METHOD FOR INCREASING CELLULOSE

IN SUGARCANE

FIELD OF THE INVENTION

The invention provides a method for increasing the cellulose and/or glucose content of sugarcane.

BACKGROUND TO THE INVENTION

Declining fossil fuel reserves have stimulated research into renewable energy sources such as plant-derived bioethanol. First generation biofuel technology relies on simple carbohydrates such as sucrose or glucose derived from starch as fermentable substrates, and therefore competes for limited resources, possibly jeopardizing food security (Abramson et al., 2010; Waclawovsky et al., 2010; Karp and Richter, 201 1 ). For the production of second generation biofuels, agricultural byproducts such as lignocellulosic biomass have been utilized (Pauly and Keegstra, 2010; Harris and DeBolt, 2010; Cook and Devoto, 201 1 ).

Cellulose is a rich source of fermentable glucose (Carroll and Somerville 2009) and is a major component of the cell wall. However, in the plant cell wall it cross-links with hemicellulosic polysaccharides and lignin (Vogel 2008). The interwoven matrix created by these three components renders lignocellulosic biomass recalcitrant to cellulolytic enzymatic hydrolysis (Abramson et al., 2010; Harris and DeBolt, 2010). This is ascribed to both the phenylpropanoid polymer lignin as well as to hemicellulose polysaccharides being coated in cellulose, which limits access of cellulolytic enzymes (Mosier et al., 2005). Moreover, when lignin and hemicellulosic polymers have been partially removed, cellulose remains resistant to enzymatic hydrolysis, due to the crystalline nature of the cellulose microfibrils (Himmel et al., 2007; Igarashi et al., 201 1 ). In order to efficiently utilize lignocellulosic biomass as a feedstock for biofuel production, it is important to reduce its natural resistance to saccharification. Sugarcane {Saccharum spp. hybrids) has been identified as a leading potential feedstock for biofuel production (Somerville et al., 2010). This perennial C ₄ grass has the ability to accumulate high lignocellulosic biomass and requires minimal light, water and nitrogen resources (Taylor et al., 2010; Waclawovsky et al., 2010). However, sugarcane bagasse is an underutilized substrate for bioethanol production due to the recalcitrant structure of its cell wall.

There is therefore a need for the development of crop plants from which cellulose can be accessed by hydrolytic enzymes for the production of second generation biofuel.

SUMMARY OF THE INVENTION

According to a first embodiment of the invention, there is provided a method for increasing the cellulose content of sugarcane, the method comprising the step of expressing in the sugarcane a heterologous polynucleotide which encodes a cellulose synthase having an amino acid sequence which is at least 70% identical to SEQ ID NO: 1 . The cellulose synthase may be from Ciona savignyi.

The cellulose synthase may have an amino acid sequence which is at least 80% identical to SEQ ID NO: 1 , at least 90% identical to SEQ ID NO: 1 , at least 95% identical to SEQ ID NO: 1 , or at least 98% identical to SEQ ID NO: 1 .

The heterologous polynucleotide encoding the cellulose synthase may have a nucleotide sequence which is at least 70% identical to SEQ ID NO: 2, at least 80% identical to SEQ ID NO: 2, at least 90% identical to SEQ ID NO: 2, at least 95% identical to SEQ ID NO: 2, or at least 98% identical to SEQ ID NO: 2.

The cellulose content may be increased in at least lignocellulosic material of the sugarcane.

The method may comprise an initial step of transforming a sugarcane cell with the heterologous polynucleotide. The cell may be a callus cell, which may be regenerated into a sugarcane plant. A vector may be used to transform the sugarcane cell.

The cellulose content may be increased relative to untransformed sugarcane.

Expression of the heterologous poynucleotide preferably does not result in a decrease in the sucrose content of the sugarcane relative to untransformed sugarcane.

According to a second embodiment of the invention, there is provided a method for increasing the yield of glucose from lignocellulosic material from sugarcane, the method comprising the step of:

saccharifying lignocellulosic material of sugarcane in which the heterologous polynucleotide encoding a cellulose synthase at least 70% identical to SEQ ID NO: 1 has been expressed as described above; and

recovering glucose from the saccharified lignocellulosic material.

The lignocellulosic material may be from sugarcane bagasse, i.e. from sugarcane from which sucrose juice has been extracted.

The saccharification step may be carried out without first pre-treating the lignocellulosic material, such as by heat treating the lignocellulosic material or treating it with an acid or base.

Alternatively, the lignocellulosic material may be subjected to a pre-treatment step (such as acid, base or heat treatment) prior to saccharification.

