; Deshmukh et al., 2003).
Accordingly, the cutin monomers can be linked by
either a linear (on the primary OH) or a branched (with
a secondary OH) pattern. Previous studies have demonstrated that both linear and branched cross-links occur in
tomato cutin. However, the relative proportion of the
linear versus branched esters remains a matter of debate.
Oxidation experiments have indicated that almost all of
the primary cutin OH groups (94%) were involved in
ester bonds, whereas only 44% of the secondary OH
groups were esterified in the cutin polymer (Deas and
Holloway, 1977; Kolattukudy, 1977). Conversely, partial
depolymerization coupled with NMR studies of the released oligomers indicated that the branched secondary
esters were the major form of tomato cutin (Graça and
Lamosa, 2010). In addition, none of these studies could
decipher the ester links of the glycerol OH groups.
Additionally, the role of a GDSL lipase, involved in
cutin polymerization, was recently reported using two
different experimental approaches (Girard et al., 2012;
Yeats et al., 2012). The corresponding GDSL lipase,
named SlGDSL1 (Girard et al., 2012) or SlGDSL2 (Yeats
et al., 2012), is now named CUTIN SYNTHASE1
(SlCUS1; Yeats et al., 2014). Different mutants affected
in the expression of SlCUS1 have been generated and
constitute attractive tools to delineate the structure of
the cutin polymer (Girard et al., 2012; Petit et al., 2014).
It has been further demonstrated that 2-monoacylglycerol (2-MAG), a putative precursor of the cutin
polymer in Arabidopsis (Yang et al., 2010), can be used
by a heterologously expressed SlCUS1 (Yeats et al.,
2012) to produce in vitro linear oligomers in aqueous
solution (Yeats et al., 2014). Nevertheless, the question
of the mechanism of cutin polymerization in planta is
still open. Indeed, SlCUS1 is specifically localized
within the cutin matrix (i.e. a hydrophobic environment; Girard et al., 2012; Yeats et al., 2012), which could
impact the acyltransferase activity of the enzyme as
observed previously for lipases (Sharma et al., 2001).
By coupling O-alkylation of the nonesterified OH
groups of glycerol and fatty acids in an isolated cutin
matrix and by further analyses of O-alkylated and
nonalkylated monomers released after depolymerization, we elucidated the ester cross-link pattern of
tomato cutin. We also showed at two stages of fruit
development and in two different genetic backgrounds
that the modulation of SlCUS1 protein level, either
through RNA interference (cherry tomato ‘West Virginia 106’ [WVa106]) or mutagenesis (miniature tomato
‘Micro-Tom’), resulted in a strong alteration of the cutin
ester cross-link pattern. These results give new insights
into the polyester structure. In addition, while CUS1
esterification involves mostly primary OH groups in
vitro (Yeats et al., 2014), our data here indicate that, in
planta, deficiencies in CUS1 also affect the secondary
OH group of 9(10),16-dihydroxyhexadecanoic acid and
both the primary and secondary OH groups of glycerol.
RESULTS
Labeling of the Nonesterified OH Groups from Cutin in
the Wild Type and cus1 Mutants
To investigate the function of SlCUS1 in cutin polymerization, we used a series of transgenic and mutant lines obtained in two different tomato genetic
808
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Cutin Synthase Impacts Ester Cross-Links of Tomato Cutin
backgrounds. The first set of lines was generated as
described previously (Girard et al., 2012) in the cherry
tomato ‘WVa106’. Cherry tomatoes (var cerasiforme) are
considered close to the ancestral form of the cultivated
tomato (Ranc et al., 2008). The series consists of four
selected transgenic lines in which the down-regulation
of SlCUS1 using RNA interference results in mild to
severe alterations in cuticle thickness and cutin monomer deposition (Pro-35S:SlCUS1RNAi lines L4, L10, L3,
and L17; Girard et al., 2012). The second set of lines was
generated in the miniature cv Micro-Tom by ethyl
methanesulfonate (EMS) mutagenesis (Just et al., 2013;
Petit et al., 2014). The cus1-a mutant isolated previously
by map-based cloning carries a splicing site mutation in
the SlCUS1 gene, resulting in a strong reduction in fruit
cuticle thickness and cutin monomer deposition (Petit
et al., 2014). Two other cus1 mutants, cus1-b and cus1-c,
were identified by TILLING (Okabe et al., 2011). The
cus1-b mutant carries an Arg-229 to Leu-229 missense
mutation, whereas the cus1-c mutant carries an Asp-264
to Asn-264 missense mutation (Supplemental Fig. S1).
None of these mutations, which could potentially affect
the function of the protein, are located within the five
GDSL lipase conserved domains. In addition, they did
not affect fruit cuticle thickness (data not shown) or the
amount of cutin monomers (Table I) and, therefore,
could be used as negative controls. Analysis of SlCUS1
protein levels in the cuticle of growing fruits at 20 DPA
confirmed that the most affected Pro-35S:SlCUS1RNAi
lines were L3 and L17 and that the only cus1 mutant
lacking SlCUS1 was cus1-a (Fig. 1A).
ATR-FTIR spectroscopic analysis of the cutin matrix
isolated from red ripe fruits returned a ratio of the intensities of methylene and carbonyl stretching vibrations, also called the esterification index (Girard et al.,
2012; Heredia-Guerrero et al., 2014). Actually, SlCUS1
down-regulation induced a decrease in cutin density that
resulted in a decrease in both methylene and carbonyl
bands (Girard et al., 2012). Consequently, the esterification index of L3, a less affected transgenic line than L17,
was not altered significantly. Conversely, in the most
affected lines (L17 and cus1-a), this ratio was higher for
these SlCUS1 mutants than for the corresponding wild
types and the nonphenotypically affected SlCUS1 mutants, cus1-b and cus1-c (Fig. 1B). These spectrometric
analyses suggest a lower proportion of ester crosslinking for the most affected lines L17 and cus1-a cutins.
