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Investigation of the fate of proteins and hydrophilicity/hydrophobicity of Lycopodium clavatum spores after organic solvent-base-acid treatment Md Jasim Uddin, Noureddine Abidi, Juliusz Warzywoda, and Harvinder Singh Gill ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.9b03040 • Publication Date (Web): 13 May 2019 Downloaded from http://pubs.acs.org on May 13, 2019

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ACS Applied Materials & Interfaces

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Investigation of the Fate of Proteins and Hydrophilicity/Hydrophobicity of Lycopodium clavatum Spores After Organic Solvent-Base-Acid Treatment Md Jasim Uddina, Noureddine Abidib, Juliusz Warzywodac, and Harvinder Singh Gilla,* a Department

of Chemical Engineering, Texas Tech University, Lubbock, TX 79409, USA of Plant and Soil Science, Texas Tech University, Lubbock, TX 79409, USA c Materials Characterization Center, Whitacre College of Engineering, Texas Tech University, Lubbock, TX 79409, USA b Department

*Corresponding author: Dr. Harvinder Singh Gill Texas Tech University Department of Chemical Engineering 8th Street and Canton Ave, Mail Stop 3121, Lubbock, TX 79409-3121, USA. Phone: 806-834-3682 Fax: 806-742-3552 E-mail: [email protected]

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Abstract Microcapsules extracted from lycopodium (Lycopodium clavatum) spores have been increasingly used as an oral therapeutic carrier. A series of sequential treatments involving acetone, KOH, and H3PO4 are used to extract protein-free hollow microcapsule. This study focuses on two critical aspects of lycopodium spores: the fate of native proteins and wettability of the spores after chemical treatment. Protein-free spores are desired to prevent an allergic reaction, while wettability is critical for formulation development. While the chemically-treated lycopodium spores are generally regarded as protein-free, the studies that have reported this have not gone into significant depths to understand the nature of residual nitrogen observed even in spores thought to be protein-free. Wettability of spores has not received any significant attention. Accordingly, in this study, we performed a comprehensive analysis of natural spores, and spores after each chemical treatment step. We show that natural lycopodium spores are hydrophobic and contain low molecular weight proteins (~ 10 kD). Acetone treatment partially solubilizes unsaturated phospholipids from the spores. Nevertheless, the acetone-treated spores retain native proteins and are still hydrophobic. KOH treatment, however, removes a significant amount of proteins and partially hydrolyzes esters to carboxylic acid salts, and results in a hydrophilic spore with good wettability. Finally, we show that H3PO4 treatment removes residual proteins, hydrolyzes remaining esters to carboxylic acids, and dissolves carbohydrates. H3PO4 treatment temperature controls carbohydrate dissolution, which in turn affects the hydroxyl functional groups and hydrophilicity (wettability) of the treated spores. Spores treated at 60 °C as opposed to 160 °C are amphiphilic in nature due to the abundance of hydroxyl functional groups on the surface. In conclusion, this study confirms the removal of native proteins from treated spores and sheds light on the chemical changes that the spores undergo after chemical treatment, and correlates these changes to their wettability. Keywords: FTIR; lycopodium spore; oral vaccination; phospholipids; pollen grains; proteins; TGA; sporopollenin.


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3

1. Introduction Plant reproductive materials are naturally transported in protective microcontainers, which are collectively called spores and pollens. Their outer wall known as exine is made of sporopollenin, a highly cross-linked biopolymer of aliphatic and phenolic compounds.1-3 The sporopollenin exine wall is mechanically robust4 and remarkably resistant to attack by most chemicals.5 Chemical inertness, physical robustness, monodispersity in size, unique surface architecture, nanoporous wall, and spacious internal cavity, collectively offer an excellent opportunity to use sporopollenin exine as a microcapsule for oral therapeutics.6-7 Such microcapsules can potentially protect acid- and enzyme-labile molecules from gastric degradation. Specifically, the microcapsule extracted from natural lycopodium spore (Lycopodium clavatum) has been increasingly used as a carrier for oral drugs8-9 and vaccines.6 The primary challenge of using lycopodium spore as a microcapsule is to ensure removal of all proteinaceous materials from its core to prevent any potential allergic reaction, and to simultaneously preserve its exine architecture in its pristine state. A well-established ‘conventional treatment method’10 and it's variants11-13 have been widely used to clean spores. The conventional treatment method utilizes an organic solvent (acetone), a base (KOH), and a non-oxidizing acid (H3PO4) to obtain protein-free sporopollenin microcapsules. Whether after these treatment steps the native proteins have been removed from the spores remains an important question. To address this question different approaches have been used. DiegoTaboada et al. used liquid-based methods to extract any native proteins from treated spores and subsequently used sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and mass spectrometry to analyze the proteins.14 None of the assays tested positive for proteins and the authors concluded that the treated spores were protein free. However, since an untreated natural spore was not included in these assays, the ability of the methods to extract proteins from the spores is debatable. In other studies, proteins were characterized by elemental analysis6, 10-11 and Fourier transform infrared (FTIR) analysis.15-16 However, lycopodium spore is a complex biopolymer of lipids, proteins, carbohydrates, and sporopollenin. Sporopollenin chemical structure is not fully resolved yet, hence, its contribution in the FTIR and elemental analysis is not well known. For instance, vibrations from sporopollenin may overlap with amide bond vibrations, thereby it may confound protein characterization. Elemental analysis, on the other hand, is an indirect method that lacks both sensitivity and specificity. In this method, nitrogen that is produced upon combustion of the spores is assumed to entirely originate from proteins in the spores. This nitrogen content is then multiplied by a conversion factor to convert all the measured nitrogen into protein. Such analysis possibly overestimates protein amount17 because the method is unable to discriminate the origin of measured nitrogen, which can come from proteins or from non-proteinaceous materials such as the spore wall constituent molecules. Protein measured by this method in the treated lycopodium spores has been reported to vary from zero10, 12-13, 18 to 0.94 (wt. %).6, 11 To solve this uncertainty of whether treated spores are truly protein free or not, there is a need to effectively extract proteins and analyze them using complementary techniques to determine the fate of proteins. Another consideration regarding the use of chemically-treated protein-free spores is the ability to properly disperse them in aqueous solutions to be able to prepare pharmaceutical formulations. A hydrophilic lycopodium surface can also enhance intestinal residence time of the spores by increasing their mucoadhesion.19 However, different reports have described conflicting results, some describing the treated spores to be hydrophilic20 while others



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4 describing them to be hydrophobic.13 An examination of the chemical treatment procedures of these studies reveals that the treatment conditions were not fully described, nonetheless, a lowtemperature treatment by Wang et al. seemingly produced hydrophilic spores20, and although Binks et al. did not specify treatment temperatures but their protocol produced hydrophobic spores.13 Majority of the current lycopodium treatment literature has not specified treatment temperatures. As such, the effect of temperature on spore wettability has largely remained understudied, and not much is known how the treatment steps affect wettability and whether the chemical treatment protocol can be tuned to control hydrophilic and hydrophobic nature of the spores. Therefore, in this study, we focused our attention on two aspects, first was to identify the fate of proteins during chemical treatment, especially after each treatment step and not just upon completion of the entire process, and second was to determine the effect chemical treatment conditions especially temperature have on spore hydrophilicity and hydrophobicity. To study the fate of proteins, we augmented elemental analysis with SDS-PAGE and matrixassisted laser desorption/ionization time of flight mass spectrometry (MALDI-TOF MS) methods to analyze spore proteins. X-ray photoelectron spectroscopy (XPS) in conjunction with FTIR was employed to address the bonding states of the residual nitrogen in the treated lycopodium spore. To obtain insight into the functional groups that can affect wettability, we used FTIR to characterize the spores at different treatment steps. Particular attention was given to how the treatment process affects C=O bond in lipids (a contributor for hydrophobicity) and –OH group in carbohydrates (a contributor for hydrophilicity). We used thermogravimetric analysis (TGA) to correlate the FTIR data to the nature of material being removed at a certain chemical treatment step. Insights gained from these analyses helped us to conclude that residual nitrogen observed in the treated spores is likely due to nitrogen from non-proteinaceous material and that chemical treatment at a lower temperature can increase the hydrophilicity of the treated spores.

