Detailed characterization of Pinus ponderosa sporopollenin by infrared spectroscopy

Phytochemistry 170 (2020) 112195

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Detailed characterization of Pinus ponderosa sporopollenin by infrared spectroscopy

T

Alec Lutzkea, Kevin J. Moreya, June I. Medfordb, Matt J. Kippera,∗ a b

Department of Chemical & Biological Engineering, Colorado State University, Fort Collins, CO, 80521, USA Department of Biology, Colorado State University, Fort Collins, CO, 80523, USA

A B S T R A C T

In plant spores and pollen, sporopollenin occurs as a structural polymer with remarkable resistance to chemical degradation. This recalcitrant polymer is well-suited to analysis by non-destructive infrared spectroscopy. However, existing infrared characterization of sporopollenin has been limited in scope and occasionally contradictory. This study provides a comprehensive structural analysis of sporopollenin in the Pinus ponderosa pollen exine using infrared spectroscopy, including detailed band assignments, descriptions of chemical reactivity, and comparison to multiple reference substances. We observe that the infrared spectral characteristics of sporopollenin prepared by enzymatic digestion of the polysaccharide-based intine are largely consistent with a copolymer of aliphatic lipids and trans-4-hydroxycinnamic acid, without distinct contributions from α-pyrone or carotenoid substructures.

1. Introduction Certain biopolymers exhibit a remarkable degree of preservation in the fossil record that results from resistance to typical modes of biodegradation (de Leeuw et al., 2006; Hemsley et al., 1993). Sporopollenin is a prominent example of this form of recalcitrant biopolymer and occurs in plant pollen and spores as the major constituent of the exospore or exine (Brooks and Shaw, 1978; Wallace et al., 2011). While there is considerable interest in the unique structural characteristics that confer durability to sporopollenin, its chemical composition remains uncertain. Non-destructive analytical methods such as Fourier transform infrared (FTIR) spectroscopy are ideal characterization techniques for recalcitrant biopolymers, and have been used for pollen characterization (Depciuch et al., 2017, 2016; Jardine et al., 2015). In particular, spectroscopic methods been used to identify distinct chemical environments present in sporopollenin (Ahlers et al., 2000; Espelie et al., 1989; Guilford et al., 1988; Shaw and Yeadon, 1966; Wilmesmeier et al., 1993). Despite the modern prevalence of FTIR as an analytical method, rigorous characterization of sporopollenin with this technique is relatively uncommon and there has been a lack of refinement or expansion upon prior FTIR studies of sporopollenin structure. In fact, assignments of characteristic vibrational bands in sporopollenin FTIR spectra have been subject to considerable variation. In the FTIR spectrum of sporopollenin derived from Typha angustifolia, Bubert et al. (2002) assign a band at 1625 cm−1 to an δ(NH) mode, while Schulze-Osthoff and Wiermann (1987) generically attribute this feature to aromatics, and Wilmesmeier et al. (1993)

∗

assign it to ν(C]O) or ν(C]C) modes. The method of purification can also affect which functional groups are observed. Bands appearing between 1200 and 1000 cm−1 in the spectrum of various purified sporopollenin samples were assigned to ν(CO) modes by Kawase and Takahashi (1995), Wilmesmeier et al. (1993) and Domínguez et al. (1999, 1998), while Jardine et al. (2017) attribute a band in the same region to aromatic ν(CH) modes in untreated spores. Jardine et al. (2017) also report a distinct Amide I band at 1650 cm−1 in Lycopodium annotinum spores, yet Domínguez et al. (1999) tentatively assign a band at that position to a ν(C]C) mode in Pinus pinaster, Betula alba, Ambrosia elatior, and Capsicum annuum sporopollenin. These likely represent different functional groups, as the band identified by Jardine et al. is susceptible to acetolysis, and may be from protein, whereas the band identified by Domínguez et al. resists acid treatment. Many distinct vibrational bands remain unassigned or are assigned without clear rationalization, and this form of uncertainty has hindered the development of precise spectroscopic descriptions of sporopollenin. Renewed effort to accurately assign characteristic vibrational bands in sporopollenin may provide greater insight into the structure of this recalcitrant polymer as it exists in nature. This form of spectral interpretation is facilitated by complementary studies that have identified possible monomeric constituents, functional groups, or precursors formed during biosynthesis. For example, degradation studies provide persuasive evidence that phenylpropanoids such as trans-4-hydroxycinnamic (p-coumaric) acid occur in sporopollenin from Pinus mugo (Schulze Osthoff and Wiermann, 1987; Wehling et al., 1989), Corylus avellana (Herminghaus et al., 1988), as well as moss, horsetail, fern and

Corresponding author. E-mail address: [email protected] (M.J. Kipper).

https://doi.org/10.1016/j.phytochem.2019.112195 Received 29 July 2019; Received in revised form 11 October 2019; Accepted 31 October 2019 0031-9422/ © 2019 Elsevier Ltd. All rights reserved.

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cycad species (Nierop et al., 2019). Genes involved in exine development encode proteins that may produce sporopollenin precursors by activation and oxidation of lipid substrates. The cytochrome P450 enzymes CYP703A2 (Morant et al., 2007) and CYP704B1/CYP704B2 (Dobritsa et al., 2009; Li et al., 2010) catalyze in-chain and ω-hydroxylation of fatty acid substrates, respectively. The acyl-coenzyme A synthetase ACOS5 preferentially activates hydroxylated fatty acids (de Azevedo Souza et al., 2009), while expression of the putative fatty acylCoA reductase MS2 in tobacco or bacteria induces production of fatty alcohols (Aarts et al., 1997; Chen et al., 2011; Doan et al., 2009). Multiple degradation studies have revealed that fatty acid-like lipids are key constituents of sporopollenin and may occur in nature as phenylpropanoid conjugates. Following ozonolysis of L. clavatum and P. silvestris spore walls, Shaw and Yeadon identified 7-hydroxyhexadecanedioic acid as a significant degradation product, while Li et al. later determined that a similar C16 ester of trans-4-hydroxycinnamic acid is bound to the sporopollenin macrostructure through cyclic acetal linkages (Li et al., 2019) to poly(vinyl alcohol) (PVA)-like chains that may be terminated with α-pyrone rings. Sporopollenin degradation products and the function of exine-related lipid oxidation enzymes can be aligned with reasonable consistency (Fig. 1), although it is clear that certain biosynthesis steps have yet to be identified. Further studies of sporopollenin biosynthesis show that Arabidopsis polyketide synthases encoded by PKSA (LAP6) and PKSB (LAP5) genes form α-pyrones from various tri- and tetraketide substrates produced by ACOS5 (Dobritsa et al., 2010; Kim, 2015; Kim et al., 2010; Mizuuchi et al., 2008), while TKPR1 (DRL1) and TKPR2 (CCRL6) oxidoreductases catalyze reduction of adjacent carbonyl groups following tri- or tetraketide cyclization (Grienenberger et al., 2010) (Fig. 2a). The modern description of sporopollenin as a cutin- or suberin-like copolymer of lipids and phenylpropanoids contrasts with earlier studies that proposed a terpenoid structure that is perhaps derived from polymerization of carotenoids (Brooks and Shaw, 1968; Zetzsche et al., 1937; Zetzsche and Huggler, 1928; Zetzsche and Vicari, 1931) (Fig. 2b). In principle, IR spectroscopy may be used to compare hypothesized constituents of sporopollenin to the polymer itself to improve identification of vibrational bands. In our work, sporopollenin from Pinus ponderosa pollen was isolated by solvent extraction of soluble organic compounds in combination with removal of the intine by enzymatic digestion (1% w/v Cellulase and Macerozyme R-10) (see Materials and Methods in Supplementary Data). For comparison, we also investigated alternative isolation methods that have been commonly reported in the literature (i.e., acidolysis with phosphoric acid or acetolysis) to discern their effects on

Fig. 2. Examples of (a) α-pyrone biosynthesis in the exine and (b) carotenoids that may be oxidatively polymerized to form sporopollenin according to the hypothesis of Brooks and Shaw (1968).

the spectral characteristics of sporopollenin. The purified sporopollenin samples were analyzed by attenuated total reflectance FTIR (ATR-FTIR) and compared with various model compounds intended to represent known or hypothesized structural components of the exine. These putative small-molecule constituents include phenylpropanoids (trans-4hydroxycinnamic and trans-4-hydroxy-3-methoxycinnamic acid), carotenoids (β-carotene), fatty acids (oleic and stearic acid), fatty alcohols (1-hexadecanol), α-pyrones (4-hydroxy-6-methyl-2-pyrone), and others (Fig. 3). Spectral analysis was supplemented by several chemical treatments aimed at degrading or derivatizing particular functional groups in enzymatically-isolated sporopollenin as summarized in Table 1. The resulting spectroscopic data was used to refine the assignment of IR bands and to further elucidate structural features of this Fig. 1. Scheme depicting enzymes involved in sporopollenin synthesis and selected degradation products. (a) Mid-chain oxidation of fatty acid substrates by CYP703A2, ω-oxidation by CYP704B1 and CYP704B2, and reduction of an activated fatty acid substrate by MS2. (b) Proposed sporopollenin monomer identified by Li et al. (2019). (c) The sporopollenin degradation products trans-4-hydroxycinnamic acid, trans-4-hydroxy-3-methoxycinnamic acid, and 7-hydroxyhexadecanedioic acid.

