Synthetic polymers such as polyethylene, polyvinylchloride or polystyrene have excellent material properties that can be used in many applications. However, since they are not biodegradable, they accumulate in landfills. If not disposed properly, they may end up in the oceans, where they persist [1
]. Plastics degrade further to microplastics that can harm marine organisms [2
]. Polymer durability is usually a desired property and it is the actual littering which causes the main problem. Therefore, there is a need for the development of sustainable polymers and materials that can be used as barriers, coatings or films. The new products should biodegrade in nature, or, more preferably, be compostable in controlled conditions.
Cellulose fibers, which are widely used for papermaking, paperboard, cardboard, textiles and specialty chemicals, can be recycled after use and utilized for several cycles [3
]. Before combusting or composting, the fibers can be utilized for ethanol production, nanocellulose applications or biobased chemicals according the cascading principle. Developed countries have significantly reduced the use of paper in printing but fibers are increasingly used to make packaging materials. Wood is also a preferable source for textile fibers since trees can be grown without the use of large amounts of water, fertilizers and pesticides, which are needed in cotton production. Cotton production demands large land areas that could be used for food production instead [4
]. In 2013–2014, cotton was harvested on ca. 32,430,000 hectares corresponding to 2.3% of the world’s arable land area [5
]. Novel methods have been developed in the production of textiles from kraft and dissolving pulp [6
]. Also, ionic liquids have been used [7
Textiles and packaging materials from natural biodegradable materials need functional properties such as water repellency. Paper and board as such will disintegrate gradually if wet. To prevent this, cardboard is traditionally laminated with polyethylene films. As an alternative, the lignocellulosic surfaces can be treated with natural substances derived from bark which form structures that repel water [8
Birch wood (Betula pendula
Roth. and Betula pubescens
Ehrh.) is a valuable raw material for the furniture industry and veneer manufacturing in the Nordic countries. In 2017, the Finnish forest industry alone consumed approximately 14.3 million m3
hardwood logs, mainly birch [10
]. A volume of 2.7 m3
birch roundwood contains on average 28.6 kg oven dry outer bark (10.6 kg o.d. m−3
]. Based on these figures it can be estimated that the amount of oven dry outer bark produced is over 150,000 tonnes annually in Finland. Currently, it is solely used for producing bioenergy.
Birch outer bark is rich in valuable biochemicals like the naturally occurring biopolyester suberin and the triterpenoids betulinol and lupeol [12
]. The content of betulinol and lupeol in outer bark varies between 30%–35% and the content of suberin can be up to 40%–50% [13
]. Birch bark containing both inner and outer bark from a pulp mill has been reported to contain 5.9% of suberin [18
]. Suberin is a complex polymer, built by long-chain bifunctional ω-hydroxyacids and α,ω-diacids that are both C18 midchain modified and saturated, long-chain monofunctional fatty acids, and fatty alcohols which are interlinked by ester-bonds to glycerol. When suberin is depolymerized, the main components are long-chain aliphatic acids, typically 80%–90% of depolymerisates [19
]. Suberin fatty acids are covalently linked through esterification to ferulic acid and neighboring lignin-like polyaromatics [12
]. Suberin is believed to form partly orderly lamellar structures [19
Due to its complex structure, suberin extraction requires special conditions. It has been extracted as fatty acid salts by alkaline alcohol reagents, such as ethanol with sodium hydroxide [11
]. Ionic liquids such as 1-ethyl-3-methylimidazolium hexanoate, cholinium hexanoate, cholinium octanoate, and cholinium decanoate have been applied on extraction of suberin from cork [26
]. The extraction yield of suberin varied between 30.6% and 67.2%. The maximum yield of suberin with alkaline methanolysis has been reported to be approximately 55% [20
]. Due to the problems related to the recovery of ionic liquid, we focused on conventional alkaline alcoholic extraction of suberin fatty acids (SFAs) because alcohols are relatively easy to recover due to their high vapor pressure.
Our goal is to develop a resource-efficient method to extract suberin fatty acids from birch bark and investigate the possibility of using them as hydrophobic coatings for lignocellulosic fibers. We are aiming to replace fossil raw materials with completely sustainable raw materials to create water-repellent and functional surface coatings for cellulose-based packaging materials and textiles.
3. Materials and Methods
Silver birch (Betula pendula Roth.) outer bark was manually removed from freshly cut stems (diameter of the trees was 200–300 mm) and air dried. Outer bark was then ground using a cutting mill with a sieve cassette having 4 × 4 mm2 square openings. The ground outer bark was then freeze-dried and stored in an airtight polyethylene bag.