2. Materials and methods 2.1.

Spores and chemicals

Lycopodium spore (Lycopodium clavatum) was purchased from Pfaltz & Bauer (Waterbury, CT, USA) or Sigma-Aldrich (St. Louis, MO, USA). All chemicals and materials were purchased from Fisher Scientific (PA, USA) unless otherwise stated. Lane marker reducing sample buffer and Coomassie Brilliant Blue G-250 dye were purchased from Thermo Fisher Scientific (Waltham, MA, USA). Tris/Glycine/SDS electrophoresis buffer, 4-20% Mini-PROTEAN TGX precast gels for SDS-PAGE, and protein standards (10-250 kD) were obtained from BioRad Laboratories, Inc. (Hercules, CA, USA). Sodium cacodylate buffer, 3% formaldehyde/3% glutaraldehyde solution, 4% osmium tetroxide, Reynold's lead citrate, and 4% uranyl acetate were purchased from Electron Microscopy Sciences (Hatfield, PA, USA). LX-112 resin, dodecenyl succinic anhydride (DDSA), nadic methyl anhydride (NMA), and 2,4,6-tri (dimethylaminomethyl) phenol (DMP-30) were purchased from Ladd Research (Williston, VT, USA). Ultrapure Milli-Q water (Direct-Q® 3 UV system, MilliporeSigma, MA, USA) with an electrical resistivity of 18.2 MΩ.cm at 25 °C was used in all applications. 2.2.

Preparation of hollow spore microcapsule

Protein-free lycopodium spores were prepared by the method described by Atwe et al.6 Briefly, 30 g of natural spores were stirred and refluxed in 350 ml of acetone at 70 °C for 12 h. The solution was cooled and filtered using a vacuum filtration unit (MilliporeSigma, Billerica, MA, USA). The filtrate was gently stirred and heated to 70 °C for 12 h to evaporate acetone. A light


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5 brown colored oily extract was obtained after acetone evaporation and used for analysis. After air-drying, acetone-treated spores were refluxed in 350 ml of 1 M (6% w/v) KOH for 12 h at 120 °C (KOH renewed after 6 h with a fresh solution). Spore solution was then cooled, diluted with water, filtered, and extensively washed with hot water, acetone, and 70% ethanol. It should be noted that filtrates obtained immediately after KOH treatment were dark brown in color. The water and organic solvent washings continued until a clear filtrate was obtained. The first spent KOH filtrate containing spore extract was dried at 100 °C. The dried residue was used for analysis. KOH-treated spores were next subjected to acidolysis by stirring them in 400 ml of 85% (w/w) H3PO4 at 60 °C or 160 °C for seven days. After acid treatment, spore solution was cooled, diluted with water, filtered, and extensively washed with hot water, acetone, 2 M HCl, 2 M NaOH, water, acetone, and 70% ethanol in a successive manner. Finally, H3PO4-treated spores were dried at 60 °C for 24 h in an oven and weight of the final dried spores was measured. Spore samples after each chemical treatment step were collected for analysis. All spores were stored at room temperature before use in the experiments. 2.3.

Scanning electron microscopy (SEM)

Spore surface morphology and structural integrity were visualized by an SEM (Hitachi S4300 E/N FESEM, NY, USA). To assess the extent of cellular materials removed from the spore core, lycopodium samples were manually fractured using mortar and pestle by first mixing the spores in dry ice to make them brittle, and imaged with SEM. 2.4.

CHN elemental analysis

Spore samples between 1.5 mg and 4 mg were analyzed using PerkinElmer 2400 Series II CHNS/O Analyzer (PerkinElmer, Inc., Waltham, MA, USA) to estimate carbon, hydrogen, and nitrogen content. All measurements were performed in triplicate. 2.5.

Transmission electron microscopy (TEM)

For TEM imaging, spores were fixed in 3% formaldehyde/3% glutaraldehyde solution in 0.1 M cacodylate buffer (pH 7.2) overnight at 4˚ C, washed in 0.1 M cacodylate buffer solution, and post-fixed in 1% osmium tetroxide for 2 h. Samples were then washed with water, stained in 1% uranyl acetate solution for 1 h, dehydrated in a graded series (25, 50, 75, 85, 95%, and 100%) of ethanol, and then 100% acetone. Epoxy resin was prepared by mixing 1.4% (v/v) DMP-30 in a resin mixture of 48.5% (w/w) LX-112, 28% (w/w) DDSA, and 23.5% (w/w) NMA. Fixed and dehydrated spores were then embedded in the epoxy resin according to a published protocol.21 Ultrathin sections (~60 nm) were cut with a diamond knife on a Leica UltraCut E microtome (Leica Microsystems Inc., Buffalo Grove, IL, USA). Sections were stained in 1% uranyl acetate solution for 30 min, washed with water, and stained again in Reynold's lead citrate solution for 5 min. After washing and air drying, specimens were examined using a Hitachi H-8110 TEM at 75 kV. 2.6.

SDS-PAGE

Proteins were extracted from lycopodium spores at different chemical treatment stages using a method published previously.22 Briefly, dry spores (20 mg) were first fragmented using a magnetic stir bar to facilitate protein extraction in 1 ml of phosphate buffered saline (PBS). Extracted proteins were then analyzed with SDS-PAGE.17



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6 2.7.

MALDI-TOF MS

Proteins from natural and chemically-treated lycopodium spores were extracted in ultrapure Milli-Q water (40 mg spores/ml of water) using the method described earlier in section 2.6. The extracted sample (2 µl) was mixed with 4 μl of α‐cyano‐4‐hydroxycinnamic acid (α-CHCA) matrix solution consisting of 5 mg/ml of α-CHCA in 50% acetonitrile and 0.1% trifluoroacetic acid (TFA). The mixture (0.5 μl) was spotted on a MALDI plate. The mass spectra were acquired in the reflector ion mode using 4800 Plus MALDI TOF/TOF mass spectrometer (Applied Biosystems, CA, USA), and a total of 2500 laser shots were accumulated per spectrum. The mass range was selected to be between 400 and 5000 m/z and data was collected in positive ion mode. Extracted samples were also analyzed in the linear ion mode using sinapinic acid matrix solution for the mass range between 5000 and 50000. 2.8.

Attenuated Total Reflectance (ATR)-FTIR

FTIR spectra of manually fragmented spore samples were recorded using Spotlight 400 FTIR (PerkinElmer, Inc., Waltham, MA, USA) equipped with a Universal ATR accessory and ZnSe-diamond crystal composite (single-bounce). All FTIR spectra were collected at a spectral resolution of 4 cm-1, with 32 co-added scans over the range from 4000 to 650 cm-1. A background scan of the clean ZnSe-Diamond crystal was acquired before scanning the samples. The PerkinElmer Spectrum was used to perform spectra normalization and baseline corrections. 2.9.

XPS

XPS measurements were performed on a Physical Electronics PHI 5000 VersaProbe spectrometer (Physical Electronics, Inc., Chanhassen, MN, USA) using a monochromatic Al Kα (hν = 1486.6 eV) X-ray source. Peaks reported were charge corrected using the adventitious carbon C 1s peak at 284.8 eV as the reference. 2.10.

TGA

TGA of spore samples was performed using the Pyris 1 thermogravimetric analyzer (PerkinElmer, Shelton, CT, USA). Samples between 1.5 mg and 2 mg were used in each analysis. Thermograms were recorded between 37 °C and 600 °C with a heating rate of 10 °C/min in a flow of nitrogen at 20 ml/min. The Pyris software was used to calculate the first derivatives of the thermograms (DTG) and the percent weight loss for each sample. 2.11.

Statistical analysis

Statistical analysis was performed using GraphPad Prism 6 (GraphPad Software, Inc., La Jolla, CA, USA). A two-tailed t-test was used to analyze data from elemental analysis.

3. Results 3.1.

Elemental analysis reveals presence of nitrogen in chemically treated spores

Elemental analysis has been widely used to answer the question whether, after chemical treatment, proteins have been removed from the spores or not. In this method, nitrogen in the treated spores is measured and multiplied with a factor of 6.25 to convert it into protein amount.6