Free (i.e. nonesterified) OH groups were chemically
labeled, by O-alkylation, in the cutin of tomato skins of
the different wild-type and mutant fruits. By using
2-benzyloxypyridine alkylation, the OH groups are transformed to benzyl ether adducts (Lopez and Dudley, 2008).
The corresponding benzyl ether derivatives of glycerol
and major tomato hydroxy fatty acids can be separated,
identified, and quantified by gas chromatography-mass
spectrometry (GC-MS; Supplemental Fig. S2). This procedure revealed the esterification levels of the primary
and secondary OH groups of the major cutin monomers
of the cutin polyester in various cus1 mutants and their
corresponding wild-type lines.
CUS1 Protein Knockdown and Glycerol Esterification of
the Cutin Polyester
Glycerol is a ubiquitous cutin monomer (Graça et al.,
2002; Graça and Lamosa, 2010). Whatever the developmental stage, the glycerol content of cutin varies
from about 2 mg cm22 in cv WVa106 to about 4 mg cm22
in cv Micro-Tom (Table I). These glycerol contents are
slightly below the range found previously (i.e. from 5 to
8 mg cm22) in the cuticle of different plants, including
tomato fruit (Graça et al., 2002). However, an unreported significant variability of glycerol content within
tomato cultivars must be mentioned here, even though
glycerol is still a minor cutin component in both tomato
cultivars. Regarding the formation and structure of the
cutin polyester, it is important to also consider the
molar ratio between glycerol and the major hydroxy
fatty acids, such as 9(10),16-dihydroxyhexadecanoic
acid (Table II). Indeed, this ratio is less variable between
genetic backgrounds and developmental stages, suggesting a common polymerization mechanism.
Furthermore, we observed in both cultivars that
nearly 90% of the OH groups from glycerol are esterified in wild-type 20-DPA and red ripe fruit cutins (Fig.
2; Supplemental Fig. S3). Although glycerol is a minor
cutin monomer, this result is in agreement with a significant role of glycerol in the structure of the cutin
polymer, as suggested previously (Graça et al., 2002;
Pollard et al., 2008). In addition, approximately 10% of
residual nonesterified OH groups were observed in both
the sn-1,3 and sn-2 positions of glycerol in wild-type tomato fruit cutins. In CUS1 tomato mutants, the amount of
glycerol per surface area is less affected than the hydroxy
fatty acid content, which decreases sharply (Table I).
However, considering the molar content of cutin
monomers, it is noteworthy that the glycerol-to-hydroxy
fatty acid molar ratio is higher in the cus1-a mutant
and in most affected Pro-35S:SlCUS1RNAi lines compared
with the corresponding wild-type plants (about 6- to
10-fold; Table II). Furthermore, glycerol esterification is
impacted, as evidenced by an increase of up to 5-fold in
the amount of nonesterified glycerol OH in the mutant
lines. Moreover, the increase in the free OH groups of
glycerol was closely related to the severity of SlCUS1
protein knockdown (Figs. 1A and 2).
Effect of SlCUS1 Protein Knockdown on Mid-Chain OH
Esterification of Cutin Fatty Acids
The 9(10),16-dihydroxyhexadecanoic acid is the main
cutin monomer in both tomato cultivars, which constitutes more than 80% of fruit cutin hydroxy fatty acids
(Table I). Consequently, we focused on this fatty acid in
this study (Fig. 3; Supplemental Fig. S4). Similar esterification levels were observed for both cv WVa106 and cv
Micro-Tom. Indeed, the primary OH group of this fatty
acid, in the v-position, is preferentially esterified in accordance with previous structural data (Deshmukh et al.,
2003; Fig. 3). A noticeable increase in the esterification
of the secondary OH groups was observed between the
Plant Physiol. Vol. 170, 2016
809
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
20 DPA
Glycerol
p-Coumaric acid
m-Coumaric acid
Hexadecanoic acid
1,16-Hexadecanedioic acid
16-Hydroxyhexadecanoic acid
9(10),16-Dihydroxyhexadecanoic acid
9-Hydroxy-1,16-hexadecanedioic acid
Octadecanoic acid
18-Hydroxyoctadecanoic acid