Unbleached softwood kraft pulp was obtained from a pulp mill after the blow line. The pulp was washed and screened in a laboratory using a Somerville screen (0.15 mm) to remove shives. The pulp was dewatered after screening and stored in a freezer at −20 °C. No pulp refining (beating) was carried out to obtain as untreated laboratory sheet structure as possible.
Isopropyl alcohol 99.8% (Merck KGaA, Darmstadt, Germany), sodium hydroxide 99.0% (Merck KGaA, Darmstadt, Germany), ethanol 94.0% (Altia Oyj, Rajamäki, Finland) and sulfuric acid 95% (VWR International S.A.S., Briare, France) were used in the experiments. Then, 2 molar sulfuric acid was prepared by pouring 112 mL concentrated sulfuric acid into 600 mL of maxima ultrapure water. The final volume was adjusted with ultra-pure water to 1000 mL after the solution was cooled to room temperature.
3.1. Suberin Fatty Acid Extraction and Isolation
The extraction procedure of suberin fatty acids is shown in Figure 6
. The extraction process was adapted from [11
] but the extraction time was prolonged to three hours.
A total of 100 g oven dry (o.d.) ground bark was placed in a 2000 mL round bottom flask. A volume of 900 mL isopropyl alcohol and 100 mL deionized water was mixed. Sodium hydroxide (20 g) was dissolved in the solution and it was poured in the flask. A reflux condenser was attached and the outer bark was refluxed for three hours. The extract was filtered hot and the bark residue was washed with 500 mL hot isopropyl alcohol and water mixture (9:1 v v−1) and filtered again. The two filtrates were combined and cooled to room temperature. The isopropyl alcohol was evaporated under vacuum and 1500 mL hot deionized water was then added. The water-insoluble betulinol fraction was precipitated out and filtered. The solid fraction was washed with 1000 mL hot deionized water and filtered. Again, the two filtrates were combined. The pH value of the combined filtrate containing suberin fatty acid soaps was adjusted to 4.7 using 2 M sulfuric acid. The suberin fatty acids soaps were converted back to fatty acids and the suberin fatty acid fraction was precipitated out from the solution. The suberin fatty acid fraction was washed with 1000 mL deionized water. Both the betulinol and suberin fatty acid fractions were freeze dried and weighed.
3.2. Elemental Analysis
Elemental analysis for betulinol and suberin fatty acid fractions was carried out using a closed wet HNO3
digestion method (Miller 1998 [32
]) in a microwave oven (CEM MDS 2000) and the extract was analyzed by an iCAP 6500 DUO inductively coupled plasma (ICP)-emission spectrometer (Thermo Fisher Scientific, Cambridge, UK).
3.3. GC and GC-MS Analysis
The composition of both the betulinol and suberin fatty acids fractions were quantified by gas chromatography flame ionization detection (GC-FID) and the peak identities were confirmed by gas chromatography mass spectrometry (GC-MS). The retention time and mass spectra of birch bark suberin and other low molecular weight components have previously been identified as carboxylic acid methyl esters and silylated alcohols [12
]. In this analysis the fractions were analyzed as their trimethylsilyl (TMS) esters and ethers. Cholesterol was used as standard. The fractions were silylated with a mixture of N
-Bis(trimethylsilyl)trifluoroacetamide (BSTFA): Chlorotrimethylsilane (TMCS): Pyridine (120:20:20) and heated for 50 min at 70 °C. The composition of the fractions by component, their retention time and Kovats’ retention index can be found in Table 1
The samples were analyzed on a Shimadzu GC-2010 Plus GC (Shimadzu Corporation, Kyoto, Japan) with a flame ionization detector, equipped with an AOC-20i autosampler and a split/splitless injector. The column used was a ZB-1HT (Phenomenex, Torrance, CA, USA), 20 m, 0.18 mm i.d. and film thickness 0.18 µm, coated with 100% polydimethylsiloxane. Initial temperature was 80 °C (1 min), temperature gradient was 8 °C min−1 and final temperature was 360 °C (15 min). Injection temperature was 250 °C and detector temperature was 360 °C. Split injection (1 µL) with a ratio of 25:1 was employed. Carrier gas was hydrogen at 40 cm s−1 linear velocity.
The samples were also analyzed on a Shimadzu GCMS-QP2010 Plus GC-MS (Shimadzu Corporation, Kyoto, Japan) for component identification. Gas chromatographic conditions were as reported above except for using helium as carrier gas. Mass spectrometer parameters were as follows: interface and ion source temperatures, 345 °C and 230 °C, respectively; ionization mode, electron ionization (EI) with 70 eV; acquisition mass range, 35–800 m z−1. Identifications were based on a comparison of the GC retention times (Kovats’ index) and EI spectra with those in our own database.
