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In Situ Nondestructive Analysis of Kalanchoe pinnata Leaf Surface Structure by Polarization-Modulation Infrared Reflection-Absorption Spectroscopy Tetsuya Hama, Akira Kouchi, Naoki Watanabe, Shinichi Enami, Takafumi Shimoaka, and Takeshi Hasegawa J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.7b09173 • Publication Date (Web): 17 Nov 2017 Downloaded from http://pubs.acs.org on November 18, 2017

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The Journal of Physical Chemistry

In Situ Nondestructive Analysis of Kalanchoe pinnata Leaf Surface Structure by Polarization-Modulation Infrared ReflectionAbsorption Spectroscopy Tetsuya Hama,*,a Akira Kouchi,a Naoki Watanabe,a Shinichi Enami,b Takafumi Shimoaka,c and Takeshi Hasegawac a

Institute of Low Temperature Science, Hokkaido University, Sapporo 060-0819, Japan

b

c

National Institute for Environmental Studies, 16-2 Onogawa, Tsukuba, Ibaraki 305-8506, Japan

Laboratory of Chemistry for Functionalized Surfaces, Division of Environmental Chemistry,

Institute for Chemical Research, Kyoto University, Gokasho, Uji, Kyoto 611-0011, Japan. *Corresponding Author: [email protected]

Telephone: +81-11-706-5474

1 Environment ACS Paragon Plus


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Abstract

The outermost surface of the leaves of land plants is covered with a lipid membrane called the cuticle that protects against various stress factors. Probing the molecular-level structure of the intact cuticle is highly desirable for understanding its multifunctional properties. We report the in situ characterization of the surface structure of Kalanchoe pinnata leaves using polarizationmodulation infrared reflection-absorption spectroscopy (PM-IRRAS). Without sample pretreatment, PM-IRRAS measures the IR spectra of the leaf cuticle of a potted K. pinnata plant. The peak position of the CH2-related modes shows that the cuticular waxes on the leaf surface are mainly crystalline, and the alkyl chains are highly packed in an all-trans zigzag conformation. The surface selection rule of PM-IRRAS revealed the average orientation of the cuticular molecules, as indicated by the positive and negative signals of the IR peaks. This unique property of PM-IRRAS revealed that the alkyl chains of the waxes and the main chains of polysaccharides are oriented almost perpendicular to the leaf surface. The nondestructive, background-free, and environmental gas-free nature of PM-IRRAS allows the structure and chemistry of the leaf cuticle to be studied directly in its native environment.


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Introduction Land plants have hydrophobic lipid membranes called cuticles at their leaf surfaces to minimize water loss and external stresses caused by air pollutant exposure, ultraviolet irradiation, and pathogens (Fig. 1A) 1–7. Recent studies indicate that the surface of plant leaves may act as an important sink for ozone and other volatile organic compounds, thereby influencing the regional and global atmospheric composition 8–11. However, the structure and chemistry of the leaf cuticle 5,11

underlying its multiple functions are not fully understood

. The lack of experimental

techniques for the direct, nondestructive characterization of the leaf cuticle is an important limiting factor. The primary chemical components of the leaf cuticle have been investigated using gas chromatography-mass spectroscopy (GC-MS) 1,12. The cuticle is composed of cutin, a polymer of hydroxylated fatty acids with chain lengths of 16 and 18 carbon atoms (C16 and C18) cross-linked by ester bonds

1–3,12

. In addition to cutin, polysaccharides play key roles in the structural

framework and physicochemical properties (e.g., water absorption capacity) of the leaf cuticle 2– 5,13–16

. Phenolic compounds are also distributed in the leaf cuticle

1–6

. The thickness of the leaf

cuticle ranges from the submicrometer scale to 10 µm or more (e.g., 200 µm) depending on the plant species

1,2,17

. An organic solvent-soluble wax covers the cuticle surface (epicuticular wax)

and is distributed across the leaf cuticle (intracuticular wax)

1–6,18

. These cuticular waxes are

typically a mixture of unbranched aliphatic (fully saturated) hydrocarbons, such as alkanes, alcohols, aldehydes, and fatty acids, with carbon chain lengths of C20–C40, and esters of these fatty acids and alcohols

1–3

. The thickness of epicuticular wax is estimated to be in the range of

50–375 nm, which corresponds to 20–90% of the total wax layer 18.