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7 Natural lycopodium spores were sequentially treated with acetone, KOH, and H3PO4 to remove proteinaceous materials and to obtain hollow spore shells. After treatment, lycopodium spores retained their structural integrity, homogeneity, and surface architecture as seen in the SEM images (Figure 1A: natural, B: treated). There was no change in the trilete scar (Figure 1C: natural, D: treated) or the reticulated microstructure morphology (Figure 1E: natural, F: treated) indicating that spores were not damaged. Spore size slightly decreased from 31.1 ± 1.3 µm (natural spores) to 28.7 ± 1.6 µm (H3PO4-treated spores) after chemical treatment, and this agrees with other studies.11, 20 Manually broken natural lycopodium spores showed sparsely distributed cellular material inside the spore shell (Figure 1G), which were absent after chemical treatment (Figure 1H). Approximately 77.9 ± 1.3% of the spore mass was removed after chemical treatment. Elemental analysis of natural and treated spores demonstrated a significant reduction (p < 0.0001) in nitrogen content after chemical treatment (Table 1). This reduction level is similar to our previously published result.6 It should be noted that the range of measured nitrogen in the H3PO4-treated spores was 0.94 µg/1.56 mg treated spores to 2.64 µg/2.03 mg treated spores. Since the detection limit of the instrument is 1 µg, some of these values fall below the detection limit. This adds some uncertainty as to how much nitrogen really remains in the spores after H3PO4 treatment, but on average less than about 1.63 µg nitrogen/1.74 mg treated spores is left. Although not as sensitive as elemental analysis, confocal microscopy also confirmed that the spores were clean from the inside (Figure S1). TEM images of natural spores showed the presence of both exine and intine wall (Figure 2A-B), similar to an earlier report.23 In contrast, H3PO4-treated spores showed only exine wall (Figure 2C-D). The average exine wall thickness was 1.62 ± 0.95 µm. Similar to SEM (Figure 1G) and confocal (Figure S1: Natural) micrographs, TEM image of the natural spore (Figure 2A) showed sparsely distributed cellular material inside the spore shell. Figure 1. Scanning electron micrographs of lycopodium spores. (A) Before chemical treatment. (B) After chemical treatment showing intact spores. (C) Proximal face of a natural spore before chemical treatment showing trilete (Y-shaped) scar. (D) Proximal face of a treated spore showing the inflated trilete scar. (E) Distal face of a natural lycopodium showing reticulated (honeycomb-like) architecture made from perforated muri (walls) sitting atop the intact spore shell. (F) Distal face of a treated lycopodium showing intact muri. (G) Manually broken natural spore showing cytoplasmic contents residing inside the core. (H) Manually broken treated spore showing a clean core devoid of cytoplasmic material. Figure 2. Transmission electron micrographs of ultrathin sections of lycopodium spores. (A) A natural lycopodium spore showing its double-layered wall and cytoplasmic material (CM) contained inside it. The grey wavy material in the enclosed space is the resin. (B) Two distinct layers (exine and intine) of the natural spore wall, ex = exine, int = intine. (C) A H3PO4-treated lycopodium spore. (D) Wall of the H3PO4treated spore showing that the exine layer remains after chemical treatment.

Table 1. Elemental analysis (wt. %) of lycopodium spores

Natural Acetone-treated KOH-treated H3PO4-treated

Carbon

Hydrogen

Nitrogen

65.47±0.04 60.27±2.25 58.69±0.09 72.33±0.36

9.53±0.25 8.52±0.35 8.14±0.42 7.99±0.09

1.25±0.06 1.84±0.07 0.71±0.03 0.09±0.04



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8 3.2.

Protein content based on gel electrophoresis

Since elemental analysis indicated that nitrogen was present in the H3PO4-treated spores, and this may originate from residual proteins, we decided to test for the presence of proteins in treated spores using SDS-PAGE. However, lycopodium spores do not possess any micron-sized apertures as do pollen grains,17 and this makes protein extraction challenging. Thus, perhaps not unsurprisingly, we were unable to detect proteins even in intact natural spores (Figure S2). We postulated that perhaps the proteins could be extracted if the spores could be broken. Therefore, we manually broke both the natural and the chemically treated spores and then extracted proteins. SDS-PAGE analysis of extract from these broken spores indeed showed an intense band of low molecular weight proteins (~12 kD) in the natural spores (Figure 3: lane 2). Acetone treatment did not remove these proteins since the band persisted after acetone treatment (Figure 3: lane 3). In contrast, after KOH treatment, majority of the protein band disappeared except a faint smear at less than 10 kD (Figure 3: lane 4). Finally, after H3PO4 treatment a clear lane in the gel electrophoresis image was seen (Figure 3: lane 5), indicating complete removal of proteins. Figure 3. SDS-PAGE analysis of manually broken lycopodium spores at different chemical treatment steps. Natural spores were treated with acetone, 6% (w/v) KOH, and 85% H3PO4. Spore samples at different chemical treatment stages were collected, then manually broken, and extracts were obtained from them. Lane 1: molecular weight marker; lane 2: extract from natural lycopodium spores before chemical treatment; lane 3: extract from acetone-treated spores; lane 4: extract from KOH-treated spores; lane 5: extract from H3PO4-treated spores.

3.3.

Protein analysis using mass spectrometry

Although elemental analysis had demonstrated the presence of nitrogen in the H3PO4treated spores, yet, no protein could be detected through SDS-PAGE. The inability to detect proteins through SDS-PAGE could mean that either the proteins were indeed removed via chemical treatment, or that they were present in an amount that could not be detected with gel electrophoresis. SDS-PAGE with Colloidal Coomassie Blue G-250 staining can detect up to 1 ng protein in a sample24 whereas MALDI-TOF MS can detect protein in attomolar (10-18 mol/L) concentration.25 We therefore further analyzed the spores with MALDI-TOF MS. The mass spectra obtained from lycopodium spore extracts at various treatment stages are shown in Figure 4 (reflector ion mode for peptides) and Figure S3 (linear ion mode for proteins). Proteins could be seen in natural and acetone-treated spores, both showing a similar spectrum, which indicates that as one would expect, acetone treatment does not significantly remove proteins. In contrast, KOH- and H3PO4-treated spores showed lack of proteins and peptides. The residual peaks in the H3PO4-treated spore may originate from α-CHCA matrix material that was used to ionize the sample (Figure 4) and a trace amount of sporopollenin materials (such as p-coumaric acid and ferulic acid). Figure 4. MALDI-TOF MS analysis of peptides extracted from lycopodium spores after different chemical treatment steps. Proteinaceous materials were extracted from ground spore samples using ultrapure water, mixed with α-CHCA matrix solution, and analyzed in reflector ion mode.

3.4.

FTIR analysis of lycopodium spores

We next used FTIR analysis to understand the functional groups present on the spores. We analyzed amide bond vibrations to track proteins, and vibrations of functional groups that


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9 can affect spore wettability. The FTIR spectra of natural, acetone-treated, KOH-treated, and H3PO4-treated lycopodium spores (Figure 5) showed major differences after receiving the respective treatments. Figure 5. Fourier transform infrared (FTIR) spectra of manually broken lycopodium spores at different chemical treatment stages. Natural spores were successively treated with acetone, 6% (w/v) KOH, and 85% H3PO4 (160 °C for 7 days) to obtain treated spores.

3.4.1. FTIR vibrations relevant to proteins To detect proteins, we inspected the region around 1660-1630 cm-1 and 1567-1530 cm-1 since these vibrations have been assigned to amide I (C=O stretching) and to amide II (in-plane N–H bending and C–N stretching), respectively, in proteins.16, 26-27 However, peaks at these vibrations were not observed in any of the samples - natural, acetone-treated, KOH-treated, or H3PO4-treated spores (Figure 5). We wondered why we could not observe these observations. Since we had confirmed the presence of proteins in the KOH extract using both SDS-PAGE and MALDI-TOF MS, we decided to examine the KOH extract for the protein vibrations. We first dried the KOH extract and then analyzed it. Indeed, we were able to observe distinctive peaks at 1635 and 1562 cm-1 (Figure S4), which were absent in the pure KOH sample. These vibrations can thus be respectively assigned to the amide I and amide II vibrations in proteins found in lycopodium spores. Thus, while the ATR-FTIR technique was unable to detect proteins in the spore samples, the technique successfully detected them in the KOH extract. 3.4.2. FTIR vibrations relevant to wettability analysis Hydrophilic contributors: Vibration at 3300 cm-1: This broad vibration is assigned to O–H stretching28 (hydrogen bonded) in carboxylic acids, alcohol, phenol, carbohydrates, or water. The intensity of this vibration increased and shifted to 3360 cm-1 following KOH treatment. This could be due to the removal of compounds (such as lipids) which may have been linked to –OH groups, thus, allowing water to adsorb. However, after H3PO4 treatment a dramatic decrease was seen in the intensity of this vibration. This could be due to the removal of carbohydrates or reduction in adsorbed water. Vibration at 1583 cm-1: This vibration appeared only as a sharp vibration in the spectrum of KOH-treated sample. This vibration was due to the asymmetric stretching of carboxylate (COO-)29 which can form after KOH hydrolysis of esters into carboxylic acid salt. Hydrophobic contributors: Vibration at 3007 cm-1: This is assigned to =C–H stretching vibration in aliphatics, which can arise from unsaturated fatty acid esters.30 This vibration completely disappeared after H3PO4 treatment suggesting the removal of unsaturated compounds. Vibrations at 2924 and 2854 cm-1: These two sharp vibrations are attributed to asymmetric and symmetric C–H stretching in methylene group (–CH2–), respectively.31 Chemical treatments did not appear to significantly affect the intensities of these aliphatic vibrations.