2

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Fig. 3. Model compounds used in the infrared characterization of sporopollenin.

Macerozyme R-10 were used to digest the pectocellulosic intine and a sequence of solvent extractions were performed to remove soluble organic compounds (Fig. S1). This mild approach results in apparent removal of intine polysaccharides without significantly affecting the spectroscopic characteristics of sporopollenin itself or the morphology of the exine (Fig. S2a,b). Elemental analysis by X-ray photoelectron spectroscopy (XPS) reveals that exine purified in this manner consists of carbon, nitrogen, and oxygen in proportions that are consistent with those observed in commercial Lycopodium sporopollenin (Table 2). We observed that the proportion of nitrogen in the exine initially increases following enzymatic digestion, and incubation in deionized water for 72–144 h at 40 °C is necessary to remove this contamination prior to further study. This result aligns with an earlier hypothesis that sporopollenin becomes contaminated with enzyme residue during enzymatic digestion of the intine (Ahlers et al., 2000). The nitrogen content of the pollen wall (prior to possible contamination by exogenous enzymes) decreases following treatment with protease isolated from Aspergillus oryzae, supporting the observation of Huang et al. (2013) that lipid transfer proteins may be co-localized with sporopollenin in the exine. For comparative purposes, sporopollenin was also obtained by heating solvent-extracted P. ponderosa pollen in 85% w/w phosphoric acid at 50 °C for 72 h (Guilford et al., 1988; Wilmesmeier et al., 1993; Zetzsche and Vicari, 1931) and through acetolysis over 15 min with 90% v/v acetic anhydride and 10% v/v concentrated sulfuric acid (Erdtman, 1960) (Fig. S3 and S4). Both methods are frequently reported to selectively remove intine polysaccharides without degradation of sporopollenin itself, although our observations suggest that this is not the case. Instead, treatment with phosphoric acid and acetolysis produce an exine that is spectroscopically distinct from the natural sporopollenin present in the pollen wall, despite retention of gross morphological similarity (Fig. S2c,d). The nitrogen content of the exine is

Table 1 Chemical treatments of enzymatically-isolated exine and summarized effects. Treatment

Effect

1 M KOH, 40 °C, 72 h D2O, 40 °C, 72 h Methanol-d4, 40 °C, 72 h Acetone, 1% w/v I2 Octanal, 1% w/v I2 Peracetic acid, 3% w/w, 20 °C, 72 h Peracetic acid, 32% w/w, 20 °C, 72 h Dilute Br2 in chloroform, 20 °C, 72 h NaBH3CN in THF, 20 °C, 72 h

none none Partial OH → OD exchange of alcohols Alcohol loss, acetonide formation Alcohol loss, octanal acetal formation none Bleaching of exine, loss of aromatic constituents, acetylation Bleaching of exine, loss of aromatic constituents none

recalcitrant biopolymer. 2. Results and discussion 2.1. Isolation of sporopollenin In our work, commercial pollen from P. ponderosa was used as a model system to study the isolation and spectroscopic characteristics of the sporopollenin-rich exine. We compare our isolated exines to commercially available sporopollenin from Lycopodium, used as received from the supplier. We selected Pinus pollen because of both its commercial availability and the comparatively large number of prior studies conducted using pollen from various Pinus species (Li et al., 2019; Schulze Osthoff and Wiermann, 1987; Shaw and Yeadon, 1966, 1964; Wehling et al., 1989). Exine that was spectroscopically unaltered from its condition in the natural pollen wall was prepared by adaptation of the Ahlers et al. (1999) protocol, in which commercial Cellulase and 3

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Table 2 XPS elemental composition of pollen and exine. Sample

%C

Solvent-extracted P. ponderosa pollen After Aspergillus oryzae protease Enzymatically-isolated exine After 72 h in water at 40 °C After 144 h in water at 40 °C After 72 h in 1 M KOH at 40 °C Exine after acidolysis with phosphoric acid Exine after acetolysis Commercial Lycopodium sporopollenin

61.4 65.6 67.3 67.4 67.5 73.0 75.4 68.0 68.5

%N ± ± ± ± ± ± ± ± ±

0.8 0.7 0.8 1.3 1.0 0.8 0.8 0.8 0.5

4.2 2.3 6.4 4.5 4.2 1.1 1.3 – 2.5

± ± ± ± ± ± ±

%O 0.7 0.2 0.8 0.2 0.5 0.1 0.1

± 0.1

34.5 32.1 26.3 28.1 28.4 25.9 23.3 30.1 29.0

%S ± ± ± ± ± ± ± ± ±

0.4 0.6 0.2 1.1 0.6 0.7 0.9 0.8 0.4

– – – – – – – 1.9 ± 0.3 –

additional band at approximately 3060 cm−1 is potentially present, but emerges only weakly from the ν(OH) absorbance and cannot be conclusively identified. Both bands are likely associated with trans-4-hydroxycinnamyl moieties, and various chemical treatments (vide infra) reduce or eliminate these features in unison with other absorbances assigned to that substructure. The more prominent of these two bands remains detectable at 3027 cm−1 when sporopollenin is isolated by treatment with phosphoric acid and at 3020 cm−1 following acetolysis. In the spectrum of enzymatically-isolated sporopollenin, the intense absorptions at 2926 and 2855 cm−1 arise from νasym(CH2) and νsym(CH2) modes of methylene groups and are not accompanied by discernible methyl νasym(CH3) (~2950 cm−1) or νsym(CH3) bands (~2860 cm−1) (Coates, 2000). While methyl νasym(CH3) and νsym(CH3) bands are fully resolved in spectra of n-octane (Fig. S8) and stearic acid (Fig. S9), these features merge with methylene ν(CH2) absorbances in oleic acid and glyceryl trioleate, and similar overlap in the spectrum of sporopollenin cannot be excluded. For this reason, the lack of detectable methyl ν(CH3) bands does not reliably indicate the absence of methyl groups. The methylene νasym(CH2) and νsym(CH2) bands remain present after P. ponderosa pollen is treated with phosphoric acid (2926 and 2855 cm−1) or subjected to acetolysis (2928 and 2857 cm−1). In the latter case, evidence elsewhere in the spectrum indicates that acetyl groups are present, yet methyl νasym(CH3) and νsym(CH3) bands are not

significantly reduced by treatment with phosphoric acid and is eliminated altogether by acetolysis. However, sporopollenin samples isolated with phosphoric acid or acetolysis exhibit a derivative macromolecular structure with a non-native chemical composition, and are therefore largely unsuitable for IR study of the natural biopolymer. For these reasons, we believe that only the enzymatically-isolated exine is consistent with P. ponderosa sporopollenin as it exists in nature.

2.2. Carbon-hydrogen vibrational bands The hypothesized lipid and phenylpropanoid constituents of sporopollenin can be distinguished in IR spectral regions characteristic of methyl, methylene, alkenyl, and aromatic carbon-hydrogen vibrational modes. ATR-FTIR spectra (4000-650 cm−1) corresponding to each isolation method are depicted in Fig. 4 and assignments are provided in Table 3. The carbon-hydrogen vibrational bands present in the ATRFTIR spectrum of native P. ponderosa pollen are uniformly retained after enzymatic digestion of the intine. At approximately 3026 cm−1, an alkene or aromatic ν(CH) band overlaps with the broader ν(OH) absorbance. This absorbance occurs at a significantly higher frequency than equivalent features in oleic acid (3004 cm−1; Fig. S5) or glyceryl trioleate (3004 cm−1; Fig. S6) and is positionally consistent with the band at 3025 cm−1 in trans-4-hydroxycinnamic acid (Fig. S7). An

Fig. 4. ATR-FTIR spectra of (i) P. ponderosa pollen, (ii) enzymatically-isolated sporopollenin, (iii) sporopollenin isolated by acidolysis with phosphoric acid, and (iv) sporopollenin isolated by acetolysis. Original data files are deposited with the accompanying Data in Brief article (Lutzke et al., 2019). 4

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Table 3 ATR-FTIR band assignments. Band position (cm−1)a P. Ponderosa pollen

Enz.-isolated sporopollenin

H3PO4 acidolyzed sporopollenin

Acetolyzed sporopollenin

Major assignmentc

3400–2400 3325 3011b 2919 2850 2745b 2689b 2623b

3400–2400 3350 3026b 2926 2855 2741b 2689b 2621b

3400–2400 3384 3026b 2926 2855 2736b 2689b 2626b 1775b [A′]

3400–2400 3394

Carboxylic acid ν(OH) Alcohol ν(OH) Alkenyl or aromatic ν(CH) νasym(CH2) νsym(CH2) Carboxylic acid overtone or combination bands

2928 2857 2646b 1775b [A′]

1743b 1740d 1707b 1678

1710b [A] 1680 [B]

1702 1680b

1630

1630 [C]

1605 1585b 1550b 1515

1605 [D] 1588 [E] 1556b [F] 1515 [G]

1632 1614 [D′] 1607 1590b

1464b 1439 1412b

1463b [H] 1437 [I] 1411b [J]

1378 1347b 1324 1309b 1283 1262 1233

1379 [K] 1348b [L] 1323 [M] 1308b [N] 1283b [O] 1261 [P] 1233b [Q]

1380 1342 1324b 1307b 1273b

1203 1168

1202 [R] 1168 [S] 1125b [T] 1103 [U]