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The physicochemical properties of organic thin film materials, including lipid membranes, are determined by the primary chemical structure and the molecular arrangement in the film 19,20. However, the fine chemical structure of the leaf cuticle is not detected by conventional mass spectroscopy analysis dedicated to the identification of the primary chemical components. Alternative methods using surface-sensitive techniques based on electrons, ions, or X-rays can damage the sample because of the high-energy radiation, particularly in biological systems. In addition, they often require sample pretreatments and work best under a vacuum

21

. Therefore,

the study of the intact cuticle structure is restricted by these technical problems, and many aspects of the cuticle structure related to its chemical composition remain unclear 4,5. Here, we describe a method for in situ nondestructive IR measurements of the cuticle membrane of the leaf surface of Kalanchoe pinnata (Lam.) Pers. using polarization-modulation infrared reflection–absorption spectroscopy (PM-IRRAS) (Fig. 1B). PM-IRRAS was originally developed for high-sensitivity measurements of organic thin films on a substrate 22–28, where the absorbance is weak because the film thickness (d) is much smaller than the IR wavelength (d ≪ λ). PM-IRRAS uses a Fourier transform infrared (FT-IR) spectrometer equipped with a linear polarizer and a photoelastic modulator (PEM), which alternately (typically 50–100 kHz) generates the s- and p-polarized IR light to obtain a ratio spectrum (PM-IRRAS signal), S, of =

− + + −

1

where Rp and Rs are the reflectance of the s- and p-polarizations, respectively (Fig. 1A). J2(ϕ0) and J0(ϕ0) are the second- and zero-order Bessel functions of the maximum dephasing, and ϕ0 is introduced by the PEM. C is a constant accounting for the signal processing. Previous studies demonstrated that the surface selection rule of the ratio spectrum (equation 1) of a thin film on a dielectric (nonmetallic) substrate can be substituted qualitatively with that of external reflection


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spectroscopy (Fig. 1A)

24–29

. In the present study, the surface selection rule is described as

follows: (i) the surface-parallel component of a transition moment yields a positive peak and (ii) the surface-normal component yields a negative peak (Fig. 1C) because the angle of incidence is set to 76°, which is larger than Brewster’s angle of the air/leaf interface 24–29. PM-IRRAS is a nondestructive analytical tool because it uses a moderate continuum IR light. Moreover, it has the advantage of being a background-free measurement technique because the ratio spectrum (S) is obtained using Rp and Rs (equation 1). This is in contrast to conventional IR measurements, which require single-beam background and sample measurements to obtain a sample spectrum. Because it is intrinsically impossible to measure the background spectrum of a bare leaf surface (substrate) without the cuticle (sample) nondestructively, this characteristic is well suited for the method described in the present study. In addition, the absorption due to environmental gases (e.g., H2O and CO2) is eliminated from the ratio spectrum because its isotropic nature contributes equally to both Rp and Rs. Thus, the nondestructive, background-free, and environmental gas-free nature of PM-IRRAS allows in situ analysis of a living specimen without damage. We also show that the application of PM-IRRAS to the leaf surface has the unique advantage that the average molecular orientations in the intact cuticle can be determined from the surface selection rule.

Experimental methods Kalanchoe pinnata (Lam.) Pers. was used as a model plant, because the leaf surface of K. pinnata is sufficiently flat for PM-IRRAS. Kalanchoe pinnata was grown indoors in the Institute of Low Temperature Science, Hokkaido University, Japan. The leaves were 5–10 cm long in the direction of the main vein. n-Hexane (>96%, Junsei Chemical Co., Ltd.) and chloroform (99%;



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Wako Pure Chemical Industries, Ltd.) were used for immersion experiments. PM-IRRAS measurements were performed on an FT-IR spectrophotometer (Nicolet iS50, Thermo Fisher Scientific) at room temperature (Fig. S1). The modulation frequency of the interferometer in the FT-IR spectrophotometer was 20 kHz and the band resolution was 4 cm–1. The angle of incidence was 76°, which is an optimal angle of incidence 27. The interference IR light was led out of FT-IR and passed though the PEM with an intrinsic resonance frequency of 50 kHz to alternately generate s- and p-polarized light by a PEM controller (PEM-100, Hinds Instruments). The double-modulated (interferometer and PEM) IR light was introduced directly onto the leaf surface, and the reflected light was fed to a mercury cadmium telluride (MCT) detector. The focal size of the IR light on the leaf surface was approximately 4 mm in diameter, and it was larger along the direction of the IR light because of the large incidence angle of 76°. The PMIRRAS signal (S) was obtained using a synchronous sampling demodulator (SSD-100-15, GWC Technologies). The lowest wavenumber limit was approximately 900 cm−1, which was the cutoff wavenumber of the CaF2 lens on the MCT detector (Figs. 1B and S1). The accumulation number of the interferogram collection was 2000, which took approximately 2377 s. A spectrum with a good signal-to-noise ratio was obtained with a measurement time of >120 s (Fig. S2). The chemical composition of the cuticular waxes was identified using GC-MS with an Agilent Technologies 6890N GC/5973AMSD system. Waxes were extracted by immersion of a K. pinnata leaf in a 4:1 mixture of n-hexane and ethyl acetate (>99.8 %; Wako Pure Chemical Industries, Ltd.) at room temperature.