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10 Vibration at 1744 cm-1: This strong vibration is assigned to C=O stretching of aliphatic ester groups, primarily from fatty acids or lipids.28 This vibration disappeared after H3PO4 treatment. Vibration at 1710 cm-1: This is attributed to the carbonyl group (C=O) stretching vibration that can originate from alkyl esters or carboxylic acids.5, 32 Presence of this peak in the spectrum of natural lycopodium indicated high amounts of alkyl esters. A slight decrease was noticed after acetone treatment. However, treatment with aqueous KOH drastically reduced the amount of these alkyl esters. The peak reappeared after H3PO4 treatment suggesting formation of carboxylic acids. Vibration at 1654 cm-1: This vibration is assigned to C=C in sporopollenin.32 This vibration was present in all lycopodium spectra. Vibration at 1606 cm-1: This vibration is assigned to C=C vibration in the aromatic ring of sporopollenin.16 It was present in all spectra except in the spectrum of KOH-treated lycopodium where it appears as a shoulder. Vibration at 1517 cm-1: This vibration appeared in all samples and is attributed to C=C vibration in phenolic components.16 The intensity decreased after H3PO4 treatment and shifted to 1500 cm-1. Vibration at 1440 cm-1: This is attributed to C–H vibration in the aromatic ring of sporopollenin.16 This vibration exists as a sharp peak in all the spectra of lycopodium samples. This vibration shifted to 1448 cm-1 after H3PO4 treatment. Vibration at 1377 cm-1: This is attributed to C–H bending vibration in methyl (–CH3) group of lipids or carbohydrates.31 This vibration was present in all the lycopodium spectra except H3PO4 treated sample where it appears as a shoulder. This suggests that H3PO4 treatment drastically reduced carbohydrates. Vibration at 1344 cm-1: This is attributed C–CH3 bending26 in carbohydrates. The vibration appeared as a shoulder in natural and acetone-treated spores. A distinctive vibration was observed in the KOH-treated spores while the vibration disappeared after H3PO4 treatment. Vibration at 1280 cm-1: This is attributed to C–O stretching of the alkyl esters. This vibration exists in the spectra of the natural and acetone-treated samples. However, this vibration completely disappeared after KOH treatment. Vibration at 1260 cm-1: This vibration is due to C–O stretching in carbohydrates26 which was seen in all spore samples except H3PO4-treated spores. Vibrations at 1140 and 1110 cm-1: These vibrations were sharp in the spectra of natural and acetone-treated lycopodium spore samples. The intensity of the 1110 cm-1 band slightly decreased while the vibration 1140 cm-1 shifted to 1132 cm-1 following KOH treatment. Both vibrations completely disappeared after H3PO4 treatment. These vibrations are attributed to the pyranose ring of sugars (skeletal vibration of C–OH and C–O–C).33 This result indicates that the treatment with H3PO4 affected carbohydrate backbone, and possibly removed cellulosic intine. Vibration at 999 cm-1: This vibration is attributed to the C–O–C stretching in carbohydrates34 and it completely disappeared after H3PO4 treatment.


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11

Vibrations around 847-814 cm-1: These vibrations are attributed to the aromatic ring vibration in sporopollenin.34 Vibration around 723 cm-1: This vibration is attributed to C–H rocking (bending) in methylene group.35 This vibration was present in all spore samples. 3.5.

XPS analysis of lycopodium spore surface reveals non-proteinaceous nitrogenous component

Elemental analysis showed that trace amounts of nitrogenous components remained in the H3PO4-treated spores (Table 1) while MALDI-TOF MS analysis showed an absence of proteins in the same sample, suggesting that the nitrogen may be part of the sporopollenin constituent compounds and not proteins. Hence, we used XPS to study the nature of the nitrogenous components present in the lycopodium spores. Survey scan XPS spectra (analysis depth ~7 nm) showed the presence of C, N, and O in both the natural and treated lycopodium spore samples (Figure S5 and Table S1). Other elements, namely Al, P, S, and K were also observed in natural spores, out of which just P was seen in the final H3PO4-treated spores. After KOH treatment a trace amount of Mg and Si appeared, which can be attributed to the presence of these elements in commercial KOH pellets used to prepare the KOH solution. However, they were removed after H3PO4 treatment. After H3PO4 treatment, the percentage of N decreased significantly. In addition, the element Na was observed, which could be due to the NaOH wash step of the sample. High-resolution scan XPS spectra (Figure 6) were also acquired to obtain the bonding states of C, N, and O on the lycopodium spore surface. The C 1s peak (Figure 6) was deconvoluted into four components: 284.8 eV (C–C or C–H); 286.2 eV (C–N or C–OH); 287.5 eV (C=O); and 289.1 eV (C=O in ester).36-37 The C 1s peak of H3PO4-treated lycopodium lacked 289.1 eV peak, suggesting the absence of ester on the spore surface. Although sporopollenin is generally considered to be a nitrogen-free structure,1, 16 trace amounts of nitrogenous components were observed in the XPS analysis. The N 1s peak was resolved into 398.3-398.5 eV (pyridine-like structure, i.e., C–N=C) and 399.9 eV (pyrrolic-N) components (Figure 6).37-39 Thus, the N 1s spectra suggest that the source of nitrogen in the sporopollenin is likely of non-proteinaceous origin. The O 1s peak (Figure 6) was deconvoluted into three components: 531.7 eV (C=O in carboxylic acid or ester); 532.5 eV (C–OH); and 533.6 eV (C–O in ester or O–H in carboxylic acid).36 A new peak appeared in the KOH-treated lycopodium at 531.2 eV instead of 531.7 eV suggesting a formation of COO-.36, 40 The deconvolution of O 1s peak for H3PO4-treated lycopodium spores is less certain, owing to the presence of traces of Na in this sample (Figure S5 and Table S1) and the resulting possible overlap of the Auger Na KLL lines and the photoelectron O 1s line.41

Figure 6. High-resolution scan X-ray photoelectron spectroscopy (XPS) spectra of C 1s, N 1s, and O 1s in lycopodium spores at different chemical treatment stages. Dots represent the measured value and the solid black line represents the XPS fitted curve.



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12 3.6.

Assessment of wettability of lycopodium spores

To assess whether the spores after undergoing the specific chemical treatment steps were hydrophobic or hydrophilic, we used two qualitative methods: first the ability of spores to form ‘liquid marbles’42 and second their degree of dispersion in water. If spores are hydrophobic they are expected to form liquid marbles and to not disperse well in water, and vice versa. It was seen that natural, acetone-treated, and H3PO4-treated spores formed liquid marbles, whereas KOH-treated spores did not form liquid marbles (Figure 7). Conversely, natural, acetone-treated, and H3PO4-treated spores did not disperse well in water, whereas KOH-treated spores did (Figure 7). Figure 7. Wettability of lycopodium spores after different chemical treatment. Natural spores were successively treated with acetone, 6% (w/v) KOH, and 85% H3PO4 (160 °C) to obtain treated spores. Photographs of lycopodium-coated water marbles are resting on a glass slide (top row). Dispersion of lycopodium spores in water (bottom row). Only KOH-treated spores did not form water marble and readily dispersed in water.

3.7.

Acetone treatment partially removed phospholipids from natural lycopodium spores

Acetone is an organic solvent and is expected to remove lipids. However, after acetone treatment, the spores still showed hydrophobic behavior. Therefore, we wanted to better understand the effect of acetone treatment on lycopodium spores. We filtered the acetone-spore mixture to recover spent acetone, evaporated the acetone from the filtrate by gentle heating, and analyzed the recovered oil with FTIR and TGA (Figure 8). The FTIR spectrum of the extracted oil showed strong lipid-associated vibrations at 3006, 2923, 2853, 1744, 1711, 1464 (C–H scissoring in the methylene group),16, 34 and 723 cm-1 (Figure 8A). The spectrum also provides critical information regarding the nature of the lipids in lycopodium spores. The vibrations at 1239, 1163, and 1097 cm-1 are the PO2- asymmetric stretching, C–O–PO2stretching, and PO2- symmetric stretching in phospholipids, respectively.26, 43 Hence, this shows that acetone treatment removed unsaturated phospholipids from the natural lycopodium spores. The removal of lipids was however partial since the lipids continued to remain associated with the spores, which is indicated by lipid-associated vibrations in the FTIR spectrum of acetonetreated spores (Figure 5). This can explain why acetone-treated spores retained hydrophobicity (Figure 7). TGA of the extracted phospholipids (Figure 8B, C) showed a two-stage decomposition similar to that reported in an earlier study.44 Light volatile compounds decomposed first with the maximum rate of decomposition at 213 ˚C and accounted for 24% weight loss; the rest of the material decomposed at the second stage with the maximum rate of decomposition at 390 ˚C. Figure 8. Characterization of the oily extract obtained from natural lycopodium spores using acetone treatment. (A) Fourier transform infrared (FTIR) spectrum of oily extract (photograph), (B) thermogram of the extract, and (C) first derivative thermogram (DTG) of the extract.