According to a third embodiment of the invention, there is provided a method for producing a biofuel from sugarcane, the method comprising the steps of:

saccharifying lignocellulosic material of sugarcane in which the heterologous polynucleotide encoding a cellulose synthase at least 70% identical to SEQ ID NO: 1 has been expressed as described above;

recovering glucose from the saccharified lignocellulosic material; and using the recovered glucose as a feedstock to produce the biofuel. The biofuel may be ethanol.

According to a further embodiment of the invention, there is provided a sugarcane plant cell, plant part or plant which has been transformed with a heterologous polynucleotide encoding a cellulose synthase at least 70% identical to SEQ ID NO: 1 as described above.

According to a further embodiment of the invention, there is provided a vector comprising a polynucleotide encoding a cellulose synthase at least 70% identical to SEQ ID NO: 1 as described above.

BRIEF DESCRIPTION OF THE FIGURES

Figure 1 : Phenotypic evaluation of greenhouse grown transgenic sugarcane and analysis of CsCesA integration and expression level in mature transgenic sugarcane plants. WT, Wild-type sugarcane; pCel8.3, pCel6.2 and pCeM .1 , Transgenic sugarcane plants, (a) Phenotypic evaluation of 18 months old greenhouse grown transgenic sugarcane in comparison with wild type, (b) PCR gel analysis of integration of CsCesA in genomic DNA of sugarcane, (c) RT-PCR gel analysis of CsCesA expression at mRNA level.

Figure 2: Analysis of total cellulose synthase enzyme activity and cellulose content in transgenic sugarcane in comparison with wild type control plants, (a) Total cellulose synthase enzyme activity in transgenic sugarcane internodal tissues. Y;

Younger internodes (internode1 -4); M; Mature internodes (internodes 14-15). WT;

Wild-type sugarcane, pCel transgenic sugarcane lines (b) Total cellulose content in transgenic sugarcane internodal tissues. Values are means ±SE (n=3). Values set in bold type were determined by the t test to be significantly (*) different (P<0.05) from the respective wild-type (WT) internodes.

Figure 3: Analysis of total uronic acids in transgenic sugarcane and wild-type internodal tissues. Y; Younger internodes (internode1 -4); M; Mature internodes (internodes 14-15) WT; Wild-type sugarcane, pCel transgenic sugarcane. Values are means ±SE (n=3). Values set in bold type were determined by the t test to be significantly (*) different (P<0.05) from the respective wild-type (WT) internodes.

Figure 4: Evaluation of saccharification efficiency of transgenic sugarcane and wild-type plants. Wild type, ·; pCel1.1 , D;pCel6.2, pCel8.3■; (a) Saccharification of young internodal tissues, (b) Saccharification of mature internodal tissues, (c) Correlation of total cellulose content and glucose released during saccharification of lignocellulose biomass. (d) Correlation of total lignin content and glucose released during saccharification of lignocellulose biomass. (e) Correlation of total lignin monomer unit content and glucose released during saccharification of lignocellulose biomass. Values are means ±SE (n=3). Values were determined by the t test to be significantly different (P<0.05) from the respective wild type (WT).

Figure 5: Amino acid sequence of cellulose synthase from Ciona savignyi (SEQ ID NO: 1 ).

Figure 6: cDNA sequence of cellulose synthase from Ciona savignyi (SEQ ID NO: 2).

DETAILED DESCRIPTION OF THE INVENTION

A method for increasing the cellulose and/or glucose content of sugarcane is described herein, in which a heterologous cellulose synthase DNA is expressed in the sugarcane plants.

