Lipophilic extractives naturally occurring in wood tend to coalesce during pulping to form pitch deposits, which have particularly undesirable effects on the pulping process and quality of pulp produced. A chemical characterization of different eucalypt species [Eucalyptus nitens, E. grandis, and E. dunnii (of different site qualities)] wood and generated pulp was performed. This study aimed at determining the effects of wood storage at -20 °C (for 6 months), by examining their chemical composition and indigenous microflora. Fatty acids were the main lipophilic compounds among E. dunnii (SQ3 and SQ4) and E. grandis wood extractives. The wood of E. nitens posed the least risk for pitch deposit formation, making it the most suitable Eucalyptus species for pulping. Storage of wood chips at -20 °C had a similar effect as the traditional method of seasoning (storage of wood outdoors prior to pulping) used for the reduction of lipophilic extractives. A 25 to 44% reduction of total extractives was observed in the raw material after storage. Variations in bacterial and fungal communities were observed after storage, and should be taken into consideration when conducting lab scale trials. If storage of wood chips is necessary for lab testing, it should be retained for a maximum of 3 months at -20 °C.
The Effects of Wood Storage on the Chemical Composition and Indigenous Microflora of Eucalyptus species Used in the Pulping Industry
Lucretia Ramnath,a Bruce Sithole,b,c and Roshini Govinden a*
Lipophilic extractives naturally occurring in wood tend to coalesce during pulping to form pitch deposits, which have particularly undesirable effects on the pulping process and quality of pulp produced. A chemical characterization of different eucalypt species [Eucalyptus nitens, E. grandis, and E. dunnii (of different site qualities)] wood and generated pulp was performed. This study aimed at determining the effects of wood storage at -20 °C (for 6 months), by examining their chemical composition and indigenous microflora. Fatty acids were the main lipophilic compounds among E. dunnii (SQ3 and SQ4) and E. grandis wood extractives. The wood of E. nitens posed the least risk for pitch deposit formation, making it the most suitable Eucalyptusspecies for pulping. Storage of wood chips at -20 °C had a similar effect as the traditional method of seasoning (storage of wood outdoors prior to pulping) used for the reduction of lipophilic extractives. A 25 to 44% reduction of total extractives was observed in the raw material after storage. Variations in bacterial and fungal communities were observed after storage, and should be taken into consideration when conducting lab scale trials. If storage of wood chips is necessary for lab testing, it should be retained for a maximum of 3 months at -20 °C.
Keywords: Lipophilic extractives; Eucalyptus; Storage; Microflora
Contact information: a: Discipline of Microbiology, School of Life Sciences, College of Agriculture, Engineering and Science, University of KwaZulu-Natal, Westville Campus, P/Bag X54001, Durban 4000, South Africa; b: Biorefinery Industry Development Facility, Council for Scientific and Industrial Research, Durban 4000, South Africa; c: Discipline of Chemical Engineering, University of KwaZulu-Natal, Durban 4000, South Africa; *Corresponding author: firstname.lastname@example.org
Eucalyptus species are the most extensively planted hardwood species in the world and are primarily used in the pulp and paper industry (Sappi 2012). Extractives make up the minor fraction of hardwoods but have the greatest potential to cause problems during pulping (Leskinen et al. 2015). The extractive content of wood is highly variable amongst species (Moodley 2011; Yang and Jaakkola 2011); therefore careful selection of Eucalyptus species for pulping is required and currently not taken into consideration in commercial pulping operations (Swain and Gardner 2003). The lipophilic fraction of wood extractives is responsible for the formation of pitch deposits during pulp production (Sjöström 1993; Kontkanen et al. 2004; Wang and Jiang 2006), resulting in poor pulp quality and gumming up of machinery (Hillis and Sumimoto 1989; Patrick 2004; Sarja 2007). Traditionally, pitch deposits are reduced by seasoning logs and wood chips (outdoor storage of wood prior to processing) (Allen et al. 1991; Back and Allen 2000). Storage reduces the extractive content by hydrolytic or oxidative conversion by plant enzymes and the activity of indigenous microflora. The reduction of extractive content occurs much faster in woodchips rather than log form, as the oxidation processes proceed more easily and rapidly (Gutiérrez et al. 1998; Burnes et al. 2000). Temperature is also known to have an effect on the degradation of lipophilic compounds (Silvério et al. 2008). It has been reported that during winter the hydrolysis of wood esters, which forms part of the pitch, is drastically reduced compared to the summer months (Almeida and Silva 2001). In industry, seasoning wood at ambient temperatures usually significantly reduces the lipophilic content in the wood. However, care has to be taken, as prolonged seasoning could lead to uncontrolled action of microorganisms and insects (Bajpai 1999; Gutiérrez et al. 2001). The activity of such organisms could lead to cellulose and hemicellulose loss in the wood, ultimately resulting in poor pulp quality (Burnes 2000). Similarly, care has to be taken when storing wood for scientific investigations. Under laboratory conditions, wood chips are usually stored at -20 °C to prohibit any natural degradation of the lipophilic extractives, contamination by foreign organisms and to preserve the original state of the wood (Venn 1980; Petäistö 2006).
The lipophilic extractives of E. globulus and certain hybrids have been extensively characterized (Gutiérrez et al. 1999; Gutiérrez et al. 2001; Freire et al. 2002, 2006; Prinsen et al. 2012); however, there is a lack of information on extractives from E. grandis, E. urograndis (Freire et al. 2006), E. urophylla, E. camaldulensis (Silvério et al. 2007), E. nitens, and E. dunnii (Rencoret et al. 2007). In addition, limited information on the effect of site quality (soil quality and environmental conditions at the plantation site) on extractive content of Eucalyptus is available. Incorporating knowledge on the lipophilic content of different Eucalyptus species when selecting wood for pulping will go a long way towards reducing the overall lipophilic content of pulp, subsequently reducing the risks of pitch formation. In the present study, the chemical composition of the wood and pulp of four Eucalyptus species, E. grandis, E. nitens, E. dunnii site quality 3 (SQ3) and E. dunnii SQ4 were evaluated. Furthermore, the effects of storage at -20 °C on the chemical composition of different Eucalyptus spp. and their indigenous microflora were investigated.