3.8.

Thermal degradation characteristics of lycopodium spores

To help correlate the changes observed in functional groups to the known constituent materials that are naturally found in the spores, we performed TGA. The characteristic degradation temperatures of lipids, proteins, and carbohydrates are different, and this can be used to study their presence in a complex mixture. To study thermal decomposition


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13 characteristics, lycopodium spore samples after each chemical treatment step were pyrolyzed in a controlled nitrogen environment. Figure 9A shows the percent weight loss as a function of temperature. All samples showed a continuous weight loss from 37 °C to 600 °C. After pyrolysis, natural, acetone-treated, KOH-treated, and H3PO4-treated spores lost approximately 85, 86, 80, and 56% of their mass, respectively. DTG of the spore samples showed five distinct regions of thermal decomposition (Figure 9B). Region I, located between 37 °C and 100 °C (Figure 9B), accounts for 1.7% weight loss for natural lycopodium, 2.3% for acetone-treated, 4.5% for KOH-treated, and 1% for H3PO4treated lycopodium. In this region, the weight loss was essentially due to adsorbed water. This water was weakly adsorbed as it was removed as soon as the temperature started to increase. H3PO4-treated spores adsorbed the least amount of water. Region II, between 100 °C and 250 °C (Figure 9B), accounts for 7% weight loss for natural spore and 4.9% for the acetone-treated spore sample. The KOH- and H3PO4-treated samples did not show a decomposition peak in this region but instead had a flat profile. This shows that KOH treatment leads to removal of components that are responsible for weight loss in Region II marked by the peak around 199-229 °C. Since KOH treatment hydrolyzed and removed proteins (Figure 3-4 and Figure S3-S4),45 the decomposition peak in this region could be due to proteins. A previous report also suggests similar protein content in natural lycopodium spores.6 Region III is located between 250 °C and 325 °C (Figure 9B). It accounts for 12.2% weight loss for natural spores. A decomposition peak at 300 °C, exhibited by natural spores, was missing in the acetone-treated and subsequent samples (Figure 9B). This suggests that treatment with acetone removed the compounds responsible for the weight loss in Region III marked by the peak at 300 °C. Since acetone treatment at least partially removed lipidic components (Figure 8),17 the decomposition at 300 °C in natural spores could be due to the lipids. Region IV, located between 325 °C and 390 °C (Figure 9B), is attributed to cellulose pyrolysis and decomposition. This is because the temperature for the maximum rate of decomposition for cellulose is 355 °C.46 It accounts for 28.5% weight loss for natural lycopodium, 60.7% for acetone-treated spores, and 48.3% for KOH-treated spores. The decomposition peak of Region IV was around 343-369 °C and was found in all spore samples except those treated with H3PO4. This suggests that H3PO4 treatment likely removed cellulose from spore intine. TEM images (Figure 2) also showed a lack of distinct intine wall in the H3PO4treated spore. Our result supports a previous notion that H3PO4 treatment removes cellulosic intine from lycopodium spores.47 Region V, located between 390 °C and 500 °C (Figure 9B), can be attributed to partial decomposition of the sporopollenin wall materials17 because by the end of H3PO4-treatment, the spores are already devoid of proteins, lipids, and cellulose and contain primarily sporopollenin. The sporopollenin decomposition peak in region V was present in all spore samples. However, this degradation behavior was slightly different in KOH-treated samples. KOH-treated spores contain not only sporopollenin but also cellulosic material, which could explain why the degradation behavior of sporopollenin is different. Specifically, it has been seen previously that the decomposition behavior of pure components changes in the presence of other components.48 Thus, it seems reasonable to speculate that the presence of cellulose and other materials can change the degradation profile of sporopollenin.



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14

Figure 9. Thermal decomposition of lycopodium spores after each chemical treatment stage. Natural spores were successively treated with acetone, 6% (w/v) KOH, and 85% H3PO4 (160 °C) to obtain treated spores. (A) TGA thermograms, (B) first derivative thermograms (DTG) of lycopodium spores showed five distinct regions defined by the Roman numerals.

3.9.

Temperature in H3PO4 treatment step controls hydrophilicity of the spore

Spores treated with H3PO4 at 160 ˚C were hydrophobic and were poorly dispersed in water (Figure 7), which is an undesirable property when making drug formulations. Thus, we wondered if we could devise a chemical treatment process, which could yield spores that are more readily dispersible in water. Our FTIR analysis of the spores coupled with the thermal analysis suggested that the spores after H3PO4 treatment loose significant amount of cellulosic and other complex carbohydrate molecules, which in turn reduces the –OH groups. In contrast, spores after KOH treatment were hydrophilic and had a stronger O–H vibration. Thus, we postulated that if –OH groups could be retained by reducing degradation of cellulose, the hydrophilic nature of spores could be retained. It is important to note that the H3PO4 treatment step cannot be simply eliminated because it is responsible for removing a significant amount of nitrogen-containing material, very likely proteins (Table 1). Thus, to reduce loss of cellulose during H3PO4 treatment, we reduced the treatment temperature to 60 ˚C hypothesizing that the lower temperature will reduce the rate of cellulose degradation. Lycopodium spores treated with H3PO4 at 60 ˚C for 7 days (Figure 10A) showed a marked difference in FTIR spectra compared to those treated at 160 ˚C (Figure 5). Specifically, O–H vibration (3360 cm-1) and vibrations in the carbohydrate fingerprint region (1200-900 cm-1) increased after low-temperature treatment. Furthermore, TGA of the spores treated at 60 ˚C (Figure 10B) showed one additional degradation of cellulose at 351 ˚C suggesting that the increase in O–H vibration was due to the carbohydrate material. This had a profound impact on sporopollenin shell aqueous dispersion. Hydrophilicity drastically increased when spores were treated with H3PO4 at 60 ˚C instead of 160 ˚C. Spores treated at 60 ˚C were easily dispersible in water and canola oil (Figure 10C), indicating an amphiphilic nature of the H3PO4-treated spores. Besides H3PO4 treatment temperature, H3PO4 treatment time also affects surface functional groups. Spores treated for three days regardless of treatment temperature (60 ˚C or 160 ˚C) showed strong COO- vibration at 1583 cm-1 (Figure 10A). Prolonged H3PO4 treatment gradually converted COO- to carboxylic acids, thus COO- vibration disappeared after seven days of treatment for both 60 ˚C (Figure 10A) and 160 ˚C (Figure 5) treated spores. Hydrophilicity of the spores treated at 160 ˚C was thus affected by this change since the treated spores lacked both COO- and –OH groups. However, the loss of hydrophilic COO- was counterbalanced by the –OH groups in the spores that were treated at 60 ˚C. It is to be noted that spores treated at 60 ˚C showed similar mass spectra as those treated at 160 ˚C (data not shown), suggesting that spores are protein-free. Therefore, spores treated at 60 ˚C produced protein-free shell with improved hydrophilicity. Figure 10. Effects of H3PO4 treatment time and temperature on lycopodium spore chemistry. KOHtreated spores were treated with H3PO4 for 3 days or 7 days at 60 ˚C or 160 ˚C. (A) Fourier transform infrared (FTIR) spectra of the treated spores, (B) thermogram and first derivative thermogram (DTG) of the spores treated at 60 ˚C for 7 days, and (C) dispersion of H3PO4-treated spores (Day 7, 60 ˚C) in water and canola oil showing amphiphilic nature of the treated spores.