1204 1170 1129 1107 1074

1735b 1729 [Ac-1] 1706 1645 [C′]

1103 1072 1034

1605

1516 1493b [G′] 1462b 1437 1404 [J′]

1515 1488b [G′] 1463b 1431 1399b 1371 [Ac-2]

1236 1233 [Ac-3] 1196 1170 1125

1021 [V″] 1013b 989 940b 917b 833 719 a b c d

985 [V] 943b [W] 912b [X] 833 [Y] 774 720 [Z]

Lactone ν(C]O)d Lipid esters, ν(C]O) Ester ν(C]O) Acetyl ν(C]O) Carboxylic acid ν(C]O) α,β-Unsaturated ester ν(C]O) Alkenyl ν(C]C)d Alkenyl ν(C]C)d Aromatic ring modes

1006 [V′] 937b

946 907 834

833 777 722

719

Protein, Amide II Aromatic ring mode Unassigned δscissors(CH2) Carboxylic acid δ(OH), δscissors(CH2) δscissors(CH2) Acetyl δsym(CH3) δsym(CH3) γwagging(CH2) or methine deformation Alcohol δ(OH), overlapping Methine deformation Carboxylic acid ν(CO) Ester ν(CO) Ester ν(CO) Acetyl ν(CO) Aromatic δ(CH) Aromatic δ(CH), ν(CO) Ether or alcohol ν(CO) Ether or alcohol ν(CO) Polysaccharides Polysaccharides Ether or alcohol ν(CO) Polysaccharides Alkenyl γ(CH) Carboxylic acid γ(OH) Unassigned Aromatic γ(CH) Unassigned δrocking(CH2)n, n ≥ 4

Reported as the mean ± standard deviation of three replicate experiments. Shoulder or overlapping band; position estimated from the second derivative spectrum. Anticipated major vibrational contributions from stretching (ν), in-plane bending (δ), or out-of-plane bending (γ). Low confidence band or assignment.

detected. This outcome supports the hypothesis that methyl ν(CH3) bands are readily obscured by overlap with equivalent methylene absorbances. If a weak aldehyde ν(CH) band (~2850 cm−1) were present in this region, it would be concealed by the prominent methylene νsym(CH2) feature (Saier et al., 1962a). In contrast, the Fermi resonance band of the aldehyde doublet is potentially present as a subtle feature at 2741 cm−1, although this absorbance is ambiguous and more plausibly arises from an unrelated overtone or combination band (Saier et al., 1962b). In eight aliphatic model compounds, the methylene δscissors(CH2) mode manifests as a doublet with mean positions of 1468 ± 3 and 1459 ± 2 cm−1 and may overlap with the methyl δasym(CH3) band (Snyder, 1961) (Table S1). The ATR-FTIR spectrum of enzymaticallyisolated sporopollenin exhibits a positionally similar shoulder at 1463 cm−1 that we attribute to the δscissors(CH2) vibration of lipid-like

alkyl chains. This region is shown in greater detail in Fig. 5. This feature also appears in the spectra of sporopollenin obtained from acidolysis (1462 cm−1) and acetolysis (1463 cm−1) with remarkable positional consistency. Regardless of the isolation method, this shoulder is not a principal feature in the spectrum of sporopollenin and the dominant band in this region occurs at lower frequency. In the case of sporopollenin prepared via enzymatic digestion or acidolysis of the intine, the major band appears at 1437 cm−1, while acetolysis shifts the absorbance maximum to 1431 cm−1. This band has been assigned elsewhere to δ(CH2) or δ(CH3) vibrations (Wilmesmeier et al., 1993) and its continued assignment to a methylene δscissors(CH2) mode is feasible, although the maximum is bathochromically displaced to lower frequency than anticipated for simple hydrocarbons. This displacement could indicate the presence of nearby electron withdrawing groups (EWG) such as alcohols (Zerbi et al., 1987), as demonstrated by the 5

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Fig. 5. ATR-FTIR spectra of (i) enzymatically-isolated sporopollenin, (ii) sporopollenin isolated by acidolysis with phosphoric acid, and (iii) sporopollenin isolated by acetolysis. Distinct bands or shoulders with diagnostic importance are assigned a unique identifier in Table 3. Original data files are deposited with the accompanying Data in Brief article (Lutzke et al., 2019).

appearance of the δscissors(CH2) band at 1430 cm−1 in poly(vinyl alcohol) (PVA; Fig. S10) (Krimm et al., 1956a). The band may be alternatively assigned to the δ(OH) mode of carboxylic acids, which also manifests in the spectra of oleic (1432 cm−1) and stearic (1430 cm−1) acids (Pudney et al., 2009). In any event, the band is positionally inconsistent with the major δscissors(CH2) mode of saturated fatty acids or other uninterrupted alkyl chains. Adhering to a δscissors(CH2) assignment obligates a sporopollenin structural model with highly substituted aliphatic regions that are not predicted by single in-chain and ω-hydroxylation of fatty acid substrates, as effected by CYP703A2 (Morant et al., 2007) and CYP704B1/CYP704B2 (Dobritsa et al., 2009; Li et al., 2010). Interestingly, a highly substituted architecture may align with the repeating 1,3-diol motif proposed by Li et al., 2019, which is attributed to polyketide synthase-mediated elongation of fatty acids. A greater EWG-related positional displacement occurs when the methylene group is adjacent to carbonyls, as in the case of α-methylene δscissors(CH2) bands in the spectra of octanal (1410 cm−1; Fig. S11), stearic acid (1411 cm−1), and oleic acid (1412 cm−1) (Holland and Nielsen, 1962). This effect is likely responsible for the shoulder at 1411 cm−1 in the spectrum of enzymatically-isolated sporopollenin, which is apparently unrelated to aromatic constituents (vide infra). In comparison, the corresponding vibration in the ester glyceryl trioleate appears at 1417 cm−1. When sporopollenin is isolated by treatment with phosphoric acid, the band at 1411 cm−1 in the spectrum of enzymatically-isolated sporopollenin is apparently shifted to 1404 cm−1, and a further displacement to 1399 cm−1 occurs following acetolysis (Fig. 5). In the case of acidolyzed sporopollenin, this shift can be largely reversed by exposure to aqueous base (1 M KOH, 40 °C, 72 h), which results in the appearance of a shoulder at 1409 cm−1. This phenomenon conceivably arises from chemical alteration of a reactive functional group that influences the frequency of the α-methylene δscissors(CH2) mode. In the spectrum of enzymatically-isolated sporopollenin, the minor band at 1379 cm−1 is readily assigned to the characteristic methyl δsym(CH3) mode. This vibrational mode is sensitive to local structure and appreciable shifts in band position are induced by electronegative substituents (Sheppard, 1955). In the spectrum of n-octane, the δsym(CH3) band is found at 1378 cm−1, while the methoxy δsym(CH3)

band of trans-4-hydroxy-3-methoxycinnamic acid appears at 1411 cm−1 (Fig. S12), and the spectrum of β-carotene includes three distinct methyl δsym(CH3) bands at 1394, 1381, and 1367 cm−1 (Fig. S13) (Berezin and Nechaev, 2005). Open-chain carotenoids (e.g., lycopene) and other polyterpenes like natural rubber may not exhibit three distinct δsym(CH3) maxima due to the absence of the β-ionone ring, and the fact that this series is undetected does not necessarily support the lack of polyterpenoid structures (Binder, 1963). However, the position and isolation of the presumptive δsym(CH3) band in sporopollenin suggest that putative methyl groups do not occur in the form of methoxy ethers or as part of a β-carotene-like structure. Furthermore, the band at approximately 1379 cm−1 endures after sporopollenin is subjected to chemical degradation and is almost certainly derived from monomeric constituents of the resilient, largely aliphatic fraction of the biopolymer. This band is detected at 1380 cm−1 in the spectrum of sporopollenin prepared by treatment with phosphoric acid, but is concealed by a more prominent band at 1371 cm−1 following acetolysis (Fig. 5). In the latter case, this independent methyl δsym(CH3) band arises from formation of acetate esters by reaction with acetic anhydride. Exposure to aqueous alkali (1 M KOH, 40 °C, 72 h) hydrolyzes these acetate esters and the band at 1371 cm−1 is consequently reduced in intensity. It is possible that methyl groups occur as unoxidized termini of fatty acids, yet the roles of the cytochrome P450 enzymes CYP704B1 and CYP704B2 in sporopollenin biosynthesis are apparently catalysis of ωhydroxylation (Dobritsa et al., 2009; Li et al., 2010). If fatty acids are the source of the 1379 cm−1 band, CYP704B2-catalyzed ω-hydroxylation is clearly not an obligatory step that must precede incorporation into the sporopollenin macromolecule. Moreover, known C16 fatty acid-derived monomers obtained from sporopollenin (e.g., 7-hydroxyhexadecanedioic acid) clearly possess oxidized termini and cannot exhibit a δsym(CH3) vibrational mode (Li et al., 2019; Shaw and Yeadon, 1966). The possibility that the absorbance at 1379 cm−1 is unrelated to methyl deformation must also be considered, particularly given the absence of resolved methyl νasym(CH3) or νsym(CH3) bands. For example, PVA exhibits a weak band near 1380 cm−1 that is not necessarily derived from residual acetate esters and may arise from a γwagging(CH2) phenomenon (Krimm et al., 1956a; Snyder and 6