9(10),18-Dihydroxyoctadecanoic acid
9,10-Epoxy-18-hydroxyoctadecenoic acid
9,10-Epoxy-18-hydroxyoctadecanoic acid
9,10,18-Trihydroxyoctadecanoic acid
24-Hydroxytetracosanoic acid
Unidentified compounds
Red Ripe
Glycerol
p-Coumaric acid
m-Coumaric acid
Hexadecanoic acid
1,16-Hexadecanedioic acid
16-Hydroxyhexadecanoic acid
9(10),16-Dihydroxyhexadecanoic acid
9-Hydroxy-1,16-hexadecanedioic acid
Octadecanoic acid
18-Hydroxyoctadecanoic acid
9(10),18-Dihydroxyoctadecanoic acid
9,10-Epoxy-18-hydroxyoctadecenoic acid
9,10-Epoxy-18-hydroxyoctadecanoic acid
9,10,18-Trihydroxyoctadecenoic acid
9,10,18-Trihydroxyoctadecanoic acid
24-Hydroxytetracosanoic acid
Unidentified compounds
Compounds
2.0
0.2
0.5
0.8
0.8
0.2
29.1
1.5
1.1
0.2
1.5
0.9
1.8
3.4
0.0
3.8
1.4
0.9
4.7
0.8
0.1
2.2
89.5
4.4
0.1
0.3
1.5
0.4
0.1
0.3
0.4
1.3
2.2
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
4.5
2.5
5.2
1.8
1.8
22.2
537.8
35.1
2.7
0.8
4.9
4.0
8.4
6.1
0.0
17.9
4.2
4.8
15.1
1.8
3.3
21.1
588.4
19.7
1.6
1.0
10.4
1.2
0.8
1.6
2.9
1.4
42.2
Wild Type
7.3
4.0
13.5
4.1
0.3
1.5
135.1
6.4
3.0
0.3
2.7
0.8
3.8
0.0
2.8
4.2
25.5
7.9
1.9
3.4
0.8
0.2
0.9
93.4
5.5
1.1
0.1
1.1
0.9
5.8
2.8
1.9
8.1
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
0.9*
1.2
2.1
0.3
0.2
0.4
7.4*
1.0
1.7
0.1
0.0
0.2
1.1
0.0
0.1
0.9
5.7
0.7*
0.0
0.9
0.2
0.1
0.0
0.4**
0.0
0.3
0.0
0.2
0.1
0.4
0.8
0.0
2.7
cus1-a
4.5
3.0
13.1
2.1
2.2
23.3
730.2
23.5
2.0
1.1
11.5
1.6
1.0
1.9
4.9
0.6
50.7
5.2
2.5
6.2
2.0
1.3
23.6
609.9
22.8
2.5
1.1
8.5
6.1
9.5
3.9
0.0
17.3
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
0.7
0.1
1.0
0.2
0.1
1.3
45.2
2.8
0.2
0.1
0.0
0.2
0.1
0.1
0.2
0.0
0.2
0.5
0.5
0.4
0.6
1.7
1.2
39.5
2.3
0.4
0.2
2.6
2.1
2.7
1.9
0.0
11.4
cus1-b
cv Micro-Tom
3.6
1.5
9.3
1.7
1.7
12.9
511.5
11.3
1.6
0.6
6.6
0.9
0.8
1.3
2.5
0.4
33.3
3.1
2.3
5.0
1.7
1.4
20.5
511.4
27.4
2.9
0.8
4.0
6.1
10.6
5.0
0.0
22.2
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
1.4
0.1
2.4
0.4
0.2
0.5
10.3
2.6
0.6
0.0
0.2
0.0
0.1
0.1
0.1
0.1
0.0
0.7
0.2
0.4
0.6
0.3
2.9
16.4
4.9
1.1
0.2
0.7
1.4
1.9
2.0
0.0
10.4
cus1-c
1.9
1.6
4.5
2.1
2.0
23.0
571.5
32.5
2.2
1.3
7.5
0.8
0.9
1.3
4.9
0.5
34.4
2.4
2.7
2.7
1.2
0.9
18.4
523.0
44.8
1.4
1.0
3.5
1.5
4.3
3.5
0.3
33.8
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
0.0
0.5
0.7
0.2
0.1
3.3
6.4
10.0
0.1
0.1
1.3
0.1
0.0
0.7
1.0
0.1
14.8
0.1
1.8
0.7
1.1
0.1
0.8
28.2
7.6
0.8
0.4
0.0
1.6
0.4
0.2
0.1
10.2
Wild Type
2.1
1.9
5.4
2.1
1.6
23.2
550.5
32.9
1.5
1.4
5.9
1.3
1.4
2.0
4.9
0.6
39.7
3.5
1.1
1.7
1.1
2.1
13.0
450.1
37.1
2.1
0.5
3.2
1.7
5.9
2.1
0.5
27.2
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
L4
0.0
0.3
1.1
0.2
0.2
1.2
27.4
0.4
0.3
0.2
0.7
0.2
0.2
0.3
2.7
0.1
1.5
0.6
0.0
0.1
0.5
0.3
0.2
22.5
0.9
1.2
0.1
0.3
0.0
0.4
0.2
0.0
0.3
2.3
1.3
4.2
1.0
0.6
11.4
459.1
24.8
1.5
0.8
7.4
0.9
1.8
1.8
4.6
0.6
36.6
2.6
0.9
1.4
0.6
0.5
9.8
348.7
32.4
1.1
0.3
2.3
0.4
4.9
1.8
0.9
22.3
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
L10
0.7
0.2
0.2
0.0
0.2
3.1
16.4*
3.0
0.0
0.1
1.7
0.0
0.6
0.5
0.2
0.0
2.3
1.0
0.2
0.4
0.1
0.3
4.7
37.4*
3.3
0.5
0.1
0.3
0.2
1.5
0.0
0.9
0.9
cv WVa106
1.8
0.3
0.7
1.1
0.1
1.1
114.9
3.6
0.6
0.1
1.7
0.4
0.0
0.0
3.5
12.3
31.3
3.9
0.5
0.3
0.6
0.1
1.9
141.9
9.9
0.8
0.1
0.8
0.2
7.0
1.6
1.9
10.1
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
0.2
0.1
0.1
0.2
0.0
0.3
11.2***
0.6
0.0
0.0
0.1
0.0
0.0
0.0
0.6
0.8
1.0
0.2**
0.0
0.0
0.2
0.0
0.1
0.6**
0.2
0.3
0.0
0.0
0.0
0.2
0.0
0.2
0.1
L3
1.9
0.2
0.5
1.0
0.0
0.6
86.0
2.4
0.5
0.1
1.4
0.3
0.0
0.0
2.9
11.1
25.4
3.8
0.3
0.3
1.0
0.0
1.1
83.4
5.4
1.3
0.1
0.7
0.1
5.7
2.4
1.8
3.4
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
0.1
0.0
0.0
0.0
0.0
0.0
1.8***
0.1
0.1
0.0
0.2
0.0
0.0
0.0
1.1
0.7
0.2
0.8
0.2
0.0
0.3
0.0
0.0
9.9**
1.8
0.6
0.0
0.3
0.1
1.6
0.5
0.4
0.5
L17
Values 6 SD are expressed in mg cm22 and are means of at least three biological replicates. Significant differences for glycerol and 9(10),16-dihydroxyhexadecanoic acid (boldface) from the
corresponding wild type are indicated with asterisks (Student’s t test; *, P , 0.05; **, P , 0.01; and ***, P , 0.001).