E. grandis, E. nitens, E. dunnii (SQ3), and E. dunnii (SQ4) were obtained from a commercial pulping mill in Umkomaas located on the south coast of South Africa. Wood chips from individual wood species were collected as they were being chipped in the wood yard. Samples were transported back to the laboratory and stored at -20 °C. Chip samples of the different wood species were milled using the Brabender® Wiley-mill to coarse sawdust. The sawdust was then passed through a 0.40 mm (40 mesh) screen and used for chemical analyses (TAPPI 2012). The following analyses were conducted on the milled wood chips before and after storage at -20 °C for 6 months. Time taken to mill and analyze the wood chips was up to 5 days. Wood chips were also pulped to determine any changes in the quality of the pulp.
Hot Water Extraction
Hot water (water soluble) extractives (waxes, fats, some resins, photosterols, and non-volatile hydrocarbons, low molecular weight carbohydrates, salts, and other water soluble substances) were evaluated by weighing out five grams of sawdust (moisture content of original sample recorded) in a 400 mL conical flask and slowly adding 100 mL of hot deionised water. The contents of the conical flask were well stirred to prevent the sawdust from floating. The conical flask was placed on a pre-heated hot plate and left to stand for 3 h. The initial level of the contents was noted and kept constant by the addition of hot deionised water. A pre-weighed number 3 crucible (Pyrex) and a vacuum pump were used to filter the sawdust which was washed with 200 mL of hot deionised water and dried at 60 ºC overnight (TAPPI Test Method T204 om-88). Experiments were performed in triplicate.
The Soxhlet extraction method was used to evaluate the solvent (water insoluble) extractives in the wood samples. Four grams of hot water extracted sawdust (moisture content of wet sawdust recorded) were weighed into an extraction thimble, which was placed in a Soxhlet apparatus that was attached to a pre-weighed 500 mL round bottomed flask containing 300 mL toluene-ethanol (2:1). The heating mantle was adjusted such that the solvent cycled through the system six times over 4 h. The extractives were rotor evaporated to dryness, cooled in desiccator and weighed (TAPPI Test Method T204 om-88). Experiments performed in triplicate.
The other chemical characteristics of the wood chips were evaluated by conducting extractive analyses such as near infra-red reflectance analyser (NIRA) for the rapid quantification of wood chemical components and high performance liquid chromatography (HPLC) for quantification of cellulose and hemicelluloses (glucose, mannose, arabinose, xylose, rhamnose, and galactose) (TAPPI Test Methods 1996-1997; Wallis et al. 1996; Wright and Wallis 1996).
Pyrolysis GC-MS analysis
The lipophilic extractives present in the samples were ascertained by Py-GC/MS analysis. A multi-shot pyrolyzer, EGA/PY-3030 D, (Frontier Lab, Japan) attached to an ultra-alloy capillary column (30 m x 0.25 mm, 0.25 μm) was used for analysis. The samples were pyrolysed at 550 °C for 20 seconds and the interface temperature was set at 350 °C. The chromatographic separation of the volatile components released by pyrolysis was performed using an ultra-alloy column. The injection temperature was set to 280 °C and the column flow rate was set to 1.0 mL/min with helium as the carrier gas. The temperature was programmed as follows: 50 °C for 2 min; rate 3 °C/min up to 200 °C and held for a further 4 min. The ion source and interface temperatures were set to 200 °C and 300°C, respectively. The scan range used for the mass selective detector was 40 to 650 m/z. Pyrolysis products were identified by comparing their mass spectra with the mass spectrum NIST library attached to instrument (modified from Sithole and Watanabe 2013).
Pulping was performed in duplicate according to the acid bi-sulphite process currently implemented in Dissolving Wood Pulp (DWP) mills. Pulp quality was determined by assessing viscosity (TAPPI T203 om-94), final pulp yield, kappa number, acid insoluble lignin (TAPPI T222 om-88), alpha cellulose, S8, S10 (TAPPI T235-OM60) and hemicelluloses (Forestry and Forest Products Research Centre, CSIR, Personal Communication 2014).