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frequency γ(CH) mode of vinyl alkenes, although numerous other functional groups may absorb in this region. In the spectrum of enzymatically-isolated sporopollenin, the major component of the prominent band at 833 cm−1 is presumably a γ(CH) mode of the 1,4-disubstituted aromatic ring of trans-4-hydroxycinnamyl units. This assignment is supported by the lack of other prominent absorbances in this region that would indicate alternate substitution patterns (Socrates, 2001). Similar bands appear in the spectrum of trans-4-hydroxycinnamic acid itself (829 cm−1; Fig. S7) and structurally analogous trans-4-methoxycinnamic acid (822 cm−1; Fig. S16). In the case of methyl trans-4-hydroxycinnamate (Fig. S17), a doublet appears in this region with maxima at 832 and 822 cm−1, while the spectrum of trans-4-hydroxy-3-methoxycinnamic acid exhibits separated bands at 851 and 803 cm−1 (Kalinowska et al., 2014). It is therefore unlikely that trans-4-hydroxy-3-methoxycinnamic acid is a significant constituent of P. ponderosa sporopollenin, since the single, largely symmetrical aromatic γ(CH) band at 833 cm−1 is more consistent with 1,4disubstituted trans-cinnamic acid derivatives. The ATR-FTIR spectrum of 4-hydroxy-6-methyl-2-pyrone includes two significant bands at 833 and 812 cm−1 (Fig. S18), however the adjacent ν(CO) band at 878 cm−1 is not distinctly present in the spectrum of sporopollenin (Seixas de Melo et al., 2001). The comparative distribution of aromatic γ(CH) bands for various model compounds is depicted in Fig. 6. The aromatic γ(CH) band at 833 cm−1 is diminished by isolation with phosphoric acid or acetolysis. In the latter case, the region becomes complex and contains a broad absorbance with several apparent maxima.

Schachtschneider, 1963). In the spectrum of enzymatically-isolated sporopollenin, shoulders at 1348 and 1308 cm−1 adjoin a maximum at 1323 cm−1, and all three features are superimposed upon a broader δ(OH) absorbance that manifests near the center of this series (Fig. 5). Given the presence of a linear aliphatic domain in sporopollenin, it may be practical to attribute these features to methylene γwagging(CH2) modes (Krimm et al., 1956b). It may also be feasible to assign the band at 1348 cm−1 to the methine deformation of secondary alcohols, ethers, or acetals, since this typically weak vibrational mode may be enhanced by the adjacent EWG (Coward, 2010). The spectra of 5-nonanol (Fig. S14) and cellulose (Fig. S15) both exhibit a similar band at 1334 cm−1. By extension, bands at 1323 and 1308 cm−1 may also arise from methine deformation. A series of three absorbances (1342, 1324, and 1307 cm−1) also appears in the spectrum of sporopollenin prepared via acidolysis, however the local absorbance maximum has shifted from 1323 to 1342 cm−1 (Fig. 5). This change in relative intensity presumably reflects a displacement of the underlying alcohol δ(OH) mode, mirroring the behavior of the corresponding ν(OH) band (vide infra). In the case of acetolyzed sporopollenin, bands in this region are likely concealed by the major acetyl δsym(CH3) absorbance at 1371 cm−1 (Fig. 5). The spectrum of enzymatically-isolated sporopollenin exhibits sharp bands at 1202 and 1168 cm−1 that are reasonably consistent with aromatic δ(CH) modes and can be compared with bands at 1214 and 1172 cm−1 in trans-4-hydroxycinnamic acid (Fig. S7). Isolation with phosphoric acid causes these bands to appear as weaker features at 1204 and 1170 cm−1 (Fig. 5). This reduction in intensity would appear to support the hypothesis that the bands originate from aromatic δ(CH) modes of trans-4-hydroxycinnamyl moieties, since all spectral features assigned in that manner are uniformly diminished after exposure to phosphoric acid. Following acetolysis, several sharp bands at 1196 and 1170 cm−1 are superimposed on the prominent ester ν(CO) absorbance at 1233 cm−1. The former band is eliminated as a resolved absorbance maximum by alkaline hydrolysis, while the band at 1170 cm−1 is revealed as a major spectral feature (Fig. 5). We hypothesize that the band at 1170 cm−1 in the spectrum of acetolyzed sporopollenin is a ν(CO) mode that is distinct from the sharper aromatic δ(CH) band observed after milder isolation methods. The IR spectra of sporopollenin samples from diverse plant species exhibit a characteristic absorbance at approximately 1000 cm−1, which has been assigned to an ether ν(CO) mode or erroneously to an aromatic ν(CH) mode (Jardine et al., 2017; Wilmesmeier et al., 1993). This feature appears as a relatively broad band with an absorbance maximum of 985 cm−1 in the spectrum of enzymatically-isolated P. ponderosa sporopollenin, and is unquestionably produced from overlap of multiple vibrational bands. This absorbance is likely to include a contribution from the trans-1,2-disubstituted alkene γ(CH) mode characteristic of cinnamic acid derivatives such as trans-4-hydroxycinnamic acid (977 cm−1; Fig. S7), trans-4-hydroxy-3-methoxycinnamic acid (972 cm−1; Fig. S12), trans-4-methoxycinnamic acid (975 cm−1; Fig. S16), and methyl trans-4-hydroxycinnamate (984 cm−1; Fig S17). The γ(CH) mode of isolated cis alkenes (e.g., oleic acid; Fig. S5) is found at substantially lower frequency and is not identified in the spectrum of enzymatically-isolated sporopollenin (Kobayashi and Kaneko, 1986; Sinclair et al., 1952). Isolation of the exine with phosphoric acid shifts the maximum in this region from 985 to 1006 cm−1, which can be rationalized as diminished contribution from the trans-1,2-disubstituted alkene γ(CH) mode of trans-4-hydroxycinnamyl constituents. Following acetolysis, there are no major bands that can be assigned to the trans1,2-disubstituted alkene γ(CH) mode of trans-4-hydroxycinnamyl moieties, although a minor shoulder is present at 971 cm−1 that is consistent with this deformation (Fig. 5). The low intensity of this putative γ(CH) band is mirrored by general loss of spectral features associated with aromatic substituents as a consequence of acetolysis. The spectrum of acetolyzed sporopollenin exhibits a band at 907 cm−1 that is not fully eliminated by aqueous alkali and may correspond to the lower

Fig. 6. ATR-FTIR spectra of (i) enzymatically-isolated sporopollenin, (ii) trans4-hydroxycinnamic acid, (iii) trans-4-hydroxy-3-methoxycinnamic acid, (iv) trans-4-methoxycinnamic acid, and (v) methyl trans-4-hydroxycinnamate. Original data files are deposited with the accompanying Data in Brief article (Lutzke et al., 2019). 7

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Fig. 7. Scheme depicting the synthesis of acetonides (cyclic ketals) from 1,2- or 1,3-diols naturally present in sporopollenin, followed by hydrolysis under acidic conditions. This process corresponds to the spectroscopic changes observed in Figure S21.

A minor band appears at 720 cm−1 in the spectrum of enzymatically-isolated sporopollenin, and an equivalent feature is present after treatment with phosphoric acid (722 cm−1) or acetolysis (719 cm−1) (Fig. 5). This band is assigned to the δrocking(CH2) mode of aliphatic chains containing multiple, contiguous methylene groups. In long-chain aliphatic solids, the δrocking(CH2) mode is partitioned into two bands that are clearly observed in the ATR-FTIR spectra of 1-hexadecanol (729 and 719 cm−1; Fig. S19) and stearic acid (728 and 719 cm−1; Fig. S9). This splitting results from vibrational interaction of methylene chains in close proximity, leading to resolution of distinct inphase and out-of-phase rocking modes (Li et al., 2004; Stein and Sutherland, 1954). In polyethylene, splitting of the δrocking(CH2) mode is also evident and arises from packing within the ordered lamellae of the crystalline phase (Krimm et al., 1956b). This splitting phenomenon is not observed in the ATR-FTIR spectra of liquids such as oleic acid and n-octane, where a single band at 722 cm−1 appears (Figs. S5 and S8). The absence of a δrocking(CH2) mode doublet in enzymatically-isolated sporopollenin and the location of the band maximum at 720 cm−1 indicate that close packing of methylene segments is unlikely. In general, analysis of carbon-hydrogen vibrational bands in P. ponderosa sporopollenin reveals a structure that consists of both aliphatic (i.e., methylene rich) and aromatic components.