Table I. Cutin monomer composition of 20-DPA and red ripe fruits in the wild type, Pro-35S:SlCUS1RNAi lines (L4, L10, L3, and L17), and cus1 mutants (cus1-a, cus1-b, and cus1-c)
Philippe et al.
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Plant Physiol. Vol. 170, 2016
Cutin Synthase Impacts Ester Cross-Links of Tomato Cutin
Figure 1. Characterization of tomato cutin
in cus1 mutants and Pro-35S:SlCUS1RNAi
lines. A, Immunoblot analysis of the SlCUS1
protein in 20-DPA fruits. B, Nonnormalized
areas of methylene (CH2; 2,978–2,838 cm21)
and carbonyl (CO; 1,750–1,690 cm21)
bands, and esterification index, the ratio of
CH2 to CO areas in red ripe fruits. Left, cus1
mutants; right, Pro-35S:SlCUS1RNAi lines. WT,
Wild type. Mean values 6 SD were calculated
from 32 independent measurements (four
biological samples and eight technical replicates each). Significant differences from the
wild type are indicated by asterisks (Student’s
t test; **, P , 0.01).
20-DPA (Fig. 3A) and red ripe (Fig. 3B) stages, especially
in cv WVa106. This result is in accordance with previous
FTIR and NMR data, suggesting an increase in ester bond
cross-links during tomato fruit development (Benítez
et al., 2004).
Remarkably, whatever the cultivar, SlCUS1 knockdown resulted, in planta, in a specific reduction of mid-chain
OH esterification of the 9(10),16-dihydroxyhexadecanoic
acid, whereas esterification of the v-OH group was not
affected.
Surface Distribution of the Nonesterified OH Groups in
the Cutin Polymer
Benzyl etherification of free OH groups gives rise to a
specific Raman band at 1,001 cm21 that was not retrieved
in the nonlabeled cutin (Supplemental Fig. S5). Moreover, Raman microspectroscopy has a high resolution
that fits with structural characterization of cuticle surfaces (Littlejohn et al., 2015). Therefore, Raman imaging
at the specific 1,001-cm21 band was used to monitor the
spatial distribution of free OH groups within the cutin
polymer. High-resolution confocal images revealed that
free OH groups were not evenly distributed (Fig. 4, A
and B) within the wild-type cutin polymer. Indeed, a
weaker signal (i.e. a higher cross-link level) was observed
in the cutin ridge at the midpoint in the cell junctions
than at its external part. Interestingly, in a previous work
(Girard et al., 2012) using SlCUS1 immunolocalization,
we observed a sharp protein signal in the region that
corresponded to the central part of the cutin ridge. In
contrast, in the Pro-35S:SlCUS1RNAi line L17, a sharper
and homogenous signal was observed (Fig. 4, C and D).
Water Permeance of Cutins from Wild-Type and
cus1 Tomato
Water permeability is an important feature related to
the biological function of the cuticle. Waxes and, in
particular, wax composition have been described as
major factors involved in cuticle permeability (Riederer
and Schreiber, 2001; Leide et al., 2007), although the
cutin matrix contributes to the transport properties of
the cuticle. In particular, it was suggested that a decrease in the permeability of chloroform-extracted fruits
observed during the development of cv Micro-Tom
fruit could be due to an increase in ester cross-links of
the cutin monomer (Leide et al., 2007).
Additionally, in a previous study, cv M82 tomato
fruits affected in SlCUS1 expression had a higher transpiration rate than its background genotype (Isaacson
et al., 2009). This experiment was conducted on whole
fruit, and it could not be deciphered whether this higher
water permeability was associated with the observed
modifications of cutin or with the wax composition.
Accordingly, the water permeance of isolated and
enzyme-treated cuticle and cutin from wild-type and
EMS mutant fruit were compared to assess the impact
of polymerization. The tomato mutant cus1-a had a
sharply reduced thickness. While comparing cuticle
Table II. Molar ratio of glycerol to 9(10),16-dihydroxyhexadecanoic
acid within cutin from 20-DPA and red ripe fruits from the wild type,
Pro-35S:SlCUS1RNAi lines (L4, L10, L3, and L17), and cus1 mutants
(cus1-a, cus1-b, and cus1-c)
Values are multiplied by 100. Significant differences from the corresponding wild type are indicated by asterisks (Student’s t test; *, P ,
0.05; **, P , 0.01; and ***, P , 0.001).
Cultivar
WVa106
Micro-Tom
Line
Wild type
L4
L10
L3
L17
Wild type
cus1-a
cus1-b
cus1-c
Plant Physiol. Vol. 170, 2016
20 DPA
1.5
2.5
2.4
8.8
14.9
2.9
26.4
2.6
2.1
6
6
6
6
6
6
6
6
6
0.0
0.4
1.2
0.4**
5.3*
1.3
3.1*
0.2
0.3
Red Ripe
1.0
1.2
1.6
5.1
6.6
2.5
15.9
1.8
1.8
6
6
6
6
6
6
6
6
6
0.1
0.1
0.4
0.1***
0.1***
0.1
0.6**
0.1*
0.6
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Philippe et al.
Figure 2. Esterification levels of glycerol OH groups of tomato fruit cutin isolated from the cus1-a mutant and Pro-35S:
SlCUS1RNAi lines. A and C, Fruits at 20 DPA. B and D, Red ripe fruits. Left, cus1-a; right, Pro-35S:SlCUS1RNAi lines. WT, Wild type.
Mean values 6 SD were calculated from nine independent measurements (three fruits and three technical replicates each).
Significant differences from the corresponding wild type are indicated by asterisks (Student’s t test; *, P , 0.05 and **, P , 0.01).
from different species, it was concluded that there is no
direct correlation between cuticle permeability and
cuticle thickness (Riederer and Schreiber, 2001). Nevertheless, when comparing cutin with similar monomer
composition, a thickness effect could not be ruled out.