DNA Isolation and Polymerase Chain Reaction (PCR)
DNA was extracted from 0.2 g milled chips using a kit as per manufacturer’s specifications (Soil DNA Extraction Kit, Zymo Research). Ribosomal genes were amplified from microbial genomic DNA using 16S and ITS2 region of 18S rRNA genes with the universal primer sets: 63F/1387R (Marchesi et al. 1998) and ITS5F/ITS4R (White et al. 1990), respectively. Each amplification reaction (50 μL) contained 1.25 mM MgCl2, 0.125 μM forward and reverse primers, 0.2 mM deoxynucleoside triphosphate (dNTPs), 0.25 U SuperTherm Taq DNA polymerase (Southern Cross Biotech), and 20 to 200 ng of template DNA. PCR was performed using the GeneAmp PCR System 9700 (Applied Biosystems). For amplification of 16S rRNA, PCR conditions were as follows: initial denaturation at 95 °C for 5 min, 30 cycles of denaturation, annealing and extension at 95 °C for 1 min, 55 °C for 1 min, 72 °C for 1.5 min and a final extension at 72 °C for 5 min, whereas 18S rRNA amplification conditions were: initial denaturation at 95 °C for 2 min, 25 cycles of 95 °C for 30 sec, 53 °C for 45 sec, 72 °C for 1 min and a final extension at 72 °C for 8 min. The amplicons were analyzed by electrophoresis on 1% agarose (SeaKem) gels in 1× TAE running buffer at 90 V for 45 min. After electrophoresis, the gels were stained in 0.5 μg/mL ethidium bromide and visualized using the Chemi-Genius 2 BioImaging System (Syngene). Upon confirmation of 16S and 18S amplicons, products were purified using a GeneJET™ PCR purification kit (Fermentas) and re-amplified in a touchdown thermal profile program using nested PCR and primers with GC clamps. PCR for 16S rRNA genes were initially performed using two primer sets: 338F-GC with a GC-clamp: 5´-CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGG-3´ and 518R 211 (237-bp fragment) (Handschur et al. 2005); 933F-GC with a GC-clamp: 5´-CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGG-3´ and 1387R (500-bp fragment) (Ji et al. 2004). The composition of the reaction mixtures was the same as that used for the first PCR. For amplification of 16S rRNA, PCR conditions were as follows: initial denaturation at 95 °C for 5 min, 30 cycles of denaturation annealing and extension at 95 °C for 1 min, 55 °C for 1 min, 72 °C for 1.5 min and a final extension at 72 °C for 5 min, whereas 18S rRNA amplification conditions were: initial denaturation at 95 °C for 2 min, 25 cycles of 95 °C for 30 sec, 53 °C for 45 sec, 72 °C for 1 min and a final extension at 72 °C for 8 min (Schabereiter-Gurtner et al. 2001).
Denaturing Gradient Gel Electrophoresis (DGGE)
DGGE was performed using the D-Code Universal Mutation Detection System (BioRad), and an optimized method (Ramnath et al. 2013) modified from Muyzer and Smalla (1998). PCR samples were loaded onto vertical perpendicular polyacrylamide gels (6% acrylamide:bisacrylamide [37.5:1]) in 1× TAE buffer prepared using 30 and 60% denaturant (100% denaturant corresponds to 7 M urea and 40% formamide). Gradients of 30% and 60% were optimal for bacterial amplicons (~237 bp) and 25% and 50% for fungal amplicons (~316 bp). A pre-run was performed at a constant voltage of 150 V at 60 °C for 30 min, following which, DGGE profiles were generated at a constant voltage of 60 V in 1× TAE buffer at 60 °C for 16 h. After electrophoresis, gels were stained in 0.5 μg/mL ethidium bromide for 60 min, destained in the same volume of 1× TAE buffer for 30 min and visualized using the Chemi-Genius 2 BioImaging System (Syngene).
RESULTS AND DISCUSSION
Hot water extraction is known to be an efficient technique for the removal of hemicellulose from wood chips prior to pulping in the production of dissolving pulp (Li et al. 2010). A comparison of the samples before and after storage at -20 °C revealed a general decrease in hot water and solvent extractives (Fig. 1).
Fig. 1. Wet chemistry of wood chips of different Eucalyptus spp. before and after storage at -20°C.
The highest hot water extractive content was observed for E. dunnii SQ3 (11.2%). The extractive contents are known to vary from tree to tree and considerably from species to species (Hillis 2014). E. dunnii SQ3 exhibited higher hot water extractive (11.2%) and solvent extractive (2%) content compared to SQ4 (6% and 1.7%, respectively), demonstrating the variation in chemical characteristics amongst Eucalyptus wood of the same species but from different site qualities. This is supported by Kilulya et al. (2014) who also observed that variations in lipophilic content are influenced by tree species and site qualities. Furthermore, their study validates our finding of higher amounts of total lipophilic extractives in E. dunnii compared to E. grandis. Wood chips with a hot-water soluble content exceeding 7 to 8% (as seen with E. dunnii and E. grandis in this study) result in obstruction of delignification due to competition of extractives with lignin for active cooking chemicals during pulping (Back and Allen, 2000). Consequently, higher amounts of active alkali are required which increases the cost and waste generated (Cohen and Mackney 1951). Extractives may cause liquor decomposition, also resulting in higher chemical requirements per ton of pulp produced (Hillis 2014).
High performance liquid chromatography (HPLC) was used for quantification of hemicelluloses (glucose, mannose, arabinose, xylose, rhamnose, and galactose) in the wood chips before and after storage. An increase in xylose content after storage for all Eucalyptus species was observed. This may be attributed to the natural degradation of glucuronoxylan during storage (Table 1). Pentosans abundant in hardwoods are mostly glucuronoxylan. The bonds between xylose units in glucuronoxylan are simple to hydrolyze (Testova et al. 2009), highlighting the potential for natural degradation during storage. Galactose and arabinose levels decreased for all Eucalyptusspp. after storage with the greatest decrease observed in E. nitens. This decrease during storage may be attributed to microbial degradation. Previous studies have shown that during the initial stages of fungal degradation, the carbohydrates most modified were sugars derived from the primary walls and middle lamella (galactose and arabinose). These are often recognized as the building blocks of hemicelluloses and/or arabinogalactan proteins in the primary wall (Skyba et al. 2013).
The mannose content of the wood material decreased by 24%, 21%, and 9% for E. dunnii (SQ3), E. grandis, and E. nitens, respectively, after storage. This decrease in mannose content may be caused by the activity of hemicellulolytic enzymes produced by microorganisms indigenous to the wood, particularly with E. dunnii (SQ3) and E. grandis. Wood treated with specific enzymes enable the extraction of high molecular weight materials, whilst non-specific enzymes were found to degrade hemicelluloses to a large extent (Azhar 2015). Thus, the increase in sugars observed in this study may be attributed to non-specific hemicellulolytic activity of indigenous microflora found in the wood material.