Results PM-IRRAS analysis of the leaf surface


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A photograph of the in situ analysis of the leaf surface of a potted K. pinnata plant is shown in Fig. 2A (see also Fig. S1). A broad range spectrum (4000–900 cm−1) was readily obtained without sample pretreatment, and positive and negative peaks appeared depending on the orientation of each functional group in the regions of 3800–2600 and 1900–900 cm−1 (Fig. 2B). The ratio spectrum baseline is a broad arch because of the Bessel function derived from the wavelength dependence of the maximum dephasing generated by the PEM modulator, which is specific to the ratio spectrum (equation 1)

22,30

. In Fig. 2B, the half-wave retardation frequency,

which provides a maximum-efficiency wavenumber for the ratio spectrum, is set to 2300 cm−1 (Fig. S3). The ratio spectra were measured with the same optical configuration using half-wave retardation frequencies of 3000 (Fig. 2C) and 1600 cm−1 (Fig. 2D) to obtain good quality spectra with a relatively flat baseline in the regions of 3800–2600 and 1900–900 cm−1, respectively (Fig. S3). In Fig. 2C, the positive peaks are assigned to the anti-symmetric stretching and symmetric CH2 stretching vibration (νa(CH2) and νs(CH2)) bands at 2914 and 2846 cm−1, respectively. A small positive peak is observed at 2951 cm−1 for the CH3 asymmetric out-of-skeleton stretching vibration (νa(CH3)os) band, where the skeleton is the molecular plane of the all-trans zigzag alkyl chain (Fig. 3A). The OH stretching vibration (ν(OH)) band appears as a broad negative peak at 3427 cm−1. The ratio spectrum in Fig. 2D has positive peaks for the C=O stretching vibration (ν(C=O)) band at 1736 and 1709 cm−1. The CH2 scissoring vibration (δ(CH2)) band appears as doublet positive peaks at 1473 and 1462 cm−1. Strong negative peaks of the ν(C–O) related bands are observed in the region of 1200–1000 cm−1. The OH2 bending vibration of water (δ(OH2)) appears as broad negative and positive bands at 1700–1550 cm−1. We also measured the ratio



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spectra of the adaxial and abaxial surfaces of a K. pinnata leaf cut from a plant, and obtained qualitatively similar spectra (Fig. S4). Representative bands used to characterize K. pinnata leaves are listed in Table 1. The positions of other typical IR bands for cutin, hemicellulosic polysaccharides, xyloglucan, and cellulose are also shown in Table 1 6,15,31–35. The peak positions of the intense negative ν(C–O) bands in the region of 1200–1000 cm−1 are close to those of hemicellulosic polysaccharides and xyloglucan, except for a small peak at 1105 cm−1 that probably arose from cutin. The characteristic peak of pectic polysaccharides around 1145 cm−1 is not observed

32–34

. The contribution of cellulose to the peaks at 1105, 1059, and 1039 cm−1 may

be minor, considering the absence of a peak at 1160 cm−1 (Fig. 2D). As shown in Table 1, the peaks at 1105, 1059, and 1039 cm−1 can also arise from cutin (1105–1101 cm−1), hemicellulosic polysaccharides (1061–1041 cm−1), and xyloglucan (1041 cm−1), respectively. The characteristic peaks of aromatic rings (phenolic compounds) and C=C double bonds around 3000 and 1600 cm−1 are not observed, even when the wavenumbers are set at the half-wave retardation frequencies

6,28,29

. The abundances of unsaturated species including cutan, an organic solvent-

insoluble and non-hydrolyzable polymer with aromatic rings and/or C=C double bonds

6,31

, are

thus below the detection limit. Phenolic compounds were also not identified by GC-MS analysis of the cuticular waxes of K. pinnata.

Solubility test for cuticular wax PM-IRRAS can be used to study the structural changes of the cuticle in response to external physical or chemical stress. In the present study, solvent invasion into the leaf surface was investigated. The ratio spectra of a K. pinnata leaf cut from a plant before solvent treatment, after immersion in n-hexane for 16 h, and after immersion in chloroform for a further 16 h are