Another EMS tomato mutant, cutin-deficient1 (cud1),
highlighted a thinner cutin with similar cutin composition than that shown previously (Petit et al., 2014) and
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Cutin Synthase Impacts Ester Cross-Links of Tomato Cutin
Figure 3. Esterification levels of 9(10),16-dihydroxyhexadecanoic acid [9(10),16-diOHC16 acid] OH groups of tomato fruit cutin
from the cus1-a mutant and Pro-35S:SlCUS1RNAi lines. A and C, Fruits at 20 DPA. B and D, Red ripe fruits. Left, cus1-a; right,
Pro-35S:SlCUS1RNAi lines. WT, Wild type. Mean values 6 SD were calculated from nine independent measurements (three fruits
and three technical replicates each). Significant differences from the corresponding wild type are indicated by asterisks (Student’s
t test; *, P , 0.05; **, P , 0.01; and ***, P , 0.001).
confirmed in this study (Fig. 5A), without a defect in
ester cross-linking according to the labeling of the free
OH derivatization (Fig. 5B). Accordingly, cud1 was
used as a control in the water permeance experiments. The water permeance of the isolated enzymetreated cuticle from the different cutin mutants, either
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Philippe et al.
Figure 4. Raman mapping of nonesterified OH
groups within cutin polyester from tomato.
Cutin from wild-type (WT; A and B) and
Pro-35S:SlCUS1RNAi line L17 (D and D) red ripe
fruit were benzyl etherified and analyzed by Raman
microspectroscopy. Each image was obtained by
mapping the characteristic band of aromatic groups
at 1,001 cm21. Images B and D correspond to higher
magnification views (dashed squares) of the Raman
images (A and C, respectively).
affected (cus1-a) or not (cud1) in cutin cross-linking, was
compared (Fig. 6A), and no significant differences were
observed between the different cv Micro-Tom mutants
and the wild type. Conversely, the median water permeance of the enzyme-treated dewaxed cuticle of the
cus1-a mutant was significantly (P , 0.05) higher than
the permeance of the wild type as well as the cus1-b and
cus1-c cutin (Fig. 6B). However, the water permeances
of the cud1 and cus1-a cutins were not statistically different, while both mutants highlighted a reduced cutin
thickness compared with the wild-type fruits (Petit
et al., 2014). These data suggest that modification of the
polymerization pattern of cus1-a cutin is not the major
factor that explains its higher cutin water permeance.
DISCUSSION
SlCUS1 Catalysis Impacts the Ester Cross-Linking of
Fruit Cutin
The cutin polyester is essential for the functional
properties of plant cuticles (Nawrath, 2006; Schreiber,
2010; Yeats and Rose, 2013). While cutin monomer
biosynthesis is now relatively well described (Li-Beisson
et al., 2010, 2013; Beisson et al., 2012), the extracellular
assembly of the monomers in a three-dimensional network is less documented. In this regard, the role of
SlCUS1, a member of the GDSL lipase, expressed in the
tomato fruit exocarp, on cutin polymerization, has been
elucidated by two concomitant independent studies
(Girard et al., 2012; Yeats et al., 2012). GDSL lipases are
ubiquitous proteins in the plant kingdom suggesting a
CUS1-like conserved pathway for cutin polymerization
(Yeats et al., 2014). With the discovery of the key function
of glycerol-3-phosphate acyltransferases in cutin deposition (Li et al., 2007), it has been hypothesized that
2-MAG could be involved in glycerol incorporation within
cutin polymer. Besides, it has been shown, in vitro, that
these 2-MAGs produced by glycerol-3-phosphate acyltransferases are substrates of SlCUS1 (Yeats et al., 2012,
2014; Yeats and Rose, 2013). Consequently, a mechanism
involving the self-transesterification of 2-MAG precursors by CUS1 has been proposed, leading first to the
formation of a 9(10),16-dihydroxyhexadecanoyl dimer
and then, in a cascade, to oligomers of higher molecular
mass. This mechanism implies the release of glycerol and
is consistent with the low level of glycerol in tomato
cuticles and, indeed, in most plant cutins (Yeats et al.,
2014). This catalysis involves only the primary OH
group of the dihydroxy fatty acid and leaves the medium
chain OH nonesterified, as demonstrated by the NMR
analysis of the oligomers produced in vitro.
In this study, we observed a lower molar proportion of
glycerol and, in particular, a lower molar ratio of glycerol
to OH fatty acids in the wild-type cutin than in the
SlCUS1 cutin of knockdown lines and cus1-a mutants.
Considering the proposed polymerization mechanism,
in which the residual glycerol is due to a residual monoglyceride unit during polyester growth (Yeats et al.,
2014), these results may indicate that polyesters of
shorter chain length are formed in the cutins of cus1-a
mutants than in those of wild-type fruits. Remarkably, in
these mutants, we also observed a significant decrease in
the esterification level of the secondary OH group of the
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Cutin Synthase Impacts Ester Cross-Links of Tomato Cutin
Figure 5. The cutin-deficient mutant cud1 is not affected in the polymerization of 9(10),16-dihydroxyhexadecanoic acid. A, Quantification of
major cutin monomers. 16-OHC16 acid, 16-Hydroxyhexadecanoic
acid; 9(10),16-diOHC16 acid, 9(10),16-dihydroxyhexadecanoic acid;
9-OH-1,16-C16 diacid, 9-hydroxy-1,16-hexadecanedioic acid. B, Esterification level of 9(10),16-dihydroxyhexadecanoic acid OH groups from
cud1 cutin compared with wild-type (WT) and cus1-a cutins. Data are
expressed as mean values, and error bars represent SD (n = 3). Significant
differences from the wild type are indicated by asterisks (Student’s t test;
*, P , 0.05 and **, P , 0.01).