In a preferred embodiment, the cellulose synthase is CesA from Ciona savignyi (SEQ ID NOs: 1 and 2 (Figures 5 and 6)). The cellulose synthase can have an amino acid sequence which is at least 70% identical to SEQ ID NO: 1 . Alternatively, the cellulose synthase can have an amino acid sequence which is at least 70% identical to SEQ ID NO: 1 , at least 80% identical to SEQ ID NO: 1 , at least 90% identical to SEQ ID NO: 1 , at least 95% identical to SEQ ID NO: 1 , or at least 98% identical to SEQ ID NO: 1 . The heterologous polypeptide encoding the cellulose synthase can have a nucleotide sequence which is at least 70% identical to SEQ ID NO: 2. Alternatively, the heterologous polypeptide can have a nucleotide sequence which is at least 70% identical to SEQ ID NO: 2, at least 80% identical to SEQ ID NO: 2, at least 90% identical to SEQ ID NO: 2, at least 95% identical to SEQ ID NO: 2, or at least 98% identical to SEQ ID NO: 2. Cellulose is composed of unbranched β-(1 ,4)-linked glucan chains that are synthesised at the plasma membrane (Saxena and Brown 2005). It is produced by a diverse group of organisms, including plants, bacteria, cellular slime moulds and one group of marine invertebrates, the urochordates (Brown et al., 1996; Kimura and Itoh 2004). The gene implicated in cellulose synthesis encoding cellulose synthase {CesA), has been identified and functionally characterized in many of these organisms (Pear et al., 1996; Ariola et al., 1998; Matthysse et al., 2004). Heterologous expression of bacterial or native CesA genes in cotton and potato, respectively, resulted in transgenic plants with increased cellulose contents (Li et al., 2004; Oomen et al., 2004). Although a number of studies have been conducted in sugarcane to introduce new metabolic carbon sinks through production of novel sugars and biopolymers (Hamerli and Birch 201 1 ; Basnayake et al., 2012; Bauer et al., 2012), cellulose increases for lignocellulosic biomass improvement in sugarcane have not been reported. Recently a transgenic approach reduced the lignin content of sugarcane cell wall through repression of the comt gene (Jung et al., 2012). Mature transgenic sugarcane lines had improved lignocellulosic saccharification efficiencies, yielding increases of the fermentable sugar glucose, with or without diluted acid pretreatment. Over-expression of cellulose synthase cDNAs in plants have been reported (Li et al., 2004; Oomen et al., 2004). In potato, for example, over-expression of the native cellulose synthase StCesA3, resulted in potato tubers with elevated cellulose contents of up to 200% (Oomen et al., 2004). Expression of bacterial cellulose biosynthesis genes acsA and acsB from Acetobacter xylinum in cotton resulted in transgenic plants with a higher cellulose content (Li et al., 2004).

The applicant has now shown that heterologous expression of a cellulose synthase cDNA from the animal Ciona savignyi (CsCesA; Matthysse et al., 2004) leads to the generation of transgenic sugarcane plants that have increased cellulose content. Moreover, the increased cellulose content resulted in plants with high hemicellulose glucose content, reduced lignin and improved lignocellulosic saccharification efficiency.

Although CsCesA expression has been shown to restore cellulose biosynthesis in cellulose deficient Agrobacterium tumefaciens, the applicant is unaware of any earlier reports of an animal CesA having been expressed in plants.

The invention will now be described in more detail by way of the following non-limiting examples.

Materials

All chemical were obtained from Sigma-Aldrich Fluka Chemical Company (St. Louis, MO, USA) or Roche Diagnostics (Basel, Switzerland), unless stated otherwise. All auxiliary enzymes, cofactors and substrates used for enzyme assays and metabolite determinations were purchased from either Sigma-Aldrich Fluka (SAF) Chemical Company (St. Louis, MO, USA) or Roche Diagnostics (Basel, Switzerland), unless stated otherwise. Nucleic acid modifying enzymes were obtained from either Promega (South Africa) or Fermentas (Inqaba Biotech, South Africa). Oligo nucleotide primers were purchased from Integrated DNA Technologies (IDT, Whitehead Scientific, South Africa) or Fermentas (Inqaba Biotech, South Africa).

Methods

Vector construction

Cellulose synthase cDNA (SEQ ID NO: 2) was excised from the pTE3 plasmid (Matthysse et al., 2004; GenBank Accession No. AY504665.1 ) by restriction enzyme digestion using EcoRI. Approximately 4.6 kbp was separated by gel electrophoresis and extracted using GeneJET ^Tm Extraction kit (Inqaba Biotech, South Africa). It was then ligated in sense orientation to the maize ubiquitin promoter into the EcoRI site of the pUBI510 (Groenewald et al., 2000) plant transformation vector resulting in the construct pCel.

Generation of transgenic sugarcane

Sugarcane embryogenic callus initiation, transformation and selection were carried out according to standard sugarcane transformation procedures from the commercial variety NCO310 leaf roll tissue (Bower and Birch 1992; Synman et al., 1996). The cellulose synthase cDNA construct (pCel) was co-transformed with pEmuKN vector containing the npt-ll selectable marker. Transformed embryogenic callus was selected for geneticin resistance and putative transgenic plants from independent transformation events were regenerated (Snyman et al. 1996). Regenerated plants were grown in a containment greenhouse under conditions of 16 hour light periods and at 26 °C until 18 months before characterization.

Polymerase chain reaction (PCR) analysis

The presence of the cellulose synthase construct (pCel) in genomic DNA of transgenic tissue was evaluated by PCR. Genomic DNA was extracted in sugarcane tissues as described by McGarvey and Kaper (1991 ) and 50 ng DNA was used as template per reaction. Gene specific primer sets which were used are: CsCesA Fwd: TTG CAA TGA GCA GGG ATA GA (SEQ ID NO: 3) and CsCesA Rev: TCT TCC GAA TAA CCC GAT TG (SEQ ID NO: 4) (amplified fragment ±380 bp) and a primer set designed to the 35S promoter: UBI Fwd: ATA CGC TAT TTA TTT GCT TGG (SEQ ID NO: 5) in combination with CsCesA Rev amplified fragment ± 780 base pair.