diminished and displaced to 3394 cm−1 following acetolysis (Fig. 4). These shifts are consistent with structural alteration or reorganization that results in weaker alcohol hydrogen bonding (Novak, 1974). In the case of acetolyzed sporopollenin, the effect arises in part from conversion of alcohols to acetate esters. Exposure to aqueous alkali has no significant effect on the alcohol ν(OH) absorbance maximum in the spectrum of enzymatically-isolated sporopollenin (3352 cm−1), but appears to reduce the frequency of this band in the case of phosphoric acid-treated (3370 cm−1) or acetolyzed exine (3363 cm−1). The latter shift is likely to reflect hydrolysis of acetate esters, restoration of native alcohols, and concomitant alteration of hydrogen bonding within the structure of sporopollenin. Sporopollenin-like biopolymers such as cutin and suberin are hypothesized to contain 1,2-diols derived from oxidation of unsaturated fatty acids (Graça, 2015). Moreover, it has been recently proposed that the structure of sporopollenin itself may include a repeating 1,3-diol architecture (Li et al., 2019). The formation of cyclic acetals or ketals from 1,2- or 1,3-diols is diagnostic for the presence of those alcohol configurations (Fig. 7). Suspension of enzymatically-isolated sporopollenin in acetone containing 0.1% w/v iodine as an acetalization catalyst (Basu et al., 2002) at 50 °C for 72 h resulted in spectral changes that are consistent with acetone ketal (acetonide) formation (Fig. S21, ii). The alcohol ν(OH) band is largely removed and occurs as a weak absorbance from 3600 to 3000 cm−1, indicating that the majority of alcohol groups have been consumed by condensation with acetone. This process predictably results in the appearance of a methyl νasym(CH3) band at 2989 cm−1 and greater intensity of the δsym(CH3) band at 1379 cm−1. Two bands develop at 941 and 874 cm−1 that can be assigned to acetonide ν(CO) or ring deformation modes, indicating that the reaction is not arrested at the hemiketal stage (Barker et al., 1959). In contrast, heating in acetone without iodine (50 °C, 72 h) has no effect on the ATR-FTIR spectrum of sporopollenin. Ketals are sensitive to acidcatalyzed hydrolysis, and we observe that exposure of the sporopollenin-acetone ketal to hydrochloric acid (6 M, 40 °C, 72 h) eliminates IR bands associated with acetonides and generally restores the original spectrum of sporopollenin, as depicted in Fig. S21, iii. Moreover, iodine is also able to catalyze acetalization of sporopollenin with the aliphatic aldehyde octanal, resulting in the incorporation of aliphatic heptyl chains. We conclude that alcohols in sporopollenin are generally arranged in a 1,2- or 1,3-diol configuration, as depicted in Fig. 7. Interestingly, this substitution pattern is not predicted by known elements of the sporopollenin biosynthetic pathway. Regardless of isolation method, the alcohol ν(OH) band overlaps with a broader feature extending to 2400 cm−1 that is recognizable as the ν(OH) mode of carboxylic acids. This distinct absorbance appears in the spectra of oleic and stearic acids and exhibits a similar range of approximately 3400-2400 cm−1 (Figs S5 and S9). In the spectrum of enzymatically-isolated sporopollenin, a sequence of subtle features with estimated positions of 2741, 2689, and 2621 cm−1 are superimposed upon the broader carboxylic acid ν(OH) region. This pattern is also

2.3. Oxygen-hydrogen vibrational bands Alcohols and carboxylic acids exhibit both ν(OH) and δ(OH) oxygen-hydrogen vibrational modes. These functional groups are readily susceptible to hydrogen-deuterium exchange using deuterium oxide (D2O) or methanol-d4 as a source of deuterium, resulting in distinctive IR band shifts. In the spectrum of enzymatically-isolated sporopollenin, the broad band centered at 3350 cm−1 is unambiguously assigned to the ν(OH) vibrational mode of alcohols participating in hydrogen bonding. Curiously, this band is unaffected by hydrogendeuterium exchange in deuterium oxide at 40 °C for 72 h, indicating that the alcohol groups of sporopollenin do not participate in hydrogendeuterium exchange equilibria under our experimental conditions (Fig. S20, ii). The resistance of P. ponderosa sporopollenin to this method of deuteration implies that its protic functional groups are inaccessible to water, which may result from hydrophobic inhibition of diffusion. This hypothesis is congruent with our observation that dilute aqueous base is unable to deprotonate carboxylic acids in enzymatically-isolated sporopollenin (if present), and may also account for the well-characterized resistance to aqueous mineral acids displayed by the polymer. In contrast, use of methanol-d4 as the deuterium source (40 °C, 72 h) produces an appreciable degree of isotopic exchange. This exchange reduces the relative intensity of the alcohol ν(OH) absorbance centered at 3376 cm−1 and results in the development of an equivalent ν(OD) band at 2502 cm−1 (Fig. S20, iii). The ν(OH) absorbance maximum at 3350 cm−1 is shifted to 3384 cm−1 when isolated with phosphoric acid, and the band is both 8

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entirely sequestered from the aqueous medium or are not present. This form of sequestration is not necessarily preserved in acetolyzed sporopollenin, and exposure to aqueous alkali hydrolyzes acetate esters and shifts the speculative δ(OH) band from 1431 to 1436 cm−1, accompanied by a loss of intensity and peak resolution (Lutzke et al., 2019). However, the relative magnitude of the intensity change is masked by shifts in neighboring bands that render this evidence disputable. It consequently remains uncertain if the major band in this region is derived from δ(OH) or δscissors(CH2) vibrational modes, or perhaps overlap of both. In the spectrum of enzymatically-isolated sporopollenin, the δ(OH) mode of aliphatic alcohols is centered at approximately 1320 cm−1 (Fig. 5). A similar feature is observed at 1324 cm−1 in the spectrum of PVA (Krimm et al., 1956a) (Fig. S10). Although three absorbances appear in this region at 1348, 1323, and 1308 cm−1, these cannot be conclusively assigned to distinct δ(OH) modes. As discussed previously, bands in this region may be alternatively interpreted as carbon-hydrogen vibrational modes that are superimposed upon the underlying δ(OH) absorbance. The δ(OH) band shifts to a position near 1340 cm−1 when sporopollenin is isolated by treatment with phosphoric acid, and acetolysis results in the appearance of a strong acetyl νsym(CH3) absorbance at 1371 cm−1 that obscures this region. The broad feature at 985 cm−1 largely obscures two shoulders with estimated positions of 943 and 912 cm−1 in the spectrum of enzymatically-isolated sporopollenin (Fig. 5). The higher frequency shoulder is attributable to the carboxylic acid γ(OH) deformation, which also appears in the spectra of stearic (941 cm−1) and oleic acids (935 cm−1) and is predictably absent in glyceryl trioleate (Holland and Nielsen, 1962) (Figs. S5, S6 and S9). The second feature at 912 cm−1 is ambiguous and is not immediately characteristic of any particular functional group. Following treatment with phosphoric acid, only the putative γ(OH) deformation remains detectable at 937 cm−1. This feature is poorly resolved and has perhaps shifted to higher frequency or diminished. Acetolysis appears to result in significant enhancement of these spectral features, and two well-resolved bands manifest at 946 and 907 cm−1. The band at 946 cm−1 is reduced to a shoulder by aqueous alkali, supporting assignment of this feature to the carboxylic acid δ(OH) mode. In contrast, the band at 907 cm−1 remains detectable at diminished intensity and is conceivably unrelated to oxygen-hydrogen vibrational modes of carboxylic acids.

Fig. 8. ATR-FTIR spectra of (i) enzymatically-isolated sporopollenin, (ii) trans4-hydroxycinnamic acid, (iii) trans-4-hydroxy-3-methoxycinnamic acid, (iv) trans-4-methoxycinnamic acid, and (v) methyl trans-4-hydroxycinnamate. Original data files are deposited with the accompanying Data in Brief article (Lutzke et al., 2019).

present after isolation with phosphoric acid, but only an absorbance at approximately 2646 cm−1 is clearly detectable following acetolysis (Fig. 4). Additional bands at lower frequency may also be present, but do not clearly emerge from the substantial background absorbance. It has been theorized that bands near 2700 cm−1 in carboxylic acids arise from the combination of various fundamental frequencies that produce a characteristic pattern of maxima (Bratoz et al., 1956). The approximation of sporopollenin maxima at 2689 and 2621 cm−1 as Fermi resonance-enhanced overtones or combination bands derived from carboxylic acid absorbances between 1400 and 1300 cm−1 is clearly arithmetically feasible, and few other assignments are reasonable. Similar bands appear in the ATR-FTIR spectra of oleic acid at 2675 and 2557 cm−1, while equivalent features in stearic acid are present at 2685 and 2565 cm−1. The model compound 4-hydroxy-6-methyl-2-pyrone also exhibits absorbances in this region (2703, 2618, 2538 cm−1; Fig. S18), yet the spectrum of sporopollenin appears to lack other characteristic bands of the α-pyrone ring, and there is no compelling reason to accept this alternative explanation. As previously hypothesized, the δ(OH) mode of carboxylic acids potentially contributes to the absorbance at 1437 cm−1 in the spectra of sporopollenin isolated via enzymatic digestion or treatment with phosphoric acid, and to the equivalent band at 1431 cm−1 in the spectrum of acetolyzed sporopollenin (Fig. 5). Curiously, both enzymatically-isolated and phosphoric acid-treated sporopollenin are remarkably resistant to mild alkaline hydrolysis (1 M KOH, 40 °C, 72 h) and there is no evidence that deprotonation results in loss of carboxylic acid ν(OH) or δ(OH) bands. This lack of potassium carboxylate formation leads to the conclusion that putative carboxylic acids are