dihydroxy fatty acid. This decrease was associated with
a decrease of cutin polyester deposition, which means
that, from a quantitative point of view, both primary and
secondary OH group contents decreased significantly in
the cus1 mutants. Therefore, these results strongly support that, in planta, SlCUS1 is involved in cutin polymer
branching by carrying out the esterification of both primary and secondary OH groups. This mechanism was
verified in both 20-DPA and mature fruits and whatever
the genetic background of the mutant lines (i.e. cv MicroTom or cv WVa106). While cv Micro-Tom belongs to the
domesticated tomato group, the cherry-type tomato
cv WVa106 clusters with the wild species Solanum
pimpinellifolium (Ranc et al., 2008). These results suggest
that the function of SlCUS1 can be conserved across tomato
cultivars and species displaying diverse cuticle architecture
and composition (Ranc et al., 2008; Yeats et al., 2014). It is
noteworthy that the involvement of both the primary and
secondary OH groups is quite compatible with the release
of glycerol during the catalytic process and, therefore,
with a low proportion of glycerol in the cutin polyester.
Considering the contribution of glycerol to the
structure of cutin polymer, its OH groups are mostly
involved in the formation of ester bonds in wild-type
plants, whereas its esterification level decreases significantly in the Pro-35S:SlCUS1RNAi and cus1-a mutant.
The nonesterified OH groups of glycerol were observed
at both the sn-2 and sn-1,3 positions, and especially in
the mutants, the esterification of both positions was
impacted. Furthermore, a significantly higher impact
was observed on the sn-1,3 positions than on the sn-2
position. First, this means that the primary OH groups
of glycerol are involved, as expected, in the transesterification process and agree with the presence of
1-MAG in the cutin building blocks (Graça et al., 2002).
Second, these results strengthen the hypothesis that, in
tomato, 2-MAGs are precursors of the cutin polymer as
they are in Arabidopsis and, as discussed above, that
the polymer sizes of the polyesters are lower in cus1-a
mutants than in the wild type. Finally, an isomerization
from 2-MAG to 1,3-monoacylglycerol cannot be
excluded, as it was observed previously in vitro
for 2-MAG (Lyubachevskaya and Boyle-Roden, 2000;
Compton et al., 2012). The apparent discrepancy between
these results and previous in vitro experiments (Yeats
et al., 2014) can be easily explained by differences in the
environment of CUS1. Indeed, in vitro catalysis (Yeats
et al., 2014) was performed in an aqueous medium, while
in planta, the enzyme is specifically localized within the
cutin matrix (Girard et al., 2012; Yeats et al., 2012), which
means that the catalysis occurs in its native hydrophobic
environment. As observed for lipases in organic waterfree solvents (Sharma et al., 2001), this hydrophobic environment could favor the transesterification process,
thereby leading to the synthesis of high molecular mass
polyesters. These lipase-catalyzed esterifications are well
documented for both the synthesis of small esters and the
synthesis of high-Mr polyesters (Wescott and Klibanov,
1994; Krishna et al., 2002; Kobayashi, 2010). These
studies have clearly shown that the lipase-catalyzed esterification can involve both primary and secondary
alcohol groups, depending on the source of lipase and
environmental conditions (e.g. the temperature and the
type of organic solvent), and lipases immobilized on
polymer beads (e.g. acrylic resin) or grafted with
polyoxyethylene detergent. In particular, the impact
of solvent hydrophobicity on the regiospecificity of
lipase-catalyzed transesterification has already been
reported. Indeed, in the case of Pseudomonas cepacia
lipase, a shift from toluene to acetonitrile modified the
regioselectivity of the butanolysis of 1,4-dibutyryloxy2-octylbenzene, leading to either a 1-hydroxy or a
4-hydroxy compound (Rubio et al., 1991). Moreover,
although CUS1 is structurally different from these
microbial lipases, at least on the basis of the position of
the catalytic amino acid (i.e. Asp-Ser-Pro catalytic
triad) in the sequence, it is interesting to note that
CUS1 is localized in a hydrophobic environment
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Philippe et al.
Figure 6. Comparison of the water permeance of
tomato cutin in red ripe fruits from wild-type and
tomato EMS mutant plants. Data are shown as
median values 6 minimum/maximum (n = 8–10).
Differences between the wild type (WT), cus1
mutants (cus1-a, cus1-b, and cus1-c), and cud1
were tested with the Mann-Whitney U test. Asterisks indicate significant differences from the
wild type (*, P , 0.05).
controlled by wax deposition in which it is more or less
immobilized on polysaccharides.
Besides, the involvement of another protein activating CUS1, as occurs in the lipase-colipase system of the
pancreatic enzyme (Lowe, 1997), cannot be excluded.
Finally, although the same results were obtained in
two different genetic backgrounds and with two different ways to modulate CUS1 expression, the hypothesis
of an indirect effect of CUS1 down-regulation, such as an
adaptation phenomenon, on cutin structure cannot be
totally ruled out.
Changes in the Esterification in cus1 Mutants Reveal
Another Polymerization Mechanism
Other interesting and unexpected results include (1)
the sharp increase in the molar ratio of glycerol to
dihydroxy fatty acid in the SlCUS1-depleted mutants
compared with the corresponding wild-type fruit
cutins and (2) the lower esterification level of the secondary OH groups in both glycerol and dihydroxy fatty
acids. As suggested above, this means that a polyester
of lower molecular size is formed in the mutants than in
the wild-type plants. This suggests that, in the absence
of SlCUS1, another mechanism is involved in the
formation of the cutin polymer. Indeed, GDSL lipases
form a multigenic family of proteins, and more than
100 genes are found in the tomato genome (http://
solgenomics.net/). Therefore, additional GDSL lipases
with CUS activity could be expressed in the fruit epidermis, as already shown during the early stages of fruit
development (Yeats et al., 2010, 2014; Matas et al., 2011).