Table 1. Carbohydrate Content of Wood Chips Before and After Storage at -20°C
Py-GC-MS analyses provided insight into the lipophilic content of the samples. The py-GC-MS profiles of the different wood species in Fig. 2 show that the samples contained mainly lignin (3’,5’-dimethoxyacetophenone) and lipophilic (octadecanoic acid and hexadecanoic acid) degradation products due to pyrolysis. Storage at -20 °C had a discernible effect on the lipophilic content of the wood. The hexadecanoic and octadecanoic fatty acids observed are an indication of polymerised lipids. Polymerisation is the cause of incomplete removal of lipids by extraction (Tao et al. 2010). Reductions in these fatty acids were observed for E. dunnii (SQ4), E. nitens, and E. grandis. Such reductions would contribute to a decrease in pitch deposit formation and would usually be considered as a positive effect. These results confirm that storage at -20 °C resulted in degradation of pitch components as with aspen wood chips reported by Allen et al. (1991). However, if the wood chips are being stored for future experimental use, any changes from the original state of the wood would not be considered favourable. E. grandis (Fig. 1D) contained significant amounts of higher molecular weight lipophilic extractives (hexadecanoic and octadecanoic fatty acids), compared to the other wood species. These compounds were present in the other samples but at much lower levels, reflecting the inherently greater lipophilic content of E. grandis. Extractives were slightly lower than what has been reported for Eucalyptus species, which may indicate that samples were not as fresh and extractives may have transformed into higher molecular weight lipids (Kilulya et al. 2014). The phthalate and siloxane observed in E. dunnii (SQ3) are contaminants and are not part of the wood. Traces of phthalates and siloxane are frequently found in most industrial samples, and inevitably form part of the surface composition (Fardim and Durán 2003).
Fig. 2. Lipophilic profiles of different Eucalyptus species, before and after storage at -20 °C. (A) E. dunnii, (B) E. dunnii (FFP), (C) E. nitens, (D) E. grandis
The pulp produced from the wood chips was evaluated before and after storage, by measuring basic pulp chemistry: kappa number, viscosity, pulp yield, alpha cellulose, S10 and S18, and copper number. The kappa number provides an estimate of the amount of chemicals required during bleaching of wood pulp to achieve a pulp with a specified degree of whiteness. Therefore, the amount of bleach required is correlated to the lignin content of the pulp. Kappa number is used to determine the effectiveness of the lignin-extraction phase of the pulping process. It is approximately proportional to the residual lignin in the pulp (Costa and Colodette 2007). All Eucalyptus species demonstrated a reduction in the total lignin content of the pulp (Table 3). Greater removal of lignin may have been possible due to the opening up of the wood structure during storage by the enzymes produced by the indigenous microflora present. The pulp produced after storage had a lower kappa number than the starting material, but with a relatively low pulp viscosity (Table 2). Since the hemicelluloses are able to shield the cellulose chain from alkaline hydrolysis (Lindström and Teder 1995), the low pulp viscosities after storage are possibly due to removal of protective hemicelluloses (Azhar 2015). Pulp viscosity (degree of polymerization of cellulose fibers) reveals the relative consistency of the cellulose fibers after pulping. The pulp viscosity varied between the different Eucalyptus species from 28.01 to 131.7 cP. A general decrease in the viscosity was observed for all samples after 6 months of storage. Viscosity values for E. dunnii (SQ3) were very low and may be explained by the degradation of cellulose fibers by cellulase-producing microorganisms previously isolated from this particular wood species (Ramnath et al. 2013). A considerable decrease in viscosity of 41% and 66% in E. grandis and E. nitens pulp, respectively, was observed after storage. This may be attributed to endoglucanase activity, which is causes random cleavage of cellulose chains leading to lower degrees of polymerization and therefore lower viscosity. The retention of long cellulose fibers is essential for producing dissolving pulp (high grade cellulose pulp). Cellulase activity results in cellulose degradation; thus the negative effect of cellulases on pulp needs to be eliminated or minimized.
After storage, glucose levels varied in the range 80.85 to 86.19% and xylose from 5.27% to 6.44% (Table 3). This provides an approximate indication of the cellulose to hemicellulose ratio amongst the different species (Neiva et al. 2014). E. dunnii (SQ3) and E. dunnii (SQ4) displayed the lowest and highest ratio of glucose/xylose, respectively. These results support the notion that site quality does have an effect on the chemical composition of the wood material within individual species. The hemicellulose content in E. dunnii (SQ4) pulp was higher than the other Eucalyptus species examined. A general increase in xylose content (Table 3) was observed for all Eucalyptus pulp samples. Fišerová et al. (2013) found that xylose content of hydrolyzed beech and oak wood samples increased with increasing wood weight loss. Hence the increase in xylose observed may be attributed to wood weight loss during storage, which may be caused by wood-decaying fungi that became activated once thawed. Wood-decaying fungi have acquired a range of hydrolytic enzymes including cellulases, hemicellulases, and oxidative enzymes for the breakdown of lignocellulose into free monomers such as xylose, mannose, glucose, galactose, arabinose and rhamnose (Álvarez et al. 2016). In another study, greater xylose levels were observed when Eucalyptus and rice straw materials were treated with a culture supernatant of Trichoderma reesei than with traditional steam explosion as a pretreatment step (Álvarez et al. 2016). This emphasizes the xylan-hydrolyzing potential of T. reesei to release xylose oligomers and other soluble sugars (Jørgensen et al. 2007), and may explain the higher xylose concentrations documented in this study. In addition, xylanases may assist in the pulp beaching process, by removing xylan which allows for easier bleaching of the cellulose fibers, therefore consuming less bleach and energy (Garg et al. 2011). This partial degradation of xylan could also assist in the degradation of lignin by other microorganisms present, which is essential as lignin also impedes pulp bleaching (Subramaniyan and Prema 2002).