2.4. Carbon-carbon vibrational bands Sporopollenin is believed to possess alkenyl and aromatic moieties (e.g., trans-4-hydroxycinnamyl and α-pyrone) with characteristic carbon-carbon vibrational modes, permitting direct spectral comparison with putative constituents. Furthermore, targeted oxidation of alkenyl or aromatic moieties simplifies the sporopollenin spectrum through elimination of ν(C]C) bands while largely preserving the structure of saturated aliphatic hydrocarbons. The ATR-FTIR spectrum of enzymatically-isolated sporopollenin exhibits a series of adjacent bands at 1630, 1605, 1588, and 1515 cm−1 that correspond directly to counterparts in the spectrum of trans-4-hydroxycinnamic acid, where these bands appear at 1627, 1601, 1590, and 1512 cm−1 (Fig. 8). Based on previously reported assignments for trans-4-hydroxycinnamic acid, the band at 1630 cm−1 is the alkene ν(C]C) vibration, while bands at 1605, 1588, and 1515 cm−1 arise from aromatic ring modes (Swislocka et al., 2012). This pattern is appreciably altered by further substitution of the aromatic ring in trans-4-hydroxy-3-methoxycinnamic acid (1619, 1598, 1590, 1512 cm−1), and conversion to the methoxy ether (1633, 1622, 1596, 1577, 1512 cm−1) or methyl ester (1655, 1598, 1583, 1515 cm−1). These patterns are compared in Fig. 8, where similarity between the spectra of enzymatically-isolated sporopollenin, trans-4hydroxycinnamic acid, and other aromatic model compounds is apparent. Certain sporopollenin carbon-carbon bands are similar to those present in the spectrum of 4-hydroxy-6-methyl-2-pyrone, which 9

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of lignin (Ma et al., 2016). Since sporopollenin is susceptible to oxidative processes and contains lignin-like phenylpropanoid monomers, we hypothesized that it may be feasible to observe the outcome of peracetic acid-mediated oxidation by ATR-FTIR. We find that sporopollenin is resistant to dilute aqueous peracetic acid (~3% w/w, 20 °C) and exhibits few spectral changes after a 72 h exposure (Fig. S24, ii). When sporopollenin is exposed to concentrated peracetic acid (32% w/ w, 20 °C) over 72 h, bleaching is complete and the exine largely dissolves to yield a small quantity of colorless, granular residue. Examination of this residue by ATR-FTIR reveals the near absence of bands associated with trans-4-hydroxycinnamyl moieties, which remain detectable only as faint shoulders with the exception of the aromatic ring mode observed at 1511 cm−1 (Fig. S24, iii). Many of the most prominent features in the spectrum can be assigned to the ν(C]O) (1725 cm−1), δasym(CH3) (1434 cm−1), δsym(CH3) (1374 cm−1), ν(CO) (1241 cm−1), and ν(CC) (844 cm−1) modes of acetate esters (Nolin and Jones, 1956), and these assignments are supported by the presence of a methyl νasym(CH3) shoulder at 2944 cm−1. These spectral features are accompanied by methylene νasym(CH2) and νsym(CH2) bands at 2923 and 2858 cm−1, respectively, and shoulders or weak bands associated with δscissors(CH2) (1464 and 1412 cm−1) and δrocking(CH2) (721 cm−1). In general, the vibrational spectrum is consistent with a fatty acid-like, polyhydroxylated residue that no longer contains aromatic components. This residue is partially O-acetylated, presumably by acid-catalyzed esterification with acetic acid. It is therefore reasonable to hypothesize that oxidation by peracetic acid proceeds through preferential degradation of aromatic constituents in a manner analogous to its effect on lignin, while the lipid components of the exine are retained as oxidized and esterified derivatives.

displays absorbances at 1655, 1618, 1584, 1577, 1537, 1510, 1492 cm−1. If this sequence of bands is largely undisturbed by homologation of the 6-methyl group to extended alkyl chains (as in the case of hypothetical precursors produced by PKSA/PKSB) (Kim et al., 2010), then there is little reason to conclude that absorbances in this region are derived from α-pyrone vibrational modes. There is also no evidence that the structure of sporopollenin exhibits unsaturation that is distinct from the conjugated alkene in trans-4-hydroxycinnamyl units. The carbon-carbon vibrational bands are sensitive to treatment with phosphoric acid, which reduces the relative intensity of the alkene ν(C]C) band at 1632 cm−1 and aromatic ring modes at 1607 and 1516 cm−1 (Fig. 5). This outcome is further indication that acidolysis results in hydrolytic removal of trans-4-hydroxycinnamyl moieties. The aromatic ring mode found at 1589 cm−1 in the spectrum of solventextracted P. ponderosa pollen is not resolved following exposure to phosphoric acid, although a weak shoulder remains present at 1590 cm−1. The absence of the band at 1589 cm−1 coincides with the development of an absorbance at 1614 cm−1 with equal intensity to the adjacent band at 1607 cm−1. It is possible that this pattern results from significant hypsochromic shifts of bands at 1605 and 1589 cm−1 in the spectrum of solvent-extracted P. ponderosa pollen. As depicted in Fig. 8 (ii-v), the presumptive aromatic ring modes of various model compounds exhibit appreciable positional variation. It is therefore unnecessary to presume that spectral changes in this region correspond to loss of the core cinnamyl structure, although it seems probable that a degree of peripheral chemical alteration has occurred. Interestingly, aqueous potassium hydroxide reduces the band at 1614 cm−1 to a minor shoulder, and there is a corresponding increase in the intensity of the shoulder at 1585 cm−1 (Fig. S33). This may indicate that a reversible, acid- and base-sensitive process modulates the development and relative intensity of both bands. Treatment with phosphoric acid also results in the appearance of a minor band at 1497 cm−1 that is not found in P. ponderosa pollen or sporopollenin isolated by other methods. This band is positionally consistent with an aromatic ring mode, but is removed by exposure to aqueous potassium hydroxide and the source is therefore uncertain. Interestingly, partial deuteration of enzymatically-isolated sporopollenin with methanol-d4 causes the band at 1588 cm−1 to significantly decrease in intensity (Fig. S22). This behavior is mirrored in trans-4-hydroxycinnamic acid, where exposure to methanol-d4 causes the band at 1590 cm−1 to diminish and perhaps shift to 1579 cm−1 (Fig. S23). This outcome further supports assignment of bands in this region to trans-4-hydroxycinnamyl moieties. The most prominent aromatic ring modes are found in the spectrum of solvent-extracted P. ponderosa pollen at 1605 and 1515 cm−1, and remain detectable following acetolysis at identical positions with diminished intensity (Fig. 5). In contrast, the alkene ν(C]C) band at 1630 cm−1 is no longer clearly resolved, nor is the nearby ring mode at 1589 cm−1. Evidently, acetolysis exerts a deleterious effect on the structure of natural sporopollenin that results in depletion of trans-4hydroxycinnamyl substituents. It is unclear if the broad feature at 1645 cm−1 is related to the band observed at 1630 cm−1 prior to acetolysis, although assignment to the ν(C]C) mode of alkenes is sensible in either scenario. If the appearance of the band at 1645 cm−1 represents new unsaturation in sporopollenin, the probable mechanism is acid-catalyzed dehydration of alcohols. Based on our analysis, we propose that trans-4-hydroxycinnamyl units are the most abundant aromatic constituent of P. ponderosa sporopollenin; other hypothesized aromatic constituents (e.g., α-pyrones) are not uniquely detected. Furthermore, there is no spectral evidence to support the existence of polyene structural motifs derived from unsaturated fatty acids or carotenoids.

2.4.2. Effects of bromine oxidation on carbon-carbon vibrational bands The addition of bromine to sporopollenin was previously investigated by Zetzsche and Huggler (1928), who generally limited their analysis to determination of elemental composition. We observe that dramatic compositional changes are induced in enzymatically-isolated sporopollenin by exposure to dilute bromine in chloroform (20 °C, 72 h). In the ATR-FTIR spectrum of brominated sporopollenin, the alcohol ν(OH) band is appreciably diminished and the maximum is shifted from 3350 to 3411 cm−1, while an intense carbonyl ν(C]O) feature develops at 1725 cm−1, replacing the original maximum at 1680 cm−1 (Fig. S25, ii). This effect is more dramatic when bromine oxidation occurs under aqueous conditions (pH 5.0 sodium acetate buffer; Fig. S25, iii). Most significantly, bands attributed to alkene and aromatic ν(C]C) modes are absent and reveal features at 1631 and 1561 cm−1 that are assigned as Amide I and II of residual protein. This result suggests the elimination of trans-4-hydroxycinnamyl moieties, which Li et al. hypothesize to be bound by ester linkage to a C16 lipid substructure, which is in turn bound to the greater sporopollenin macrostructure by cyclic acetals (Li et al., 2019). SEM confirms that this loss of aromatic constituents occurs without dramatic morphological alteration of the exine (Fig. S26). We observe by 1H NMR that the model compound trans-4-methoxycinnamic acid reacts with a 5-fold excess of bromine in chloroform to yield the anticipated 1,2-dibrominated derivative as the major product, without cleavage of the methyl ether (Figs. S27, S28). Methyl trans-4-hydroxycinnamate gives a major product consistent with 1,2-dibromination of the alkene and electrophilic bromination of the aromatic ring ortho to the hydroxyl group, while the methyl ester is preserved (Fig. S29). It is therefore unlikely that the chemistry of bromine permits direct scission of trans-4hydroxycinnamic acid alkyl esters or aryl ethers. We therefore hypothesize that trans-4-hydroxycinnamic acid is bound by ester linkage to a substructural unit of unknown configuration, which is in turn connected to the sporopollenin macrostructure by a bromine-sensitive functional group. In the presence of water, bromine-promoted oxidation of acetals to esters (Palou, 1994; Williams et al., 1988) and the scission of alkyl