These enzymes would have a higher regiospecificity
toward primary OH groups. Since nonenzymatic transesterification processes preferentially use primary OH
groups (Benítez et al., 2015), such reactions cannot be
excluded. A self-assembly process of cutin monomers
has already been suggested for cutin formation
(Domínguez et al., 2015). Whatever the mechanisms
involved in the absence of SlCUS1 (i.e. enzymatic or
nonenzymatic), our results suggest that, in the wildtype plants, different mechanisms should participate
in the formation of the cutin polymer.
Benzyl Etherification: A Tool to Probe Cutin Ester
Cross-Link Heterogeneity
Derivatization within cutin polymers coupled with
Raman imaging is also a new tool to elucidate the
structural heterogeneity within the cutin biopolymer.
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Cutin Synthase Impacts Ester Cross-Links of Tomato Cutin
This heterogeneity has been suggested previously in
the literature (Stark and Tian, 2006) and was fully
confirmed by our data here. The most highly crosslinked areas are localized at the ridge between the
cells and coincide with high CUS1 immunolabeling
(Girard et al., 2012). This strengthens the role of CUS1 in
the extracellular polymerization of the cutin polymer.
In addition, this labeling procedure could also be applied to the study of cutin formation. Indeed, according
to different biochemical and spectrometric studies,
modification of the cross-links within the cutin polymer may be associated with the changes in cutin
composition observed during fruit development
(Baker et al., 1982; Benítez et al., 2004; Matas et al.,
2011). Such studies would indeed benefit from this
imaging approach.
The Ester Cross-Link Does Not Affect the Water
Permeance of Cutin
The question of the impact of cutin cross-linking on
the functional properties of cutin is still open. The tomato mutants with different levels of cross-linking that
were characterized in our study should be valuable
tools for further studies on the biological function of the
cutin.
A previous study has compared the water permeance
of intact, dewaxed, and peeled cv Micro-Tom red ripe
tomato in whole fruit (Leide et al., 2007). It was estimated that cutin accounted for approximatively 20% of
cuticle permeance. Although minor, this is a significant
contribution to cuticle permeability.
From our results, it can be concluded that the crosslinking pattern of the tomato cutin has a limited impact
on the water permeance of the polymer. Rather, cutin
water permeance seems to be associated with polymer
thickness. It is noteworthy that in both cus1-a and cud1
mutants, the molar ratio between glycerol and hydroxy
fatty acids is significantly higher than in the wild-type
plants (i.e. 0.159 6 0.006, 0.170 6 0.027, and 0.025 6
0.001, respectively). A potential effect of the esterified
glycerol on cutin permeability is not documented in the
literature but cannot be ruled out.
The water permeance of isolated cuticle was not
correlated with the cutin permeance, as no significant
difference in water permeance was highlighted between the cus1-a, cud1, and wild-type cuticles. The
median water permeances of the cuticles were much
lower than that of cutin, which fits with a major impact
of waxes on water permeance. It was shown previously that the wax load was higher in cus1-a (8.9 6
1.3 mg cm22) and cud1 (10.6 6 1.9 mg cm22) than in wildtype (6.2 6 0.7 mg cm22) red ripe tomato fruits (Petit
et al., 2014). This result suggests that the wax load could
have compensated for the higher cutin water permeance
of cus1-a and cud1. Waxes are either deposited on the
outer surface of the cuticle (epicuticular waxes) or embedded in the cutin polymer (intracuticular waxes), and
a recent study has established that epicuticular waxes
from Prunus laurocerasus leaves were not significantly
involved in the water-loss barrier properties of the cuticle (Zeisler and Schreiber, 2016). In this study, the ratio of
the intracuticular and epicuticular waxes of the isolated
tomato cuticles was not monitored but could explain
how the water permeance of cuticle was not associated
directly with the wax load.
CONCLUSION
The benzyl etherification of nonesterified OH groups
in cutin is a procedure that probes cutin polymerization
in planta by combining complementary GC-MS and
microspectroscopy imaging analyses of cutin building
blocks. The results confirm the major role for SlCUS1
in cutin polymerization in planta. SlCUS1 is involved
in a transesterification process that is compatible
with 2-MAG as the enzyme substrate and involves
both primary and secondary OH groups of 9(10),16dihydroxyhexadecanoic acid and glycerol. This enzymatic catalysis leads to glycerol release and, therefore, is
consistent with the low glycerol content of cutin polyester. This enzymatic process leads to a highly reticulated
network but, surprisingly, without a major relationship
with the water permeance of the corresponding cutins.
Finally, in the absence of SlCUS1, another enzymatic or
nonenzymatic mechanism that produces cutins with a
higher proportion of glycerol could take place. In wildtype plants, this mechanism could coexist with the
SlCUS1-catalyzed formation of the cutin polyester.
MATERIALS AND METHODS
Plant Materials and Isolation of Fruit Cuticles
The Pro-35S:SlCUS1RNAi lines generated in the cherry tomato (Solanum
lycopersicum var cerasiforme ‘WVa106’; L10, L4, L3, and L17) have been described previously (Girard et al., 2012). The cus1 cv Micro-Tom mutants were
identified from a tomato EMS mutant collection either by phenotypic screening
for P15C12 and renamed cus1-a (Petit et al., 2014) or by TILLING (P13E2
renamed cus1-b and P4H12 renamed cus1-c; Supplemental Fig. S1). The cutindeficient mutant P23F12 was identified from the same collection by phenotypic
screening (Petit et al., 2014) and was renamed cud1. All plants were grown in a
greenhouse under previously described conditions (Girard et al., 2012; Petit
et al., 2014).
Fruit skins were peeled and enzymatically digested in a solution of 1% (v/v)
cellulase and 2% (v/v) pectinase containing 1 mM NaN3. After drying, the
cuticles were exhaustively delipidated in CH3OH:CH2Cl2 (1:2) and dried for
analysis. These enzyme-treated and dewaxed cuticles are referred to as cutins
here.