Table 2. Characteristics of the Pulp Produced from Wood Chips Before and After Storage
Table 3. Quantification of Lignin and Hemicelluloses of the Pulp Produced from Wood Chips Before and After Storage
Prior to storage, pulping of E. nitens wood resulted in the highest pulp yield of 50.8%; however, after storage the yield was reduced to 44.8%, the lowest level amongst the Eucalyptus spp. examined. After storage E. dunnii (SQ3) generated the highest pulp yield of 48.6%. Considering the pulping characteristics of the different species without the effect of storage, E. nitens would be the most suitable for pulping as it generated the greatest pulp yield (50.8%) and highest pulp viscosity (131.7%). Integrating the effects of storage and E. dunnii (SQ4) would be the most suitable for pulping as minimal effects on pulp yield and viscosity were observed for this species.
Following successful DNA isolation and PCR, DGGE analysis of the bacterial communities present in the wood revealed some interesting findings. There was a significant increase in the diversity and abundance of bacteria present in the wood samples after 6 months of storage (Fig. 3). The microbial community of harvested crops is known to transform during storage, and is particularly dependent on the water content of the samples (Barry Kay et al. 2009). Identities of the microorganisms were inferred using a molecular weight marker of known microbial species previously isolated from Eucalyptus wood chips. Bacillus thuringiensis and B. cereus appeared to be prevalent in all of the samples after storage, except for E. dunnii (SQ4). In fact, the bacterial community detected in E. dunnii (SQ4) post storage was minimal compared to the other Eucalyptus species. Following storage of E. grandis, DGGE analysis revealed the presence of Bacillus thuringiensis, Klebsiella sp., Pantoea ananatis, Micrococcus luteus, and Inquilinus limosus, and the subsequent loss of Prauserella and Saccharomonospora spp. E. nitens on the other hand, showed loss of the Inquilinus limosus population and increase in intensity of Lecleriasp., Saccharomonospora sp., and B. cereus populations.
Fungal diversity amongst the different Eucalyptus species was evident. For all Eucalyptusspecies, various fungal species observed prior to storage, increased in abundance after storage. This was observed for the following fungal species; Phialophora alba (E. dunnii SQ3 and E. grandis), Brachyalara straminea, Loddermayces elongisporus (E. dunnii SQ4), Aspergillus fumigatus (E. grandis), and Basidiomycota sp. (E. nitens). In addition, fungal species such as Paecilomyces variotii, Curvularia sp., and Pichia scolytii were discovered only after storage in E. dunnii (SQ3), E. dunnii (SQ4), and E. grandis, respectively. This may seem anomalous, as at temperatures of -10 to -12°C, microbial growth is considered to be minimal. However, some microorganisms remain inactivated during storage and then continue growing once thawed, leading to microbial degradation of the thawed product (Hui and Sherkat 2005). Freezer storage of material may eliminate the risk of microbial activity during the storage period, but wood material is still at risk during the cooling-down and thawing processes (Hansen 1990). Petäistö (2006) found that the growth of grey mold, such as Botrytis cinerea commonly found in soil, on Norway spruce progressed at the beginning and/or thawing phase of cold storage at -3°C. Temperatures above zero before and after the cold-storage phase may intensify the threat of grey mold damage although the optimum temperature for B. cinerea is approximately 20 °C (Petäistö 2006).
Low concentrations of extractives in wood have previously been associated with low resistance to decay, as extractives may function as a fungicide (Haupt et al. 2003; Latorraca et al. 2011; Severo et al. 2016). The increase in microbial populations and diversity may have been facilitated by the lower extractives content observed in the tested samples.
Fig. 2. DGGE profiles of bacterial (A) and fungal (B) communities present in different Eucalyptus species before and after storage at -20°C. Lanes: 1- time zero E. dunnii (SQ3); 2- after storage E. dunnii (SQ3); 3- time zero E. dunnii (SQ4); 4- after storage E. dunnii (SQ4); 5- time zero E. grandis; 6- after storage E. grandis; 7- time zero E. nitens; 8- after storage E. nitens
The microorganisms identified here have been described in other studies on hardwood chips, including Paecilomyces sp., Phialophora sp., Bacillus sp., Pseudomonas sp., and Micrococcus sp. (Adair et al. 2002; Rajala et al. 2010; Kropacz and Fojutowski 2014; Zhang et al. 2014; Szulc et al. 2017). The microorganisms detected in the wood chips of different Eucalyptus species have the potential to produce cellulolytic, hemicellulolytic, and xylanolytic enzymes (Schmidt 2006; Seo et al. 2013; Nandimath et al. 2016), which would affect the chemical composition of the wood material. E. dunnii (SQ4) exhibited the lowest diversity of indigenous microorganisms, which supports the idea of minimal degradation of cellulose fibers as evidenced by the high pulp yield and viscosity levels obtained for this species, making it the most suitable species for pulping.
Some of the bands on the gels remain unidentified, therefore the microbial diversity may be underrepresented. However, the principle of DGGE that one band represents one genus/species is not always correct as a single point mutation occasionally results in two bands (Miller et al. 1999; Muyzer and Smalla 1998; Adil 2015). Therefore, there is the possibility of multiple bands for a single species due to single base pair mutations in their DNA.
Numerous changes in the chemical and biological nature of the wood were observed after six months of storage at -20 ºC. This is unfavorable, particularly if the wood is to be used for experimental purposes. Previous studies have shown that storage of wood chips at -20 °C storage retards seasoning but does not stop it altogether (Allen et al. 1991; Dunlop-Jones et al. 1991; Back and Allen, 2000). The authors therefore recommend that wood material be stored for a maximum of three months under laboratory conditions. This recommendation is based on information from available literature (Allen et al. 1991; Back and Allen 2000).
- Storage of wood chip samples at -20 °C influenced the chemical nature of Eucalyptuswood, particularly the lipophilic extractives.