2.4.1. Effects of peracetic acid oxidation on carbon-carbon vibrational bands The oxidative scope of peracetic acid is broad and its ability to oxidize aromatic compounds has led to its use in the depolymerization 10

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absent from the structure of natural sporopollenin, or more plausibly represent minor constituents that are not easily detected. Paradoxically, the ester content of the exine is potentially enriched by isolation with phosphoric acid, which results in the appearance of a distinct shoulder at 1775 cm−1 that is removed by subsequent alkaline hydrolysis (Lutzke et al., 2019). This feature appears at an identical position in the ATR-FTIR spectrum of acetolyzed exines and may originate from the action of mineral acid on the P. ponderosa pollen wall. If this shoulder is assigned to the reasonably characteristic ν(C]O) mode of five-membered cyclic esters (~1775 cm−1) (Hall and Zbinden, 1958) it is possible to speculate that their formation results from the acid-catalyzed cyclization of γ-hydroxy acids to form the corresponding γ-lactone or an equivalent process that produces strained cyclic carbonyl structures. Aldehydes also exhibit a ν(C]O) band (1725 cm−1 in the spectrum of octanal), but the aldehyde ν(CH) (~2850 cm−1) band and additional features derived from Fermi resonance are likely to overlap with other absorbances in the spectrum of sporopollenin and cannot be specifically identified or excluded (Saier et al., 1962b). In general, there is no clear spectroscopic evidence for or against the presence of aldehydes and ketones. However, suspension of sporopollenin in a solution of 1% w/v sodium cyanoborohydride in tetrahydrofuran (20 °C, 72 h) does not result in significant spectral changes (Fig. S32) (Lane, 1975). We are consequently obliged to conclude that aldehydes and ketones are not major constituents of sporopollenin or are not effectively reduced by sodium cyanoborohydride. An abundance of α-pyrone rings is likely excluded by the absence of the weak, yet characteristic ν(C]O) band occurring at 1754 cm−1 in the spectrum of 4-hydroxy-6-methyl-2-pyrone (Seixas de Melo et al., 2001) (Fig. S18). In P. ponderosa pollen, a weak shoulder appears at 1550 cm−1 that cannot be removed with organic solvent and is sensibly identified as an Amide II band, which includes contributions from δ(NH) and ν(CN) vibrational modes (Barth, 2007). This result contradicts the optimistic perspective of Hesse and Waha (1989) that protein is destroyed by organic solvent and consequently removed from the exine. Although an Amide I band arising primarily from the ν(C]O) mode of the amide carbonyl might be anticipated to appear near 1650 cm−1, an independent maximum is not distinguishable from nearby bands derived from trans-4-hydroxycinnamyl units (Barth, 2007). The Amide II band is retained following isolation of sporopollenin via enzymatic digestion, but cannot be resolved after acidolysis with phosphoric acid or acetolysis. Exposure of enzymaticallyisolated sporopollenin to 1 M KOH for 72 h at 40 °C removes the Amide II band in a manner consistent with amide hydrolysis (Fig. S33). The notion that the exine is nitrogen-free is seemingly a consequence of prior studies that utilized harsh isolation methods, and does not necessarily reflect the composition of material obtained by enzymatic digestion. This nitrogen functional groups observed in the exine are consistent with protein, however, they are likely not from sporopollenin. Both Amide I (if genuinely present) and Amide II are likely to induce positional variability in unrelated bands that occur in this region due to overlap, but are unlikely to be confused with oxygen-carbon vibrational modes of sporopollenin itself. In the spectrum of enzymatically-isolated sporopollenin, overlapping absorbances at 1283, 1261, and 1233 cm−1 can be assigned to ν(CO) modes of carboxylic acids, esters, or alcohols (Fig. 5). Two of these bands (1283 and 1233 cm−1) manifest as shoulders on the major band at 1261 cm−1. In the spectrum of trans-4-hydroxycinnamic acid, the carboxylic acid ν(CO) mode appears at 1283 cm−1 (Clavijo et al., 2009; Nolasco et al., 2009; Swislocka et al., 2012). Similar ν(CO) bands are present in the spectra of oleic (1284 cm−1) and stearic acids (1297 cm−1) (Pudney et al., 2009) (Figs. S5 and S9). In comparison, the ν(CO) bands of glyceryl trioleate appear at 1238 (minor) and 1160 cm−1 (major) (Fig. S6). Isolation of sporopollenin by treatment with phosphoric acid eliminates the distinct maximum at 1261 cm−1, and the major band in this region now appears at 1236 cm−1 with a subtle shoulder that replaces the original 1261 cm−1 band. This change

ethers to yield carboxylic acids or ketones (Deno and Potter, 1967) have been demonstrated. However, exposure of a putative sporopolleninoctanal cyclic acetal to dilute bromine in chloroform removes trans-4hydroxycinnamyl moieties without elimination of the octanal-derived lipid substructure (Fig. S30). Therefore, acetal connectivity of the type proposed by Li et al. cannot be invoked to rationalize the ability of bromine to eliminate trans-4-hydroxycinnamyl moieties, and it is necessary to speculate that an alternative bromine-sensitive linkage is interposed between the phenolic constituents of sporopollenin and the recalcitrant macrostructure. 2.5. Oxygen-carbon and nitrogen-carbon vibrational bands Sporopollenin is hypothesized to include multiple carbon-oxygen functional groups, including alcohols, ethers, acetals, and carboxylic acids. Moreover, the purified exine may contain natural lipids, polysaccharides, or proteins that exhibit carbon-oxygen or nitrogen-carbon vibrational modes that are independent from those of sporopollenin. Unprocessed P. ponderosa pollen exhibits an IR band at 1737 cm−1 that is diminished to a subtle shoulder or removed altogether following extraction with organic solvents. This band is most plausibly assigned to the ν(C]O) mode of soluble lipid esters such as triglycerides, which are known to occur in the pollen of Pinus species (Scott and Strohl, 1962). An equivalent feature is observed at 1744 cm−1 in the ATR-FTIR spectrum of commercial glyceryl trioleate (Fig. S6). Although fatty acids are inarguably present in P. ponderosa pollen, the hydrogenbonded ν(C]O) mode of such compounds manifests at lower frequency and is not distinguishable from other absorbance features in that region. The band at 1737 cm−1 may be alternatively attributed to the ν(C]O) mode of esterified D-galacturonic acid units in pectin (appearing at 1738 cm−1 in citrus pectin; Fig. S31), yet such an assignment is inconsistent with substantial reduction of this band following extraction with organic solvent. After removal of soluble lipid esters and enzymatic digestion of the intine, the major band at 1680 cm−1 is retained and can be assigned to the ν(C]O) mode of the α,β-unsaturated carbonyl of trans-4-hydroxycinnamyl units (Schulze Osthoff and Wiermann, 1987; Wehling et al., 1989). Similar bands are present in the ATR-FTIR spectra of trans-4-hydroxycinnamic acid (1668 cm−1), trans4-methoxycinnamic acid (1672 cm−1), and methyl trans-4-hydroxycinnamate (1683 cm−1) (Figs. S7, S16, and S17). The major ν(C]O) band at 1680 cm−1 is distorted in the direction of higher frequency by a shoulder with an estimated position of 1710 cm−1, and this feature is most reasonably assigned to the ν(C]O) mode of carboxylic acids similar to oleic (1708 cm−1) or stearic acids (1699 cm−1) (Figs. S5 and S9). This assignment is supported by the presence of other spectral features associated with carboxylic acids, which must therefore be present in sufficient abundance to produce a distinct ν(C]O) band in the carbonyl region. The relationship between these two bands is reversed if P. ponderosa pollen is treated with phosphoric acid, and the prominent carboxylic acid ν(C]O) band at 1702 cm−1 is accompanied by a diminished shoulder at 1681 cm−1 (Fig. 5). This shift in relative intensity can be interpreted as cleavage of hydrolytically-sensitive trans-4-hydroxycinnamyl esters from the sporopollenin macrostructure. In comparison, the spectrum of the exine after purification via acetolysis is dominated by a single feature produced from overlap of the ν(C]O) bands of acetate esters (1729 cm−1) and carboxylic acids (1706 cm−1). The α,β-unsaturated ν(C]O) mode is no longer resolved, and it is evident that hydrolytic removal of trans-4-hydroxycinnamyl units occurs rapidly during acetolysis. In the absence of conjugation or hydrogen bonding, the ester ν(C] O) mode typically manifests at higher frequencies than carboxylic acids (1744 cm−1 in the case of glyceryl trioleate) and no distinct bands appear in this region in the ATR-FTIR spectrum of the enzymaticallyisolated exine. A weak shoulder potentially appears near 1740 cm−1, although detection of this feature is unreliable and its presence cannot be confirmed. It must therefore be concluded that saturated esters are 11