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shown in Figs. 4A and B, a–c. n-Hexane partly extracts non-polar wax constituents, whereas chloroform extracts all the wax constituents 1. The νa(CH2) and νs(CH2) bands became weaker after immersion in n-hexane (Fig. 4A, b), and they almost disappeared after immersion in chloroform (Fig. 4A, c). This confirms that PM-IRRAS probes the organic solvent-soluble epicuticular and intracuticular waxes in the K. pinnata leaf cuticle. Considering the minor contribution of cellulose to the ratio spectra, the cell wall layer mainly functions as a dielectric substrate, and the leaf cuticle functions as a thin sample film in the three-layer system (air /thin sample/substrate) (Fig. 1A) 29,36. If the thickness of a sample film is larger or of the order of the IR wavelength (d ≳ λ), the ratio spectra should have derivativeshaped bands in the entire wavenumber range due to the specular-reflection component in the two-layer system (air/thick sample)

28,29

. This is not the case for the ratio spectra in the present

study (Figs. 2 and 4). Thus, the probing depth (d) of PM-IRRAS is much smaller than the IR wavelength (d ≪ λ)

29

. Since the value of λ ranges between 2.5–11.1 µm (4000–900 cm−1), the

present PM-IRRAS probes roughly less than 100-nm surface regions of the K. pinnata leaf cuticle as a thin sample film. The baseline of the ratio spectrum can be distorted in a region in which there is strong absorption by a dielectric substrate

24–27

. A precise determination of the

probing depth will require a more detailed characterization of the K. pinnata leaf cuticle, including the thickness measurement, by combined PM-IRRAS and other techniques. In the spectral region of 1900–1000 cm−1 (Figs. 4B and C), the δ(CH2) band became weak after immersion in n-hexane for 16 h, whereas the doublet peaks were still present. This implies that the cuticular waxes maintained a crystalline form even after immersion in n-hexane. The δ(CH2) band disappeared after further immersion in chloroform, indicating the complete removal of the cuticular waxes (Fig. 4C).



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In contrast, most of the strong negative ν(C–O) bands remained almost intact even after subsequent immersion in n-hexane and chloroform for 16 h each (Fig. 4B). This confirms that the cell wall layer is resistant to organic solvents and functions as the substrate for the PMIRRAS measurements, and that the negative ν(C–O) bands are attributed to organic solventinsoluble compounds in the K. pinnata leaf cuticle. For the ν(OH) band, a strong negative signal was detected at around 3450 cm−1 before solvent treatment (Fig. 4A, a). After immersion in nhexane and chloroform, the band changed to broad positive and negative signals in the range of 3800–3400 and 3400–2600 cm−1, respectively (Fig. 4A, c). These results indicate that both organic solvent-soluble and insoluble compounds have hydroxyl or carboxyl groups.

Discussion CH2 and CH3 vibrations: Crystalline epicuticular wax The νa(CH2), νs(CH2), and δ(CH2) bands are discussed first because they show a sensitive response to the molecular conformation. The νa(CH2) and νs(CH2) bands appear at 2914 and 2846 cm−1, respectively (Fig. 2C). These positions are typical for an alkyl chain with an ordered all-trans zigzag conformation with high molecular packing (Fig. 3A). An alkyl chain with a mobile or a disordered (gauche) conformation shows peaks at 2924 and 2855 cm−1 or higher 29. The solvent immersion experiments (Fig. 4) showed that the νa(CH2), νs(CH2), and δ(CH2) bands mainly arise from the organic solvent-soluble cuticular waxes. Our results indicate that the alkyl chains in the cuticular waxes on the K. pinnata leaf surface mostly adopt the all-trans zigzag conformation rather than the gauche conformation. The all-trans zigzag conformation is typical for solid molecules with sufficiently long alkyl chains, such as stearic acid (CH3(CH2)16COOH) 29

. In fact, GC-MS analysis of the cuticular waxes of K. pinnata showed that they are mainly


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composed of alkanes, alcohols, and fatty acids with carbon-chain lengths with a range of C14–C33, and terpenoids such as dehydroabietic acid (C19H29COOH) and β-Amyrin ((3β)-Olean-12-en-3ol, C30H50O) (Fig. S5). This result supports that most of the molecules in the cuticular waxes of K. pinnata have a sufficiently long alkyl chain for the all-trans zigzag conformation. Qualitative analysis of the orientation of the alkyl chains in the cuticular waxes is possible based on the positive or negative sign of the bands and considering the surface selection rule (Fig. 1C). Because the νa(CH2) and νs(CH2) bands are positive bands, both vibrations are nearly surface-parallel, which implies that the alkyl chains stand nearly perpendicularly to the K. pinnata leaf surface (Fig. 3A). Another notable finding is the doublet δ(CH2) band at 1473 and 1462 cm−1 (Fig. 2D). This band yields doublet peaks at these positions when the alkyl chains are involved in a crystalline structure

29,37

. This is the factor-group splitting, which is induced by two orthogonal

vibrational couplings between the contiguous CH2 groups of the two chains in the orthorhombic or monoclinic structure (Fig. 3B)