For SlCUS1 immunoblot analysis, the proteins were extracted from 10 mg of
20-DPA fruit pericarps ground in liquid nitrogen. Proteins were extracted with a
50 mM Tris-HCl buffer (pH 8) containing 2% SDS and 1% mercaptoethanol.
After electroblotting, SlCUS1 was detected with an Immuno-Star AP kit (BioRad) as described previously (Girard et al., 2012).
ATR-FTIR and Atomic Force Microscopy
FTIR spectra (200 scans) were recorded by ATR as described previously
(Girard et al., 2012) on a Nicolet Magna IR 550 spectrometer using a single
reflection accessory fitted with a thermostatted diamond crystal with a 45°
angle of light incidence. Two spectra were acquired on four different parts of the
fruit cutin, and four fruits per line were analyzed. Surface calculations were
conducted from the nonnormalized spectra after a baseline correction, which
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Philippe et al.
was established in the same conditions using Galactic software (Thermo Scientific). The surfaces of the methylene (CH2; 2,978–2,838 cm21) and the carbonyl
(CO; 1,750–1,695 cm21) bands were measured and used to calculate the ratio
RICH2/ICO.
Derivatization of Free OH Groups within Cutin Polyester
Free OH groups were derivatized by benzyl etherification by a procedure adapted from a procedure developed for organic alcohols (Lopez
and Dudley, 2008). Isolated cutins (3 mg) were mixed in a screw-capped
glass tube for 24 h with 15 mg of 2-benzyloxy-1-methylpyridinium triflate (Sigma-Aldrich) and 1.68 mg of magnesium oxide in 1 mL of trifluorotoluene at 90°C overnight. Cutin was then washed extensively with
CH 2 Cl 2 and dried. By comparing different sizes of cutin samples and
cutin powders, we verified that the same amount of derivatized cutin
monomer was obtained, indicating that cutin diffusion barriers did not
hinder the reaction.
Determination of Cutin Monomer Composition
Tomato cutins were depolymerized via methanolysis according to a previously described procedure (Molina et al., 2006). The reaction was performed
in dry methanol containing 15% (v/v) methyl acetate and 6% (v/v) sodium
methoxide, and 15-hydroxypentadecanoic acid was added to the reaction
medium as an internal standard. Complete depolymerization was obtained
after 6 h of heating at 60°C. Afterward, the mixture was cooled, and it was
adjusted to pH 5 with acetic acid. Fatty acids were extracted from the aqueous
mixture with methylene chloride, and the nonbenzylated OH groups of the
depolymerized fatty acids were silylated with 1% N,O-bis(trimethylsilyl)trifluoroacetamide/trimethyl chlorosilane and analyzed by GC-MS and gas
chromatography-flame ionization detection as described previously (Girard
et al., 2012).
The mass spectrometry spectra of derivatized cutin monomers were characterized by their fragmentation and the ion of mass 91 corresponding to the
benzyloxy group (Supplemental Fig. S2).
Glycerol Quantification
Glycerol was released by mild methanolysis using a modified procedure
(Graça et al., 2002). Isolated cutin slices were stirred at room temperature in a
mixture of 50 mM sodium methoxide in dry methanol with the internal
standard 1,2,3-butanetriol (Shen and Xu, 2013). We used different reaction
times to determine the time necessary to attain a plateau for glycerol release.
After 20 h, the extract was dried with a nitrogen flow, silylated with 1% N,
O-bis(trimethylsilyl)trifluoroacetamide/trimethyl chlorosilane, and analyzed by GC-MS and gas chromatography-flame ionization detection.
Raman Microspectrometry Imaging
Cutin samples (approximately 2 cm2) were placed on aluminum supports
that were mounted on the motorized xyz stage of the InVia Raman spectrometer (Renishaw) equipped with confocal capability, a 785-nm diode laser
(50 mW), and a 1,200 L mm21 grating. The instrument wavelength was
calibrated with silicon at 520 cm21. Spectra were collected with an accumulation time of 0.5 s over a fixed window of 1,065 cm21 (ranging from 838 to
1,902 cm21) every 2 mm over a line map of 100 to 200 mm. The instrument was
operated using a 503 or 1003 objective, a 65-mm slit, and a 1,040- 3 256-pixel
Renishaw CCD camera. Each data set was analyzed using the Renishaw
WIRE 4.1 software.
Measurement of Water Permeance of Tomato Fruit Cutin
Purified tomato fruit cutins (approximately 1.5 cm2) from wild-type and EMS
mutant cv Micro-Tom plants were used for water permeability experiments
using a gravimetric method as described previously (Schreiber and Riederer,
1996; Riederer and Schreiber, 2001). Isolated cuticles or cutin (eight to 10
samples of each mutant) were placed in transpiration chambers filled with
300 mL of deionized water. The amount of evaporated water across the cuticle
was monitored as a function of time to achieve a linear regression. Permeances
(m s21) were calculated from the slope of this linear regression. Water permeances were presented as medians with 25% and 75% quartiles.
Sequence data from this article can be found in the GenBank/EMBL data
libraries under accession numbers solyc11g006250; Solgenomics, SGNU585129; NCBI Gene ID, LOC101254153.
Supplemental Data
The following supplemental materials are available.
Supplemental Figure S1. EMS MicroTom mutants used this study.
Supplemental Figure S2. Benzyl etherification of non esterified OH groups
in cutin monomers.
Supplemental Figure S3. Esterification level of glycerol in cus1mutants
and Pro35S:SlCUS1RNAi cutin fruits.
Supplemental Figure S4. Esterification levels of 9(10),16-dihydroxyhexadecanoic
acid of cus1 mutants and Pro35S:SlCUS1RNAi fruit cutin.
Supplemental Figure S5. Typical Raman spectra of non-alkylated and
alkylated cutin, and alkylation reagent.
Received October 16, 2015; accepted December 7, 2015; published December 16,
2015.
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