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is rationalized as partial loss of a ν(CO) or δ(OH) mode at 1261 cm−1, accompanied by an increase in the relative intensity of the adjacent 1236 cm−1 band. The decrease in the band at 1261 cm−1 after exposure to phosphoric acid is mirrored by the reduced intensity of aromatic bands elsewhere in the spectrum, permitting tentative assignment to the ester ν(CO) mode of trans-4-hydroxycinnamyl units. This region is also significantly altered by acetolysis, which results in the appearance of the acetate ester ν(CO) band at 1233 cm−1, although this absorbance may include contributions from the ν(CO) mode of hypothesized cyclic esters. This ester band is predictably eliminated by hydrolysis with aqueous alkali, while the adjacent feature at 1170 cm−1 is preserved. The intensity of the absorbance at 1170 cm−1 renders it infeasible to attribute this band to trans-4-hydroxycinnamyl units, and the frequency is unusually high when compared with the ν(CO) modes of simple alcohols (Zeiss and Tsutsui, 1953). The frequency of ν(CO) bands in simple aliphatic alcohols is dependent upon local structure, and some degree of discrimination is feasible between primary and secondary examples (Zeiss and Tsutsui, 1953). In the spectrum of enzymatically isolated sporopollenin, the band at 1103 cm−1 is consistent with the ν(CO) mode of secondary alcohols, and similar features are found in the spectra of PVA (1096 cm−1), cellulose (1100 cm−1), and pectin (1100 cm−1) (Figs. S10, S15, and S31). The shoulder with an estimated position of 1125 cm−1 is not attributable to trans-4-hydroxycinnamyl units and is assigned to the ν(CO) mode of secondary alcohols or the ν(COC) mode of aliphatic ethers. These bands are found in the spectrum of sporopollenin obtained via acidolysis at 1129 and 1107 cm−1, while only the band at 1125 cm−1 is resolved following acetolysis. No clear absorbance maxima appear between 1100 and 1000 cm−1 following enzymatic digestion of intine polysaccharides, in contrast with the appearance of bands at 1072, 1034, and 1013 cm−1 in native P. ponderosa pollen. Although these pollen absorbances overlap with other absorbances and are subject to considerable positional uncertainty, it is likely that various ν(CO) and ring breathing modes of polysaccharides such as cellulose, hemicellulose, and pectin are partial contributors (Wiercigroch et al., 2017). This is supported by the continued presence of 1081 and 1023 cm−1 bands following extraction of the pollen with organic solvents, and detection of similar bands in cellulose (1054 and 1030 cm−1) and citrus pectin (1075 and 1009 cm−1) (Figs S15 and S31). A band at 1074 cm−1 is retained following isolation of sporopollenin with phosphoric acid, which may indicate the continued presence of polysaccharides. In the case of enzymatically-isolated exines, it is probable that various ν(CO) modes also manifest as components of the major alkenyl γ(CH) absorbance at 985 cm−1, although these occur without distinct maxima. This absorbance shifts to 1006 cm−1 in the spectrum of phosphoric acid-treated exines, and further to 1021 cm−1 after acetolysis (Fig. 5). This positional fluctuation is a consequence of decreasing contribution from the trans-1,2disubstituted alkene γ(CH) band of trans-4-hydroxycinnamyl moieties that gradually reveals the subsumed oxygen-carbon vibrational mode, rather than a genuine frequency shift (Ryu et al., 2010). Altogether, the oxygen-carbon vibrational bands in sporopollenin can be largely rationalized as the combination of those native to trans-4-hydroxycinnamyl and oxidized fatty acid-like lipid monomers. Some fraction of these monomers may be linked as α,β-unsaturated esters, but spectral evidence does not support the presence of abundant saturated esters.

(98%), trans-4-hydroxy-3-methoxycinnamic acid (99%), 4-hydroxy-6methyl-2-pyrone (98%), iodine (> 99.99%), trans-4-methoxycinnamic acid (98%), microcrystalline cellulose, 5-nonanol (98%), octanal (98%), and oleic acid (99%) were purchased from Alfa Aesar (Ward Hill, MA, USA). Peracetic acid solution (32% w/w in dilute acetic acid) and protease from Aspergillus oryzae were obtained from Sigma-Aldrich (St. Louis, MO, USA). Methyl trans-4-hydroxycinnamate (> 98.0%) and stearic acid (> 98.0%) were obtained from TCI America (Portland, OR, USA). Cellulase and Macerozyme R-10 were obtained from Yakult Pharmaceutical Industry Co, Ltd. (Tokyo, Japan). Sporopollenin (Lycopodium) was purchased from Polysciences, Inc. (Warrington, PA, USA). Pollen from Pinus ponderosa was purchased from the Canadian Pine Pollen Company (North Vancouver, BC, Canada). The pollen was harvested during May and June of 2017 in Merritt, BC, Canada, and was sifted through 200 mesh and dehydrated by solar dryer prior to use. Deionized water (18.2 MΩ cm) was supplied by a Milli-Q Synthesis A10 water purification system. Preparation of sporopollenin materials is described in detail in Supporting Data. 4. Methods Infrared spectroscopy. Attenuated total reflectance Fourier transform infrared spectroscopy (ATR-FTIR) was performed using a Nicolet iS50 FTIR spectrometer equipped with a Smart iTR ATR sampling accessory and a diamond crystal plate (Thermo Fisher Scientific, Madison, WI, USA). The instrument was operated with a DTGS/KBr detector and KBr beamsplitter (XT-KBr). Spectra were generated from 64 scans with a data spacing of 0.964 cm−1 over the range of 4000 to 650 cm−1. Spectra were processed (atmospheric suppression; ATR correction was not applied) and band positions were obtained using the OMNIC software suite. In cases of band overlap, the positions of shoulders and other indistinct features were estimated from second derivative spectra generated using Savitzky-Golay smoothing with 25 points. Original ATR-FTIR data files are deposited with the accompanying Data in Brief article (Lutzke et al., 2019). X-ray photoelectron spectroscopy. X-ray photoelectron spectroscopy (XPS) was carried out using a Physical Electronics PE-5800 X-ray photoelectron spectrometer with an Al Kα X-ray source (1486.6 eV) (Chanhassen, MN, USA). Samples were stored in a desiccator prior to analysis and were distributed as a densely packed layer on double-sided carbon tape. Spectra were acquired in the range of 1100-10 eV with a pass energy of 187.85 eV and a step size of 1.6 eV at a take-off angle of 45°. Data was processed using MultiPak version 9.3.0.3 and chemical shift was referenced to the major carbon peak (C 1s) and adjusted to a binding energy of 284.8 eV. Original ATR-FTIR data files are deposited with the accompanying Data in Brief article (Lutzke et al., 2019). Additional details of sample preparation and data collection are in the Supplementary Information. 5. Conclusions Our ATR-FTIR analysis of enzymatically-isolated P. ponderosa sporopollenin indicates that the biopolymer consists of distinct aliphatic and aromatic domains. The majority of spectral features are shared with commercial Lycopodium sporopollenin isolated under acidic conditions (Fig. S34). The similarity of natural sporopollenin to cuticular polymers and waxes is revealed by comparison of ATR-FTIR spectra (Table S2) (Heredia-Guerrero et al., 2014; Ramirez et al., 1992) as well as the existence of biosynthetic pathways common to both the cuticle and sporopollenin (Li et al., 2010) We therefore find considerable justification for the continued perspective that sporopollenin consists largely of polymerized trans-4-hydroxycinnamic and fatty acids (Morant et al., 2007), although the primary form of connectivity is unlikely to be the saturated ester linkages that are prevalent in cutin. The majority of IR bands can be assigned directly to trans-4-hydroxycinnamyl units and simple aliphatic chains with varying degrees of

3. Experimental 3.1. Materials Glyceryl trioleate (99%) and sodium cyanoborohydride (95%) were obtained from Acros Organics (Morris Plains, NJ, USA). Acetic anhydride (> 97%), bromine (99.8%), β-carotene (99%), citrus pectin, di-nhexyl ether (98%), 1-hexadecanol (98%), trans-4-hydroxycinnamic acid 12

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Fig. 9. The relationship between IR band assignments and hypothesized structural components of sporopollenin.

References

oxidation (Fig. 9). Alcohols primarily occur in the form of 1,2- or 1,3diols that permit formation of cyclic acetals or ketals. The characteristic bands of isolated α-pyrones are not detected in ATR-FTIR spectra of sporopollenin, nor is there spectral evidence that carotenoids or other terpenoid structures are major constituents of sporopollenin. The aromatic components of sporopollenin are partially removed by hot phosphoric acid or acetolysis, and are eliminated altogether by peracetic acid and bromine. The morphology of individual exine grains is largely retained after removal of aromatics by bromine, which indicates that trans-4-hydroxycinnamic acid and other phenolic compounds are not critical structural components. Following removal of aromatic constituents, the aliphatic residual fraction is no longer chemically consistent with natural sporopollenin. As hypothesized by van Bergen et al. (1993), a similar form of oxidative degradation is probably the source of recalcitrant fossil sporopollenin, which is feasibly polymerized through ether linkages in a manner analogous to cutan (Villena et al., 1999) since acetals, esters, and other hydrolytically-sensitive functional groups possess limited stability. The physical and chemical attributes of sporopollenin cannot be meaningfully studied or discussed in the absence of a clear definition of the biopolymer, which requires an isolation method that preserves the chemistry of sporopollenin as it exists in nature. Many chemical treatments clearly produce sporopollenin derivatives that are spectroscopically distinct from the in planta biopolymer. In some cases, this derivatization is extreme and the resulting polymers are entirely depleted of aromatics (Domínguez et al., 1999) or other structural components (Shaw and Apperley, 1996) that are likely characteristic of native sporopollenin. For this reason, a practical chemical definition should qualify that true sporopollenin remains spectroscopically consistent with the natural exospore or exine following isolation. This comparison is straightforward using IR spectroscopy, which remains a convenient analytical technique for the identification of sporopollenin in plant tissue and at all stages of purification.

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