37,38

, in which highly ordered packing of molecular planes of

the all-trans zigzag skeleton is necessary. The δ(CH2) band becomes a singlet peak at around 1473–1467 cm−1 when the alkyl chains have triclinic or hexagonal structures, or an amorphous form

29,37,39,40

. The observation of the doublet δ(CH2) band is consistent with the results of the

analysis of the νa(CH2) and νs(CH2) band positions (Fig. 2C), which indicates the all-trans zigzag conformation. Considering the probing depth (less than 100 nm) and the typical thickness of epicuticular wax (50–375 nm)

18

, our results show that the cuticular waxes, especially the

epicuticular waxes, of K. pinnata have the orthorhombic and/or monoclinic structures (Fig. 3B). This is also in good agreement with a previous electron and X-ray diffraction study of isolated



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epicuticular waxes from the leaves of 35 plants, which indicated that most of the waxes have an orthorhombic structure 41. Regarding the terminal CH3-related bands, the ratio spectrum showed a small positive peak at 2951–2953 cm−1 for the νa(CH3)os band (Figs. 2C, and 4A, a), whereas the two other related modes were not observed, namely, the asymmetric in-skeleton stretching vibration (νa(CH3)is) at 2962 cm−1 and the symmetric stretching vibration (νs(CH3)) at 2870 cm−1 (Fig. 3A) 29

. This result supports the discussion above, because the only νa(CH3)os band showed a highly

parallel orientation to the surface compared with that in other modes, giving a positive peak at 2951 cm−1

29

. In addition, there is a strong negative ν(OH) signal at around 3420–3460 cm−1

before solvent treatment (Figs. 2C and 4A, a). The negative ν(OH) band is also reasonable considering an alkyl chain with the terminal hydroxyl (or carboxyl) group oriented nearly perpendicular to the leaf surface, yielding a surface-normal ν(OH) vibration (Fig. 3A). The broad negative and positive δ(OH2) bands appear at 1700–1550 cm−1 (Fig. 2D). Overlap of the positive and negative signals suggests that the H2O molecules in the K. pinnata leaf cuticle are isotropic (randomly oriented).

C-O vibrations of other cuticular chemical constituents The leaf cuticle is composed of cutin, a polymer of C16 and C18 fatty acids cross-linked by ester bonds

1–3,12

. The positive ν(C=O) bands at 1736–1709 cm−1 are close to characteristic

peaks of the ester bonds in cutin 6,15,31. However, the amount of cutin alone cannot fully account for the negative ν(C–O) bands because of the following three reasons. First, the negative ν(C–O) band intensities at 1200–1000 cm−1 are much stronger than the positive ν(C=O) signals. Because the numbers of C=O and C–O groups are identical in an ester bond (Fig. S6), the


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positive/negative PM-IRRAS signal intensities of the C=O/C–O vibrations should be the same order of magnitude as those of the ester bonds in cutin, even if the difference of the absorption coefficients is considered. In fact, a stronger ν(C=O) signal than the ν(C–O) signal was reported in the IR spectra of isolated cutin from leaves 6,14, which was attributed to the stronger permanent dipole moment of the C=O group than that of the C–O group 29. The fact that the negative ν(C– O) bands are stronger than the positive ν(C=O) bands suggests that the number of C–O bonds involved in the ratio spectra is larger than that of the C=O bonds. Second, the two strong peaks at 1126 and 1082 cm−1 do not correspond to the characteristic peaks of the ester bonds in cutin (Table 1) 6. Third, the peak position at 1173–1171 cm−1 is higher than that usually observed for cutin (1168–1161 cm−1) 6,15,31. The most plausible explanation for the origin of the ν(C–O) peaks is the presence of polysaccharides in the K. pinnata leaf cuticle as described in previous studies

2,5,14,15

.

Hemicellulosic polysaccharides have characteristic absorption bands at 1173, 1126, and 1061– 1041 cm−1

32

. Xyloglucan, a major hemicellulose of growing cell walls in dicots

characteristic absorption bands at 1078 and 1041 cm−1

33,34

42

, has strong

. These peak positions are close to

those observed in the ratio spectra (Table 1). One of the most common repeat units of xyloglucan, designated as XXXG with a main chain of β-1,4-D-glucan (G) and side chains of xylose (X) is shown in Fig. 5 42. In many dicots, the main chain has three substituted glucose units (e.g., X or galactose) followed by an unsubstituted glucose residue (G) 42. According to the surface selection rule, the strong negative peaks for the ν(C–O) bands indicate a large surface-normal component of the transition moment, namely, a strong perpendicular orientation of the C–O bonds to the leaf surface. The negative ν(C–O) bands can appear if the main chain of a polysaccharide is oriented nearly perpendicular to the leaf surface,