- This effect was similar to that seen with the traditional method of seasoning used for the reduction of lipophilic extractives. This is contrary to reports that storage of wood chips at -20 °C retards seasoning.
- Changes in bacterial and fungal communities were observed after storage, which should be taken into consideration when conducting lab scale trials.
- It is therefore recommended that if storage is necessary under laboratory conditions, it should be retained for a maximum of 3 months at -20 °C.
This work was supported by the National Research Foundation (NRF) and the Biorefinery Industry Development Facility at the Council for Scientific and Industrial Research (CSIR). The authors declare no conflict of interest.
Adair, S., Kim, S. H., and Breuil, C. (2002). “A molecular approach for early monitoring of decay basidiomycetes in wood chips,” FEMS Microbiol. Lett. 211, 117-122.
Adil, E. (2015). “Corrective measures of denaturing gradient gel electrophoresis limitations,” J. Environ. Sci. Technol. 8, 1-12.
Allen, L. H., Sithole, B. B., MacLeod, J. M., Lapointe, C., and McPhee, F. J. (1991). “The importance of seasoning and debarking in the kraft pulping of aspen,” J. Pulp Paper Sci. 17, J85-J91.
Almeida, J. M., and Silva, D. J. (2001). “Effect of extractives amount and liquor accessibility on Eucalyptus kraft pulping,” in: 34th Annual Pulp and Paper Meeting, Congresso Anual da ABTCP, 22–25 October, São Paulo, Brazil, pp. 1-13.
Álvarez, C., Reyes-Sosa, F. M., and Díez, B. (2016). “Enzymatic hydrolysis of biomass from wood,” Microb. Biotechnol. 9, 149-156.
Azhar, S. (2105). “Extraction of polymeric hemicelluloses from spruce wood,” Doctoral Thesis, Wallenberg Wood Science Center, Department of Fiber and Polymer Technology, School of Chemical Science and Engineering, KTH Royal Institute of Technology, Stockholm.
Back, E. L., and Allen, L. H. (2000). Pitch Control, Wood Resin and Deresination, TAPPI Press, Atlanta, pp. 170-174.
Bajpai, P. (1999). “Chapter 2: Wood retreatment to remove toxic extractives,” in: Biotechnology for Pulp and Paper Processing, Springer Science & Business Media, pp. 33-56.
Barry Kay, A., Bousquet, J., Holt, P. G., and Kaplan, A. P. (2009). “Part 5: Allergens,” in: Allergy and Allergic Diseases, John Wiley & Sons, pp. 972.
Burnes, T. (2000). “Wood degradation and biological pulping and bleaching,” Pulp Paper Industry pp. 2646-2651.
Burnes, T. A., Blanchette, R. A., and Farrell, R. L. (2000). “Bacterial biodegradation of extractives and patterns of bordered pit membrane attack in pine wood,” Appl. Environ. Microbiol. 66, 5201-5205.
Cohen, W. E., and Mackney, A. W. (1951). “Australian Pulp & Paper,” Ind. Tech. Assoc. Proc. 5, 315.
Costa, M. M., and Colodette, J. L. (2007). “The impact of kappa number composition on Eucalyptus kraft pulp bleachability,” Braz. J. Chem. Eng. 24, 61-71.
Dunlop-Jones, N., Jialing, H., and Allen, L. H. (1991). “An analysis of the acetone extractives of the wood and bark from fresh trembling aspen: Implications for deresination and pitch control,” J. Pulp Pap. Sci. 17, 60-66.
Fardim, P., and Durán, N. (2003). “Modification of fibre surfaces during pulping and refining as analysed by SEM, XPS and ToF-SIMS,” Colloids Surf A. 223, 263-276.
Fišerová, M., Opálená, E., and Illa, A. (2013). “Comparative study of hemicelluloses extraction from beech an oak wood,” Wood Res. 58, 543-554.
Freire, C. S. R., Pinto, P. C. R., Santiago, A. S., Silvestre, A. J. D., Evtuguin, D. V., and Neto, C. P. (2006). “Comparative study of lipophilic extractives of hardwoods and corresponding ECF bleached kraft pulps,” BioResources 1, 3-17.
Freire, C. S. R., Silvestre, A. J. D., and Neto, C. P. (2002). “Identification of new hydroxy fatty acids and ferulic acid esters in the wood of Eucalyptus globulus,” Holzforschung 56, 143-149.
Garg, G., Dhiman, S. S., Mahajan, R., Kaur, A., and Sharma, J. (2011). “Bleach-boosting effect of crude xylanase from Bacillus stearothermophilus SDX on wheat straw pulp,” New Biotech. 28, 58-64.
Gutiérrez, A., del Río, J. C., González-Vila, F. J., and Martín, F. (1999). “Chemical composition of lipophilic extractives from Eucalyptus globulus Labill. Wood,” Holzforschung 53, 481-486.
Gutiérrez, A., del Río, J. C., González-Vila, F. J., and Martín, F. (1998). “Analysis of lipophilic extractives from wood and pitch deposits by solid-phase extraction and gas chromatography,” J. Chromatogr. A 823, 449-455.
Gutiérrez, A., del Río, J. C., Martínez, M. J., and Martínez, A. (2001). “The biotechnological control of pitch in paper pulp manufacturing,” Trends Biotechnol. 19, 340-348.
Handschur, M., Piñar, G., Gallist, B., Lubitz, W., and Haslberger, A. G. (2005). “Culture free DGGE and cloning based monitoring of changes in bacterial communities of salad due to processing,” Food Chem. Toxicol. 43, 1595-1605.
Hansen, E. M. (1990). “Chapter 8: Storage molds,” in: Major Diseases: Storage Molds, Forest Research Laboratory, College of Forestry, Oregon State University pp. 25-26.
Haupt, M., Leithoff, H., Meier, D., Richter, H. G., and Faix, O. (2003). “Heartwood extractives and natural durability of plantation-grown teakwood (Tectona grandis L.) – A case study,” Holz als Roh- und Werkst. 61, 473-474.