4. Discussion


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15 Lycopodium spores have lately received significant attention as an oral drug delivery system. To prepare spores for such use, they are first cleaned by subjecting them to a series of chemical treatment steps. Conventionally, a chemical treatment protocol comprising of sequential treatment of the spores with acetone, KOH, and H3PO4 is used to obtain clean spores.6, 9-10, 15, 20 The present study was motivated to better understand two key aspects of this chemical treatment process. First was to track the fate of intrinsic proteins contained in lycopodium spores after each chemical treatment steps, and second was to determine how the different chemical treatment steps affect the wettability property of the spores. This knowledge is critical because it can help to tailor and tune the chemical treatment step so that not only are the native proteins removed to eliminate risk of potential protein-based allergic reactions, but the spores can also form good suspensions to support development of drug and vaccine formulations. Proteins in the lycopodium spore are primarily located inside the core enclosed by the sporopollenin wall, which has a hydrophobic coating. Because of these barriers, gentle protein isolation methods such as incubation of spores in water or PBS are inadequate to isolate proteins. Therefore, we manually fractured the sporopollenin wall to facilitate protein extraction. To better understand the fate of proteins, different analytical methods were used, among which nitrogen elemental analysis was the least sensitive method. Elemental analysis suggested that a trace amount of nitrogen remained in the spores after treatment. SDS-PAGE and MALDI-TOF MS, both of which are more sensitive and specific techniques than elemental analysis showed the presence of low molecular weight proteins in the natural lycopodium spores. Proteins with molecular weight greater than 10 kD were not found. This is an interesting data since in the past we have seen that unlike lycopodium spores, natural pollen grains, such as the ragweed pollen, do contain large molecular weight proteins.17 Both SDS-PAGE and MALDI-TOF MS techniques showed that acetone treatment did not significantly remove the proteins, but after KOH treatment these proteins were significantly removed, and were undetectable after H3PO4 treatment. The trace amount of residual nitrogen reported by elemental analysis of the H3PO4treated spores was later confirmed to be non-proteinaceous in nature by XPS analysis. To track proteins, we also used ATR-FTIR. Despite noticing a significant amount of proteins in the natural spores using SDS-PAGE and MALDI-TOF MS techniques, we did not see the two major protein vibrations (i.e., amide I at 1660-1630 cm-1 and amide II at 1567-1530 cm-1) in the FTIR spectrum of natural lycopodium spores. We also searched the literature to determine if the FTIR amide vibrations for natural Lycopodium clavatum spores had been reported in the published literature. Based on this search (Table S2), to the best of our knowledge, previous literature has also not specifically identified FTIR amide vibrations for Lycopodium clavatum spores. Several factors could explain this. First, the ATR-FTIR technique suffers from shallow depth of penetration of infrared (IR) beam into the sample. The depth of penetration of IR beam is less than 1.2 µm at 1600 cm-1 and it decreases with the increase in wavenumber.49 Since both TEM and confocal micrographs suggested that biomolecules were located largely inside the lycopodium cavity, and the wall that surrounds it has an average thickness of 1.62 ± 0.95 µm, it is likely that thick sporopollenin wall limits the IR beam interaction with spore biomolecules.35 This hypothesis is further strengthened by the observation that SDSPAGE did not reveal protein bands when intact natural spores were used for protein extraction. This suggests that proteins are either not present on the outer surface of the spores or are present in amounts that cannot be detected. The spores had to be cracked to be able to extract the proteins. It was surprising though, that our attempts to increase the FTIR spectrum intensity by using manually broken spores was also not successful. We postulated that this could be due to the sparse distribution of the biomolecules in the natural spores. To test this postulate, we used diffuse reflectance infrared Fourier transform spectroscopy (DRIFTS) to identify amide II



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16 vibration from manually broken lycopodium spores. DRIFTS measurement method provides bulk chemical characteristics of powdered sample as opposed to ATR-FTIR which provides surface characteristics.50 Before using DRIFTS on lycopodium spores, we first confirmed that the DRIFTS can indeed correctly identify amide II vibration by examining manually broken natural ragweed pollen. We were able to confirm the presence of amide II vibration at 1551 cm-1 in this ragweed pollen sample (Figure S6 and Figure S7: Natural ragweed pollen), which agrees with our previous ATR-FTIR studies on ragweed.17, 45 However, upon using the same preparation and measurement method of DRIFTS we were unable to find a distinct amide II vibration in the manually broken natural lycopodium spores (Figure S6 and Figure S7: Natural lycopodium). We then postulated that the thick sporopollenin wall and/or low protein concentration in lycopodium spore might be hindering detection of amide II vibration. To overcome these constraints, we then attempted to bring out the lycopodium spore proteins by manually breaking the lycopodium spores, then submerging these ground natural spores in ultrapure water for 24 h, and subsequently drying at 45 ºC. However, these hydrated spore sample also did not show any presence of amide II vibration (Figure S6 and Figure S7: Natural lycopodium hydrated). This could be due to the low protein concentration that may fall below the detection limit of FTIR method. Consistent with this notion, elemental nitrogen analysis shows that lycopodium spores contain approximately 7.3% protein (wt%)6 as opposed to ragweed pollen that contains approximately 26% protein (wt%).22 The only vibration that was observed in the protein region was at 1654 cm-1. This was observed in natural spores and in spores after every chemical treatment step, including the final H3PO4 treatment step. Many earlier reports have assigned this vibration to the amide I of proteins.16, 27, 51 However, our work shows that the final spore samples after H3PO4 did not contain proteins, and neither was the second characteristic protein vibration of amide II region seen in their FTIR spectra. Thus, the vibration at 1654 cm-1 cannot be from proteins. We propose that this vibration is from the strong C=C vibration (1650 cm-1) from sporopollenin wall aromatic rings.2 A similar vibration was also present in other pollen shells such as of ragweed and sunflower, from which the proteins had also been removed.17 Acetone treatment partially removed labile phospholipids, but it did not have a major impact on protein content or hydrophobicity. KOH treatment, on the other hand, had a profound impact on sporopollenin chemistry and led to significant protein removal. Lycopodium spores underwent two major changes following KOH treatment. First, the treatment removed cytoplasmic materials and proteins as is evident visually from confocal micrographs and analytically through the various techniques that were used; the elemental analysis showed reduction in nitrogen content, the protein band was no longer visible in the gel electrophoresis image, and protein peaks in the MALDI-TOF MS analysis were no longer seen. The proof that KOH treatment was removing proteins was obtained by analyzing the spent KOH that was recovered after treatment. FTIR spectrum of the spent KOH showed strong protein related vibrations (1635 and 1562 cm-1) which concurs with our earlier study.45 TGA analysis of the KOH-treated samples also showed a lack of decomposition around 199-229 °C which again suggests that this might be due to the removal of a large fraction of proteins. The second major change was that KOH treatment drastically increased treated spore hydrophilicity. FTIR analysis provides a possible mechanism of how KOH treatment may alter wettability. Vibrations related to unsaturated fatty acid esters (3007, 1744, and 1710 cm-1) decreased while a new vibration at 1583 cm-1 attributable to COO- appeared in the KOH-treated spores. This suggests that aqueous KOH solution hydrolyzed unsaturated fatty acid esters into carboxylic acid salts (saponification) and the creation of these hydrophilic groups likely resulted in increased wettability. XPS analysis also confirmed the formation of COO- in the KOH-treated spores.


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17 There were four major changes that occurred during H3PO4 treatment. i) The treatment helped to remove all residual proteins and peptides, resulting in a decrease in nitrogen content in the elemental analysis, a clear lane in the SDS-PAGE, and an absence of peaks in the MALDI-TOF MS analysis. ii) H3PO4 treatment hydrolyzed esters into carboxylic acids, resulting in the disappearance of ester (C=O) vibration at 1744 cm-1. Furthermore, H3PO4 protonated COO- to form carboxylic acids, thus COO- vibration at 1583 cm-1 disappeared and carboxylic acid (C=O) vibration at 1710 cm-1 increased. The formation of carboxylic acids supports the inversion of zeta potential at higher pH reported by Binks et al.13 iii) H3PO4 treatment removed cellulose, hence cellulose-associated vibrations at 1377, 1344, 1260, 1140, 1110, 1051, and 999 cm-1 disappeared after acidolysis. The removal of cellulose was also reflected by a drastic decrease in O–H vibration intensity at 3360 cm-1. A characteristic thermal decomposition related to cellulose around 355 ˚C was also absent in the TGA analysis of H3PO4-treated spores. TEM of the H3PO4-treated spore showed a lack of intine wall that is primarily made of cellulose. Cellulose removal is not surprising because 85% H3PO4 has been shown to dissolve cellulose.52 iv) H3PO4-treated spores (treatment temperature: 160 ˚C) became hydrophobic, formed a liquid marble, and failed to disperse in an aqueous medium (Figure 7), which can be explained due to lack of COO- and –OH functional groups. Using this new knowledge of how each treatment step alters the chemistry of the lycopodium spores, we were able to tailor the treatment steps to produce spores that were not only devoid of proteins but were also readily dispersible in water. Our study revealed that spores after KOH treatment were hydrophilic, but this property was reversed after H3PO4 treatment. We were able to use FTIR and TGA to map this property-shift to the reduction in carbohydrate content and the associated reduction in –OH groups. This reasoning allowed us to postulate that a lower H3PO4 treatment temperature (60 ˚C) could reduce carbohydrate dissolution, which in turn could increase –OH functional groups and thus increase hydrophilicity of the spores. Indeed, spores treated with H3PO4 at 60 ˚C were readily dispersible. This illustrates the value of careful characterization of spores with a suite of complementary techniques. Phenolic components (phenylpropanoids) specifically ferulic acid and p-coumaric acid are the building block of sporopollenin.53 These components are responsible for the UV-B radiation absorption54-55 and intrinsic autofluorescence of the sporopollenin wall.56 The fact that the aromatic ring vibrations (1654, 1606, 1500, 1448, and 1164 cm-1) and the exine wall autofluorescence remained despite undergoing the full treatment suggests that these phenolic components were not removed after chemical treatment. For oral therapeutics delivery, sporopollenin microcapsule may hold great potential to protect light-sensitive encapsulated compounds from photo-oxidation and thereby increase their shelf life. Moreover, phenylpropanoids have also shown anti-inflammatory properties,57 which may have a beneficial effect during oral vaccination.