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and the surface-perpendicular orientation is dominant in the K. pinnata leaf cuticle (Fig. 5). Polysaccharides can also explain the presence of broad positive and negative ν(OH) bands in the range of 3800–2600 cm−1 after immersion in n-hexane and chloroform, i.e., removal of the cuticular waxes (Fig. 4A, c), because they generally have many OH bonds. Overlap of the positive and negative signals implies the random orientation of the OH bonds in polysaccharides compared with the main chain (Fig. 5). Molecular orientation of cellulose and polysaccharides in the cell wall plays a critical role in determining the mechanical properties of the cell wall 43. The perpendicular orientation of the alkyl chains of the cuticular waxes and the main chains of polysaccharides to the leaf surface suggested by the present findings in K. pinnata could provide new insights into the molecular-level mechanism of the physics and chemistry of the leaf cuticle. For example, it would be related to the mechanical properties of plant leaf surfaces, the bidirectional transport of matter (gas, liquid, and solid) between plant leaves and the surrounding environment, and the resistance against biotic and abiotic stresses 5. Polysaccharides in the leaf cuticule is still a topic of current research 5,13–16. For example, polysaccharides are usually considered to be localized at the innermost cuticle region in contact with the cell wall, whereas recent transmission electron microscopy studies reported that polysaccharides are also present in the outer cuticle region immediately under the epicuticular wax for Eucalyptus globulus, Populus×canescens, and Pyrus communis

5,13

. The present PM-

IRRAS also implies that polysaccharides are widely distributed across the K. pinnata leaf cuticle including the outer region based on (i) their association with epicuticular and intracuticular waxes, (ii) the probing depth of less than about 100 nm, and (iii) the minor contribution of cellulose (cell wall) to the ratio spectra

36

. The PM-IRRAS signal intensity of polysaccharides

(ν(C–O) bands) was stronger than that of cutin (ν(C=O) bands) (Fig. 2D). Moreover, the


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methylene groups of cutin do not contribute substantially to the ratio spectra, because the νa(CH2) and νs(CH2) bands became very weak after immersion in n-hexane and chloroform (Fig. 4A, c). There results indicate a large abundance of polysaccharides in the K. pinnata leaf cuticle compared with cutin and/or the isotropic ester and methylene groups in cutin. In the latter case, the positive and negative ν(C=O), νa(CH2), and νs(CH2) signals were mostly overlapped in the ratio spectra, resulting in small net positive signals

24,27

. Further investigation is needed to

determine the location and amount of polysaccharides in the leaf cuticle precisely and clarify the role of their molecular orientation. The background-free and nondestructive nature of PMIRRAS allows the simultaneous or subsequent use of other analytical tools, including microscopy and mass spectroscopy techniques. Combination studies may reveal the fine chemical structure of the leaf cuticle.

Conclusions The present study describes the in situ IR analysis of the intact cuticle of K. pinnata leaves using PM-IRRAS. This technique enables the selective observation of the surface wax layer and its interface with the top surface of the cell walls without background interference. PMIRRAS determines the conformation, crystallinity, and average orientation of the cuticular molecules. The positive νa(CH2), νs(CH2), and δ(CH2) bands indicated that the alkyl chains of the cuticular waxes are dominantly crystalline, highly packed in an all-trans zigzag conformation, and oriented nearly perpendicular to the leaf surface. The strong negative ν(C–O) bands arose from organic solvent-insoluble compounds, most likely polysaccharides. The negative sign suggested that the C–O bonds in the main chains are oriented perpendicular to the leaf surface.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

These findings provide helpful information for the molecular-level interpretation of the physicochemical properties of the leaf cuticle.

Supporting Information Photograph of the PM-IRRAS measurement of a leaf surface of K. pinnata, ratio spectra of a leaf surface of K. pinnata, GC-MS analysis of the cuticular waxes of K. pinnata, and schematic of an ester bond (PDF).

This information is available free of charge via the Internet at

http://pubs.acs.org Acknowledgments We thank Prof. Y. Chikaraishi, and Ms. Y. Takizawa for GC-MS analysis of the cuticular waxes of K. pinnata and fruitful discussions. We are also grateful to Profs. R. Tanaka, Y. Kimura, H. Hidaka, Y. Oba, and Drs. K. K. Tanaka, T. Yamazaki, and S. Ishizuka for helpful comments and suggestions. We thank Drs. K. Ono and R. Sato, and Mrs. R. Taniguchi for taking care of the plants. This work was supported by Japan Society for the Promotion of Science KAKENHI grants 24224012, 16H06024 (Hama), and 15H02185 (Hasegawa), Ministry of Education, Culture, Sports, Science and Technology KAKENHI grant 25108002, and the Collaborative Research Program of Institute for Chemical Research, Kyoto University (grants 2016-76 and 2017-87). Author contributions T.H., A.K., and N.W. conceived and designed the experiments. T.H. carried out the experimental measurements. T.H., S.E., T.S., and Ta.H. performed data analysis. All authors discussed the results and co-wrote the paper.