Hillis, W. E. (2014). Wood Extractives and Their Significance to the Pulp and Paper Industries, Academic Press, ISBN 1483258610, 9781483258614
Hillis, W. E., and Sumimoto, M. (1989). “Effect of extractives on pulping,” in: Natural Products of Woody Plants, J.W. Rowe, (ed.), Springer-Verlag, Berlin, Germany pp. 880-920.
Hui, Y. H., and Sherkat, F. (2005). “Chapter 39: Freezing seafood and seafood products: Principles and applications,” in: Handbook of Food Science, Technology, and Engineering, CRC Press, pp. 15.
Ji, N., Peng, B., Wang, G., Wang, S., and Peng, X. (2004). “Universal primer PCR with DGGE for rapid detection of bacterial pathogens,” J. Microbiol. Methods. 57, 409-413.
Jørgensen, H., Kristensen, J. B., and Felby C. (2007). “Enzymatic conversion of lignocellulose into fermentable sugars: challenges and opportunities,” Biofuels Bioprod. Bioref. 1, 119-134.
Kilulya, K. F., Msagati, T. A. M., Mamba, B. B., Ngila, J. C., and Bush, T. (2014). “Effect of site, species and tree size on the quantitative variation of lipophilic extractives in Eucalyptus woods used for pulping in South Africa,” Ind. Crops Prod. 56, 166-174.
Kontkanen, H., Tenkanen, M., Fagerstrom, R., and Reinikainen, T. (2004). “Characterisation of steryl esterase activities,” J. Biotechnol. 108, 51-59.
Kropacz, A., and Fojutowski, A. (2014). “Colonization by fungi of wood chips stored in industrial conditions,” Drewno 57, 60-80.
Latorraca, J. V., Dünisch, O., and Koch, G. (2011). “Chemical composition and natural durability of juvenile and mature heartwood of Robinia pseudoacacia L.” An. Acad. Bras. Ciênc. 83, 1059-1068.
Leskinen, T., Salas, C., Kelley, S. S., and Argyropoulos, D. S. (2015). “Wood extractives promote cellulase activity on cellulosic substrates,” Biomacromolecules 16, 3226-3234.
Li, H., Saeed, A., Jahan, M. S., Ni, Y., and Van Heiningen, A. (2010). “Hemicellulose removal from hardwood chips in the pre-hydrolysis step of the kraft-based dissolving pulp production process,” J. Wood Chem. Technol. 30, 48-60.
Lindström, M. E., and Teder, A. (1995). “The effect of polysulfide pretreatment when kraft pulping to very low kappa numbers,” Nordic Pulp Paper Res. J. 10, 8-11.
Marchesi, J. R., Sato, T., Weightman, A. J., Fry, J. C., and Hiom, S. J. (1998). “Design and evaluation of useful bacterial-specific PCR primers that amplify genes coding for bacterial 16S rRNA,” Appl. Environ. Microbiol. 64, 795-799.
Miller, K. M., Ming, T. J., Schulze, A. D., and Withler, R. E. (1999). “Denaturing gradient gel electrophoresis (DGGE): A rapid and sensitive technique to screen nucleotide sequence variation in populations,” Biotechniques 27, 1016-1020.
Moodley, P. (2011). “Characterization and quantification of wood extractives and their impact on pitch,” Masters Thesis, University of KwaZulu-Natal, Durban, South Africa.
Muyzer, G., and Smalla, K. (1998). “Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology,” A. Van Leeuw. 73, 127-141.
Nandimath, A. P., Kharat, K. R., Gupta, S. G., and Kharat, A. S. (2016). “Optimization of cellulase production for Bacillus sp. and Pseudomonas sp. soil isolates,” Afri. J. Microbiol. Res. 10, 410-419.
Neiva, D. M., Gominho, J., and Pereira, H. (2014). “Modelling and optimization of Eucalyptus globulus bark and wood delignification using response surface methodology,” BioResources 9, 2907-2921.
Patrick, K. (2004). “Enzyme technology improves efficiency, cost, safety of stickies removal program,” Paper Age, September, 22-25.
Petäistö, R.-L. (2006). Botrytis cinerea and Norway Spruce seedlings in cold storage. Baltic Forestry 11, 24-33.
Prinsen, P., Gutiérrez, A., Rencoret, J., Nieto, L., Jiménez-Barbero, J., Burnet, A., Petit-Conil, M., Colodette, J. L., Martínez, A. T., and del Río, J. C. (2012). “Morphological characteristics and composition of lipophilic extractives and lignin in Brazilian woods from different eucalypt hybrids,” Ind. Crops Prod. 36, 572-583.
Rajala, T., Peltoniemi, M., Pennanen, T., and Mäkipää, R. (2010). “Relationship between wood-inhabiting fungi determined by molecular analysis (denaturing gradient gel electrophoresis) and quality of decaying logs,” Can. J. Res. 40, 2384-2397.
Ramnath, L., Bush, T., and Govinden, R. (2013). “Method optimization for denaturing gradient gel electrophoresis (DGGE) analysis of microflora from Eucalyptus sp. wood chips intended for pulping,” Afri. J. Biotechnol. 13, 356-365.
Rencoret, J., Gutiérrez, A., and del Rio, J.C. (2007). “Lipid and lignin composition of woods from different eucalypt species,” Holzforschung 61, 165-174.
Sappi 2012. www.typek.co.za/typek/applications/typek/templates/…/Eucalyptus.pdf
Sappi Unlimited. (2017). “Sustainability FAQs Dissolving wood pulp,” Document no. SCA-27, Revision no. 5, Revision date 22 March 2017. https://cdn-s3.sappi.com/s3fs-public/Sappi-FAQs-Dissolving-wood-pulp.pdf
Sarja, T. (2007). “Measurement, nature and removal of stickies in deinked pulp,” Doctoral Thesis, Department of Process and Environmental Engineering, Faculty of Technology, University of Oulu, Finland Acta Univ. Oul. C 275.