3.4. Preparation of Laboratory Sheets (Lignocellulosic Fiber Network)
Laboratory sheets were prepared in a standard sheet former with 60 g m−2 target grammage. Wet sheets were pressed twice, first 5 min at 400 kPa, then 2 min at 400 kPa. The sheets were dried in conditioned air (23 °C temperature and 50% relative humidity) using drying plates. Dried sheets were cut to squares having a 0.02 m2 area and stored in aluminum foil.
3.5. Suberin Fatty Acid Impregnation and Curing
A curing agent solution containing 50 mg ml−1
suberin fatty acids and 50 mg ml−1
maleic anhydride in ethanol was prepared. Dry solids of 10, 20 and 30 g m−2
(2.0, 4.0 and 6.0 mL curing agent) were applied on the laboratory sheets and the laboratory sheets were placed on an aluminum foil. The dosages were selected on the basis of commercial polymer coating grammage of paperboard [33
]. The volume of ethanol in 10 g m−2
was so small that 2 mL of additional ethanol was used in order to distribute suberin fatty acids and maleic anhydride evenly in the laboratory sheets. The solvent was evaporated in an oven at 70 °C for 30 min and the laboratory sheets were removed from the aluminum foil. The temperature was then increased to 150 °C and the laboratory sheets were kept in the oven overnight. One reference set of laboratory sheets was only heat treated (0 g m−2
) and was composed only of pure fibers (pure laboratory sheet).
3.6. Paper Technical Properties
Thickness, grammage, tensile strength, tear strength, brightness, yellowness and air permeability were measured for the laboratory sheets according to the following ISO standards:
Thickness, density, grammage: ISO 534 Paper and board—Determination of thickness, density and specific volume.
Tensile strength: ISO 1924 Paper and board—Determination of tensile strength.
Tear strength: ISO 1974 Paper—Determination of tearing resistance.
Optical properties: ISO 2469 Paper, board and pulps—Measurement of diffuse radiance factor (diffuse reflectance factor) and ISO 2470-1 Paper, board and pulps—Measurement of diffuse blue reflectance factor—Part 1: Indoor daylight conditions (ISO brightness).
Air permeance: ISO 5636 Paper and board—Determination of air permeance and air resistance (medium range).
3.7. Water Vapor Transmission Rate
The water vapor transmission rate (WVTR) was determined, using the relative humidity (RH) gradient of 0%/54%. Fiber sheets were sealed on aluminum cups containing 43 g CaCl2 as a desiccant, with the top side of the laboratory sheet facing up towards the moist side. There was an air gap of 6 mm width between the salt and the wire side of the laboratory sheet. The cups were placed in a desiccator cabinet equipped with a fan to circulate the air above the samples at a speed of 0.15 m s−1. The cabinet was kept at constant temperature of 24 °C and the RH was maintained at 54% using saturated Mg(NO3)2 solution. The cups were weighed after 0, 115, 1120, 1340 and 1555 min. The temperature and the RH of the cabinet were measured using a Rotronic RH meter (Rotronic AG, Bassersdorf, Switzerland) before each weighing. The water vapor transmission rate was calculated from the linear regression of the slope of weight gain vs. time by dividing the slope by the treated laboratory sheet area. Three replicates of each paperboard type were tested. The thickness of the specimens was measured before testing at five points with a micrometer (Lorentzen & Wettre, Kista, Sweden, precision 1 µm).
3.8. Contact Angle Measurement
The contact angles of ultrapure water on the coated paperboards were measured using a CAM 200 Series Optical Contact Angle and Surface Tension meter (KSV Instruments Ltd.; Helsinki, Finland, now part of Biolin Scientific, Stockholm, Sweden). The drop size was set to 4 µl and three parallel measurements were performed for each substrate. The contact angle was calculated as an average of the right and left angles 1 s after the drop was detached. The measurement time was 60 s.
3.9. Scanning Electron Microscopy
Scanning electron microscopy (SEM) images were obtained by Zeiss GeminiSEM 450 field emission scanning electron microscope (Carl Zeiss Microscopy GmbH, Jena, Germany) equipped with secondary electron analyzer. Acceleration voltage was 0.500 kV and probe current 50 pA. Gold sputtering was applied on the samples prior to imaging.
3.10. Time-of-Flight Secondary-Ion Mass Spectrometry
Secondary ion mass spectra were obtained using a Physical Electronics ToF-SIMS TRIFT II spectrometer (Physical Electronics Inc., Chanhassen, MN, USA). A primary ion beam of 69 Ga+ liquid metal ion source (LIMS) with 25 kV accelerating voltage and 600 pA beam current (in DC mode) was used in both positive and negative modes. The measurements were done from an area 200 × 200 µm and the analysis depth is in the order of few nanometers. The measurement time of 5 min was used and the total ion dose was <1012 ions cm−2. Charge compensation was obtained with an electron flood gun pulsed out of phase with respect to the ion gun.