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Notes: The authors declare no competing financial interests.



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Table 1. Observed band positions in the ratio spectra of the K. pinnata leaf, and characteristic IR absorption bands of plant cutin, polysaccharides, and cellulose.a,b Peak band Group Positive/ Cutin 6,15,31 Hemicellulosic Xyloglucan Cellulose 33,34 34,35 polysaccharides positions vibrations negative 32 (cm−1) 3420–3460 (broad)

ν(OH)

Negative

3390–3304 (broad)

3440 (broad)

-

-

2952

νa(CH3)os

Positive

-

-

-

-

2915

νa(CH2)

Positive

2922–2918

-

-

-

2847

νs(CH2)

Positive

2853–2849

-

-

-

1736

ν(C=O)

Positive

1732–1728

-

-

-

1709

1707–1700

1700–1550 (broad)

δ(OH2) (water)

Positive/ Negative

-

-

-

-

1473, 1462

δ(CH2)

Positive, doublet

1463–1457

-

-

-

1172

ν(C-O)

Negative

1168–1161

1173

-

1160

1126

ν(C-O)

Negative

-

1126

1120

-

1105

ν(C-O)

Negative

1105–1101

-

-

1109

1082

ν(C-O)

Negative

-

-

1078

-

1059

ν(C-O)

Negative

-

1061–1041

-

1060–1057

1039

ν(C-O)

Negative

-

-

1041

1040–1030

a

Peak position varies ± 1 cm−1 on K. pinnata leaves.

b

The bands at 1450–1200 cm−1 are not discussed in detail because the band signals were ambiguous.


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Figure legends

Fig. 1. Schematic overview of the PM-IRRAS technique. (A, left) Schematic of the leaf cuticle in PM-IRRAS. PS: polysaccharides. (A, right) Schematics of external reflection spectroscopy. The plane formed by the incident and reflected IR light is called the incident plane. The polarizations of the radiation with electric field vectors parallel and perpendicular to the incident plane were the p- and s-polarizations, respectively. θ is the angle of incidence. (B) Schematic of the PM-IRRAS set-up on the leaf surface. FT-IR: Fourier transform infrared spectrometer; MCT: mercury cadmium telluride detector; PEM: photoelastic modulator. (C) Schematic of the surface selection rule on a dielectric material surface with a large angle of incidence (θ = 76º). The surface-parallel and perpendicular components of a transition moment yield a positive and negative peak, respectively. 25 Environment ACS Paragon Plus


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Fig. 2. PM-IRRAS measurements of K. pinnata leaves. (A) Photograph of the PM-IRRAS measurement of a leaf surface of a potted K. pinnata plant. The red arrows show the IR light path. The size of the leaf is approximately 5 cm along the direction of the main vein. Ratio spectra with half-wave retardation frequencies set to (B) 2300, (C) 3000, and (D) 1600 cm−1.


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A

νa(CH3)os positive CH3

ν s(CH3)

B b

ν a(CH3)is

H2C CH2 νa(CH2) positive

νs(CH2) positive

H

H2C

a

CH2 H2C CH2

C H H C H

n

H2C O H Leaf

ν(OH) negative

a c

b

Fig. 3. Structure of the alkyl chains in the cuticular waxes. (A) All-trans zigzag conformation of an alkyl chain with the terminal hydroxyl group almost perpendicular to the substrate. The arrows show the direction of the transition moment of the group vibrations. ⊗ shows the direction of a transition moment, which is parallel to the substrate and perpendicular to the molecular plane of the all-trans zigzag conformation. (B) Orthorhombic structure involving two chains and four methylene groups viewed along the molecular c-axis. The c-axis is parallel to the long axes of the all-trans zigzag alkyl chains. Three axes, a, b, and c, are distinct (a ≠ b ≠ c) and intersect at 90° angles. The monoclinic structure is similar to the orthorhombic structure, whereas they differ in that the alkyl chains (c-axis) are tilted with respect to the ab plane 38.