5. Conclusion Microcapsules derived from natural lycopodium spores via acetone-KOH-H3PO4 treatment have shown promising prospects for delivering delicate, acid- and enzyme-labile therapeutics orally. This study addressed two fundamentally important areas of lycopodium chemistry: the fate of native proteins and spore wettability following different chemical treatments. To begin, natural lycopodium spores were hydrophobic and contained low molecular weight (~10 kD) proteins. Acetone treatment although partially removed phospholipids, the acetone-treated spores retained their intrinsic proteins and hydrophobicity. KOH treatment led to significant removal of proteinaceous materials and hydrolyzed the esters into carboxylic acid salts, thereby rendering the spores hydrophilic. H3PO4 treatment removed the trace amount of



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18 residual peptides that were left after KOH treatment, converted COO- to carboxylic acids, and solubilized carbohydrates from the spore wall. Temperature during H3PO4 treatment regulated the degree of carbohydrate dissolution, which subsequently affected spore hydrophilicity and wettability. Reducing H3PO4 treatment temperature from 160 ˚C to 60 ˚C significantly improved the hydrophilicity of the treated spore. Insights gained from this study helped us to understand the chemical changes lycopodium spores undergo during each chemical treatment step. This knowledge helped us to devise a treatment protocol to improve spore hydrophilicity while maintaining a protein-free hollow structure.

Supporting Information Confocal laser scanning microscopy (CLSM) images of lycopodium spores, SDS-PAGE analysis of intact lycopodium spores, MALDI-TOF MS analysis of lycopodium spore proteins, FTIR analysis of lycopodium spore extracts removed by KOH hydrolysis, XPS analysis of lycopodium spore surface, a list of FTIR vibrations from Lycopodium clavatum spores, diffuse reflectance infrared Fourier transform spectroscopy (DRIFTS) spectra of lycopodium spores.

Acknowledgments The authors thank Rajeev Rajbhandari and Erandi Rajakaruna in the Department of Plant and Soil Science at Texas Tech University for their help in TGA and FTIR. The authors also thank Mary Catherine Hastert in the College of Arts & Sciences Microscopy of Texas Tech University for her help in TEM imaging. The authors acknowledge the help from Dr. Masoud Zabet-Moghaddam in the Center for Biotechnology and Genomics at Texas Tech University for his help in MALDI-TOF MS. This research was supported by the National Institutes of Health (NIH) [grant number DP2HD075691] and the Defense Advanced Research Projects Agency (DARPA) [grant number N66001-12-1-4251].

Conflict of Interest HSG is a co-inventor on a patent related to the development of pollen grains for oral vaccines. This potential conflict of interest has been disclosed and is managed by Texas Tech University.

References (1) Mackenzie, G.; Boa, A. N.; Diego-Taboada, A.; Atkin, S. L.; Sathyapalan, T. Sporopollenin, the Least Known Yet Toughest Natural Biopolymer. Front. Mater. 2015, 2, 66. (2) Domínguez, E.; Mercado, J. A.; Quesada, M. A.; Heredia, A. Pollen Sporopollenin: Degradation and Structural Elucidation. Sex. Plant Reprod. 1999, 12, 171-178. (3) Li, F.-S.; Phyo, P.; Jacobowitz, J.; Hong, M.; Weng, J.-K. The Molecular Structure of Plant Sporopollenin. Nat. Plants 2019, 5, 41-46. (4) Katifori, E.; Alben, S.; Cerda, E.; Nelson, D. R.; Dumais, J. Foldable Structures and the Natural Design of Pollen Grains. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 7635-7639. (5) Jardine, P. E.; Fraser, W. T.; Lomax, B. H.; Gosling, W. D. The Impact of Oxidation on Spore and Pollen Chemistry. J. Micropalaeontol. 2015, 34, 139-149.


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20 (21) Jia, P.-F.; Li, H.-J.; Yang, W.-C. Transmission Electron Microscopy (TEM) to Study Histology of Pollen and Pollen Tubes. In Plant germline development: methods and protocols; Schmidt, A., Ed.; Springer New York: New York, NY, 2017; pp 181-189. (22) Uddin, M. J.; Gill, H. S. Ragweed Pollen as an Oral Vaccine Delivery System: Mechanistic Insights. J. Control. Release 2017, 268, 416-426. (23) Tryon, A. F.; Lugardon, B. Spores of the Pteridophyta, Springer, New York, NY: 1991. (24) Candiano, G.; Bruschi, M.; Musante, L.; Santucci, L.; Ghiggeri, G. M.; Carnemolla, B.; Orecchia, P.; Zardi, L.; Righetti, P. G. Blue Silver: A Very Sensitive Colloidal Coomassie G-250 Staining for Proteome Analysis. Electrophoresis 2004, 25, 1327-1333. (25) Clark, A. E.; Kaleta, E. J.; Arora, A.; Wolk, D. M. Matrix-Assisted Laser Desorption Ionization–Time of Flight Mass Spectrometry: A Fundamental Shift in the Routine Practice of Clinical Microbiology. Clin. Microbiol. Rev. 2013, 26, 547-603. (26) Stuart, B. H. Infrared Spectroscopy: Fundamentals and Applications. Ando, D. J., Ed.; John Wiley and Sons: New York, 2005; pp 137-165. (27) Bağcıoğlu, M.; Kohler, A.; Seifert, S.; Kneipp, J.; Zimmermann, B. Monitoring of Plant– Environment Interactions by High-Throughput FTIR Spectroscopy of Pollen. Methods Ecol. Evol. 2017, 8, 870-880. (28) Pappas, C. S.; Tarantilis, P. A.; Harizanis, P. C.; Polissiou, M. G. New Method for Pollen Identification by FT-IR Spectroscopy. Appl. Spectrosc. 2003, 57, 23-27. (29) Max, J.-J.; Chapados, C. Infrared Spectroscopy of Aqueous Carboxylic Acids: Comparison Between Different Acids and Their Salts. J. Phys. Chem. A 2004, 108, 3324-3337. (30) Gottardini, E.; Rossi, S.; Cristofolini, F.; Benedetti, L. Use of Fourier Transform Infrared (FT-IR) Spectroscopy as a Tool for Pollen Identification. Aerobiologia 2007, 23, 211-219. (31) Fraser, W. T.; Scott, A. C.; Forbes, A. E. S.; Glasspool, I. J.; Plotnick, R. E.; Kenig, F.; Lomax, B. H. Evolutionary Stasis of Sporopollenin Biochemistry Revealed by Unaltered Pennsylvanian Spores. New Phytol. 2012, 196, 397-401. (32) Julier, A. C. M.; Jardine, P. E.; Coe, A. L.; Gosling, W. D.; Lomax, B. H.; Fraser, W. T. Chemotaxonomy as a Tool for Interpreting the Cryptic Diversity of Poaceae Pollen. Rev. Palaeobot. Palynol. 2016, 235, 140-147. (33) Domínguez, E.; Mercado, J. A.; Quesada, M. A.; Heredia, A. Isolation of Intact Pollen Exine Using Anhydrous Hydrogen Fluoride. Grana 1998, 37, 93-96. (34) Bagcioglu, M.; Zimmermann, B.; Kohler, A. A Multiscale Vibrational Spectroscopic Approach for Identification and Biochemical Characterization of Pollen. PLoS ONE 2015, 10, e0137899. (35) Zimmermann, B.; Kohler, A. Infrared Spectroscopy of Pollen Identifies Plant Species and Genus As Well As Environmental Conditions. PLoS ONE 2014, 9, e95417. (36) Rouxhet, P. G.; Genet, M. J. XPS Analysis of Bio-Organic Systems. Surf. Interface Anal. 2011, 43, 1453-1470. (37) Ahlers, F.; Bubert, H.; Steuernage, S.; Wiermann, R. The Nature of Oxygen in Sporopollenin From the Pollen of Typha angustifolia L. znc 2000, 55, 129-136. (38) Gammon, W. J.; Kraft, O.; Reilly, A. C.; Holloway, B. C. Experimental Comparison of N(1s) X-Ray Photoelectron Spectroscopy Binding Energies of Hard and Elastic Amorphous Carbon Nitride Films with Reference Organic Compounds. Carbon 2003, 41, 1917-1923.