Schabereiter-Gurtner, C., Piñar, G., Lubitz, W., and Rölleke, S. (2001). “Analysis of fungal communities on historical church window glass by denaturing gradient gel electrophoresis and phylogenetic 18S rDNA sequences analysis,” J. Microbiol. Methods. 47, 345-354.
Schmidt, O. (2006). “Wood and tree fungi,” in: Biology, Damage, Protection, and Use, Springer-Verlag, Berlin, Heidelberg, pp. 94.
Seo, J. K., Park, T. S., Kwon, I. H., Piao, M. Y., Lee, C. H., and Ha, J. K. (2013). “Characterization of cellulolytic and xylanolytic enzymes of Bacillus licheniformis JK7 isolated from the rumen of a native Korean goat,” Asian-Australas. J. Anim. Sci. 26, 50-58.
Silvério, F. O., Barbosa, L. C. A., Maltha, C. R. A., Fidêncio, P. H., Cruz, M. P., Veloso, D. P., and Milanez, A. F. (2008). “Effect of storage time on the composition and content of wood extractives in Eucalyptus cultivated in Brazil,” Bioresour. Technol. 99, 4878-4886.
Silvério, F. O., Barbosa, L. C. A., Maltha, C. R. A., Silvestre, A. J. D., Pilo-Veloso, D., and Gomide, J. L. (2007). “Comparative study on the chemical composition of lipophilic fractions from three wood tissues of Eucalyptus species by gas chromatography and mass spectrometry (GC–MS) analysis,” J. Wood Sci. 53, 533-540.
Sithole, B., and Watanabe, C. (2013). “Using Py-GC/MS to detect and measure silicone defoamers in pulp fibres and mill deposits,” J. Anal. Appl. Pyrolysis 103, 8-16.
Sjöström, E. (1993). “Wood Chemistry,” in: Fundamentals and Applications, Academic Press, San Diego, CA.
Skyba, O., Douglas, C. J., and Mansfield, S. D. (2013). “Syringyl-rich lignin renders poplars more resistant to degradation by wood decay fungi, Appl. Environ. Microbiol. 79, 2560-2571.
Subramaniyan, S., and Prema, P. (2002). “Biotechnology of microbial xylanases: Enzymology, molecular biology, and application,” Crit. Rev. Biotechnol. 22, 33-64.
Swain, T. L., and Gardner, R. (2003). “Use of site-species matching and genetic gain to maximise yield – A South African example,” in: Eucalyptus Plantations – Research, Management and Development, Proceedings of the International Symposium, R.-P. Wei, and D. Xu (ed.), World Scientific, 04 Sep 2003 – Science – 432 pages.
Szulc, J., Otlewska, A., Okrasa, M., Majchrzycka, K., Sulyok, M., and Gutarowska, B. (2017). “Microbiological contamination at workspaces in a combined heat and power (CPH) station processing plant biomass,” Int. J. Environ. Res. Public Health 14, 99-117.
Tao, F., Liu, Y., and An, N. (2010). “On the necessity of organic solvent extraction for carbon isotopic analysis of α-cellulose: implications for environmental reconstructions,” Intern. J. Environ. Anal. Chem. 90, 605-619.
TAPPI. (1996) Test methods T249 cm-85: Carbohydrate anaylsis; T222 om-88: Acid insoluble lignin of wood and pulp; T235 OM- 60: Alkali solubility of pulp; T204 om-88: Solvent extractives; T203 om-94: Viscosity of pulp (capillary viscometer method). TAPPI Test Methods(1996-1997), TAPPI Press, Atlanta, USA.
Testova, L., Vilonen, K., Pynnönen, H., Tenkanen, M., and Sixta, H. (2009). “Isolation of hemicelluloses from birch wood: Distribution of wood components and preliminary trails in dehydration of hemicelluloses,” Lenzinger Berichte. 87, 58-65.
Venn, K. (1980). “Winter vigour in Picea abies (L) Karst. VII. Development of injury to seedlings during overwinter cold storage A Literature Review,” Meddr. Norsk Inst. Stogforsk. 35, 483-530.
Wallis, A. F. A., Wearne, R. H., and Wright, P. J. (1996). “Chemical analysis of polysaccharides in plantation eucalypt woods and pulps,” Appita J. 49, 258-262.
Wang, X. H., and Jiang, C. (2006). “Papermaking, Part I, Surface and colloid chemistry of pulping process,” in: Encyclopedia of Surface and Colloid Science, Vol. 6, P. Somasundaran (ed.), CRC Press pp. 4415-4434.
White, T. J., Burns, T., Lee, S., and Taylor, J. (1990). “Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics,” in: PCR Protocols, Innes et al. (ed.), Academic Press, San Diego, California, USA pp. 315-322.
Wright, P. J., and Wallis, A. F. A. (1996). “Rapid determination of carbohydrates in hardwoods,” Holzforschung 50, 518-524.
Yang, G., and Jaakkola, P. (2011). “Wood chemistry and isolation of extractives from wood,” Literature study for BIOTULI project, Saimaa University of Applied Sciences, South Karelia, Finland.
Zhang, X. H., Zhao, G. H., Li, D. W., Li, S. P., and Hong, Q. (2014). “Identification and evaluation of strain B37 of Bacillus subtilis antagonistic to sapstain fungi on poplar wood,” Scientific World J. 2014, article ID 149342.
Article submitted: July 11, 2017; Peer review completed: October 8, 2017; Revised version received: October 30, 2017; Accepted: October 31, 2017; Published: November 3, 2017.