5

A

4

S

3

(a) Before immersion (b) n-Hexane (c) Chloroform (a) 3450

2

(b)

2916

2848

2953 2916 2848 2918 2850

(c) 1 0 4000 3800 3600 3400 3200 3000 2800 2600 2400 2200 2000 -1

Wavenumber (cm )

B

5

(a) 4

S

3 2

(b) (c)

1 0 1900 1800 1700 1600 1500 1400 1300 1200 1100 1000

900

-1

Wavenumber (cm )

C

4.4 4.3 4.2

1473

(a)

1462

(b) (c)

S

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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4.1 4.0 3.9 3.8 1510

1490

1470

1450

1430

-1

Wavenumber (cm )

Fig. 4. Comparison of ratio spectra of a K. pinnata leaf cut from a living plant. (a) Before solvent treatment, (b) after immersion in n-hexane (500 mL) for 16 h, and (c) after additional immersion in chloroform (150 mL) for 16 h. The half-wave retardation frequencies were set to (A) 3000 and (B) 1600 cm−1. (C) Magnification of spectra in (B) in the range of 1510–1430 cm−1.


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Negative peak

Xyl

Xyl

Xyl

Glc

Leaf Fig. 5. Structure of xyloglucan. A representative repeat unit of the XXXG oligosaccharide is shown. XXXG has a main chain of β-1,4-D-glucan (G) and side chains of xylose (X). Glc: glucose; Xyl: xylose. When the main chain is oriented perpendicular to the leaf surface, the total number of C–O bonds perpendicular to the leaf surface (eight red circles) is greater than that of bonds parallel to the leaf surface (six blue circles), even assuming parallel orientations of all side chains to the leaf surface. As a result, the transition moment over XXXG has a surface-normal component, yielding the negative ν(C–O) bands.



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TOC GRAPHICS


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Fig. 1. Schematic overview of the PM-IRRAS technique. (A, left) Schematic of the leaf cuticle in PM-IRRAS. PS: polysaccharides. (A, right) Schematics of external reflection spectroscopy. The plane formed by the incident and reflected IR light is called the incident plane. The polarizations of the radiation with electric field vectors parallel and perpendicular to the incident plane were the p- and s-polarizations, respectively. θ is the angle of incidence. (B) Schematic of the PM-IRRAS set-up on the leaf surface. FT-IR: Fourier transform infrared spectrometer; MCT: mercury cadmium telluride detector; PEM: photoelastic modulator. (C) Schematic of the surface selection rule on a dielectric material surface with a large angle of incidence (θ = 76º). The surface-parallel and perpendicular components of a transition moment yield a positive and negative peak, respectively. 300x350mm (300 x 300 DPI)

ACS Paragon Plus Environment


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Fig. 2. PM-IRRAS measurements of K. pinnata leaves. (A) Photograph of the PM-IRRAS measurement of a leaf surface of a potted K. pinnata plant. The red arrows show the IR light path. The size of the leaf is approximately 5 cm along the direction of the main vein. Ratio spectra with half-wave retardation frequencies set to (B) 2300, (C) 3000, and (D) 1600 cm−1. 190x142mm (300 x 300 DPI)


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Fig. 3. Structure of the alkyl chains in the cuticular waxes. (A) All-trans zigzag conformation of an alkyl chain with the terminal hydroxyl group almost perpendicular to the substrate. The arrows show the direction of the transition moment of the group vibrations. ⊗ shows the direction of a transition moment, which is parallel to the substrate and perpendicular to the molecular plane of the all-trans zigzag conformation. (B) Orthorhombic structure involving two chains and four methylene groups viewed along the molecular c-axis. The c-axis is parallel to the long axes of the all-trans zigzag alkyl chains. Three axes, a, b, and c, are distinct (a ≠ b ≠ c) and intersect at 90° angles. The monoclinic structure is similar to the orthorhombic structure, whereas they differ in that the alkyl chains (c-axis) are tilted with respect to the ab plane 34. 400x266mm (300 x 300 DPI)



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Fig. 4. Comparison of ratio spectra of a K. pinnata leaf cut from a living plant. (a) Before solvent treatment, (b) after immersion in n-hexane (500 mL) for 16 h, and (c) after additional immersion in chloroform (150 mL) for 16 h. The half-wave retardation frequencies were set to (A) 3000 and (B) 1600 cm−1. (C) Magnification of spectra in (B) in the range of 1510–1430 cm−1. 254x338mm (300 x 300 DPI)


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Fig. 5. Structure of xyloglucan. A representative repeat unit of the XXXG oligosaccharide is shown. XXXG has a main chain of β-1,4-D-glucan (G) and side chains of xylose (X). Glc: glucose; Xyl: xylose. When the main chain is oriented perpendicular to the leaf surface, the total number of C–O bonds perpendicular to the leaf surface (eight red circles) is greater than that of bonds parallel to the leaf surface (six blue circles), even assuming parallel orientations of all side chains to the leaf surface. As a result, the transition moment over XXXG has a surface-normal component, yielding the negative ν(C–O) bands. 266x350mm (300 x 300 DPI)



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Table of contents 209x148mm (300 x 300 DPI)


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In Situ Nondestructive Analysis of

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