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21 (39) Kondo, T.; Casolo, S.; Suzuki, T.; Shikano, T.; Sakurai, M.; Harada, Y.; Saito, M.; Oshima, M.; Trioni, M. I.; Tantardini, G. F.; Nakamura, J. Atomic-Scale Characterization of NitrogenDoped Graphite: Effects of Dopant Nitrogen on the Local Electronic Structure of the Surrounding Carbon Atoms. Phys. Rev. B 2012, 86, 035436. (40) Wu, N.; Fu, L.; Su, M.; Aslam, M.; Wong, K. C.; Dravid, V. P. Interaction of Fatty Acid Monolayers with Cobalt Nanoparticles. Nano Lett. 2004, 4, 383-386. (41) Moulder, J. F.; Stickle, W. F.; Sobol, P. E.; Bomben, K. D. Handbook of X-Ray Photoelectron Spectroscopy, Physical Electronics USA, Inc.: 1995. (42) Aussillous, P.; Quéré, D. Liquid Marbles. Nature 2001, 411, 924–927. (43) Meng, X.; Pan, Q.; Ding, Y.; Jiang, L. Rapid Determination of Phospholipid Content of Vegetable Oils by FTIR Spectroscopy Combined with Partial Least-Square Regression. Food Chem. 2014, 147, 272-278. (44) Naktiyok, J.; Bayrakçeken, H.; Özer, A. K.; Gülaboğlu, M. Ş. Kinetics of Thermal Decomposition of Phospholipids Obtained From Phosphate Rock. Fuel Process. Technol. 2013, 116, 158-164. (45) Gonzalez Cruz, P.; Uddin, M. J.; Atwe, S. U.; Abidi, N.; Gill, H. S. Chemical Treatment Method for Obtaining Clean and Intact Pollen Shells of Different Species. ACS Biomater. Sci. Eng. 2018, 4, 2319-2329. (46) Yang, H.; Yan, R.; Chen, H.; Lee, D. H.; Zheng, C. Characteristics of Hemicellulose, Cellulose and Lignin Pyrolysis. Fuel 2007, 86, 1781-1788. (47) Diego-Taboada, A.; Cousson, P.; Raynaud, E.; Huang, Y.; Lorch, M.; Binks, B. P.; Queneau, Y.; Boa, A. N.; Atkin, S. L.; Beckett, S. T.; Mackenzie, G. Sequestration of Edible Oil From Emulsions Using New Single and Double Layered Microcapsules from Plant Spores. J. Mater. Chem. 2012, 22, 9767-9773. (48) Burhenne, L.; Messmer, J.; Aicher, T.; Laborie, M.-P. The Effect of the Biomass Components Lignin, Cellulose and Hemicellulose on TGA and Fixed Bed Pyrolysis. J. Anal. Appl. Pyrolysis 2013, 101, 177-184. (49) Mojet, B. L.; Ebbesen, S. D.; Lefferts, L. Light at the Interface: The Potential of Attenuated Total Reflection Infrared Spectroscopy for Understanding Heterogeneous Catalysis in Water. Chem. Soc. Rev. 2010, 39, 4643-4655. (50) Wilfong, W. C.; Srikanth, C. S.; Chuang, S. S. C. In Situ ATR and DRIFTS Studies of the Nature of Adsorbed CO2 on Tetraethylenepentamine Films. ACS Appl. Mater. Interfaces 2014, 6, 13617-13626. (51) Dell’Anna, R.; Lazzeri, P.; Frisanco, M.; Monti, F.; Malvezzi Campeggi, F.; Gottardini, E.; Bersani, M. Pollen Discrimination and Classification by Fourier Transform Infrared (FT-IR) Microspectroscopy and Machine Learning. Anal. Bioanal. Chem. 2009, 394, 1443-1452. (52) Zhang, Y. H. P.; Cui, J.; Lynd, L. R.; Kuang, L. R. A Transition From Cellulose Swelling to Cellulose Dissolution by O-Phosphoric Acid: Evidence From Enzymatic Hydrolysis and Supramolecular Structure. Biomacromolecules 2006, 7, 644-648. (53) Fraser, W. T.; Lomax, B. H.; Jardine, P. E.; Gosling, W. D.; Sephton, M. A. Pollen and Spores as a Passive Monitor of Ultraviolet Radiation. Front. Ecol. Evol. 2014, 2. (54) Blokker, P.; Boelen, P.; Broekman, R.; Rozema, J. The Occurrence of P-Coumaric Acid and Ferulic Acid in Fossil Plant Materials and Their Use as UV-Proxy. Plant Ecol. 2006, 182, 197-207.



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Page 23 of 33 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


For Table of Contents Only

Acetone treatment KOH treatment H3PO4 treatment

Natural (Hydrophobic)

(Hydrophilic)

Acetone-treated (Hydrophobic)

Lipids Proteins

KOH-treated (Hydrophilic) (Hydrophobic) H3PO4-treated


ACS Applied Materials & Interfaces 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

Page 24 of 33

Figure 1 A

C

E

G

B

D

F

H


Page 25 of 33 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


Figure 2

A

B

ex

resin CM

C

int

D ex



Figure 3

kDa

1

2

3

4

5

250 – 150 – 100 – 75 – 50 – 37 – 25 – 20 – 15 – 10 –


Page 26 of 33

2243.5

2611.7

666.1

1388.9 1503.9 1688 1858.2

2785.8

498.4

Acetone-treated

3100

1388.9

Figure 4

Natural

2611.7 2785.8 3100

498.4

568.2

682.1 909.1

KOH-treated

441.1

H3PO4-treated

861.1

656.1

568.2 441.1

-CHCA

622.1 763.5 1144.7

Intensity (a. u.)

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


1503.9 1688.1 1859.2

Page 27 of 33

1000

2000

3000

4000

Mass (m/z)


5000


814

Natural

Acetone-treated

KOH-treated

H3PO4-treated

3600

3200

2800

2400

2000

1600 -1

Wavenumber (cm )


1200

800

721

999

1744 1710 1654 1606 1583 1517 1440 1377 1344 1280 1260 1140 1110

3007 2924 2854

3300

Figure 5

Absorbance

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

Page 28 of 33

Page 29 of 33

Figure 6 O 1s

N 1s

C-OH C-O/ O-H

Natural

C 1s C-C

C-N C-N / C-OH

C=O

C=O C=O

Acetonetreated

KOHtreated

Intensity (a.u.)

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


COO-

C-N=C

H3PO4treated

536 534 532 530 528 404 402 400 398 396 394 291 289 287 285 283 281

Binding Energy (eV)



Page 30 of 33

Figure 7

Natural

Acetone-treated

KOH-treated


H3PO4-treated

Page 31 of 33

Absorbance

723

1239 1163 1097

1377

1464

1744 1711

A

3006 2923 2853

Figure 8

1.5 Extract after acetone treatment

1.0

0.5

0.0 3600

3200

2800

2400

2000

1600

1200

800

-1

Wavenumber (cm )

B

C 100

Derivative weight (% / min)

Residual weight (%)

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


80 60 40 Extract after acetone treatment

20 0 0

100

200

300

400

500

600

0 -3 -6

213

Extract after acetone treatment

-9

390

-12 0

100

200

300

400

Temperature (°C)

Temperature (°C)


500

600


A 100 80 60

Natural Acetone-treated KOH-treated H3PO4-treated

40 20 0 0

100

200

300

400

500

600

Temperature (°C)

B

I

II

41

199

Region III IV

V

300

Natural


Figure 9

Residual weight (%)

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

Page 32 of 33

45

352

400

229 435

Acetone-treated

369

45

KOH-treated

343

441

37

H3PO4-treated 0

100

200

300

424

400

Temperature (°C)

500


600

Page 33 of 33

1200

974

1170

1600

1076

1583

A

1705

3360

Figure 10

Day 3 60 C

Absorbance

Day 3 160 C

Day 7 60 C

3600

3200

2800

2400

2000

Wavenumber (cm-1)

Residual weight (%)

B

100

1

80

-1

60

-3 351

40 20

-5 -7

Day 7 60 C

453

0 0

100

200

300

400

500

-9

C

Water


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


600

Temperature (°C)


Canola oil

800

hydrophobicity of

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