Total RNA extraction and first strand cDNA synthesis

Mature greenhouse grown sugarcane plants internodal tissues, internode 1 -4 (younger internodes) and internodes 14-15 (mature internodes) were used to screen for expression of the CsCesA cDNA. Total RNA was extracted from 2 g internodal tissue according to Bugos et al. (1995). Total RNA was treated with RNase free DNase I (Fermentas) according to the manufacturer's instructions. First strand cDNA was prepared using ^g of DNase treated total RNA and reversed transcribed using Revertaid™ H minus first strand cDNA synthesis kit (Fermentas, Inqaba Biotech, South Africa) according the manufacturer's instructions.

Analysis of Ciona savignyi CsCesA transcript level in sugarcane by semi-quantitative RT-PCR

Specific primers (CsCESA Fwd and CsCESA Rev) for CsCesA were used to amplify a fragment of approximately 380 bp. The forward: Fwd 25S RNA (TGA AAG CGT GGC CTA TCG ATC CTT (SEQ ID NO: 6)) and reverse: Rev 25S RNA (AGG ATT GGA CCA ACC GAT GAC GAT (SEQ ID NO: 7)) primers for sugarcane 25S ribosomal RNA reference gene for RT-PCR normalization were designed as described by Iskandar et al. (2004). The cDNA was diluted to 10 ng with milliQ water before being used in the semi quantitative RT-PCR. Amplicons were separated on a 1 % agarose gel and spot densitometry was conducted by ascertaining the integrated density value of each PCR fragment, using the auto background option of AlphaEase image-analysis software version 4.0.1 (Alpha Innotech, San Leandro, CA, USA).

Cellulose synthase activity determination

Protein extracts were prepared from young and mature internodal tissues according to a modified method of Schafer et al. (2004). Internodal tissues were homogenized in ice cold extraction buffer (50 mM Tris-HCI pH 7.2, 10 mM MgCI ₂ , 2 mM EDTA, 5 mM DTT, 10% (v/v) glycerol and 0.0016 mg/l Complete protease inhibitor) before being centrifuged at 4°C for 15 min at 5 000 g. The supernatant was filtered through two layers of nylon cloth at 4°C and centrifuged at 4°C for 60 min at 100 000 g. The resulting pellet was re-suspended in a buffer containing 50 mM Tris-HCI pH 7.5, 1 % (v/v) Triton X100. The crude protein extract was incubated with reaction buffer containing 50 mM Tris-HCI pH 7.5, 8 mM MgCI ₂ , 8mM CaCI ₂ , 20 mM cellobiose, 1 mM (UDP-glucose and UDP-D-[U- ¹⁴ C] glucose) and the reaction incubated at room temperature for 30 min. The reaction was stopped by the addition of 0.5 M NaOH and heated at 65 °C for 20 min. The samples were filtered through glass fiber filters, washed twice with milliQ water, three times with 70% (v/v) ethanol and dried. The filters were submerged in 4 ml scintillation cocktail and radioactivity measured in a scintillation counter (Beckman Coulter).

Preparation of alcohol insoluble residue (AIR) and removal of starch

2 g of young (internodes 1 -4) and mature (internodes 14-15) internodal tissues was added to 80% (v/v) ethanol and incubated at 80 °C for 1 hour. The samples were centrifuged at 5 000 g for 10 minutes and the supernatant discarded. The extraction process was repeated three times. The alcohol insoluble residue (AIR) was washed with acetone, vacuum dried and stored in a desiccator under vacuum until required.

AIR (1 g ± 10 mg) was mixed with 50 mM sodium acetate buffer pH 4.8 and incubated at l OCC for 1 hour. After cooling to room temperature, 10 U amyloglucosidase from Aspergillus niger (Fluka) was added and incubated overnight (16-18 hours) at 55°C and the reaction terminated by incubating the sample at l OCC for 10 minutes. Samples were centrifuged at 10 000 g for 10 minutes and washed three times with 80% (v/v) ethanol. AIR was washed with acetone and vacuum dried and stored as before.

Cellulose content determination

The cellulose content was determined according to a modified Updegraff method (1969). In brief, 10 mg ± 1 mg of destarched AIR was boiled in 1 ml acetic- nitric acid reagent (acetic acid: nitric acid: water: 8:1 :2) for 30 minutes and samples cooled to room temperature and centrifuged at 8 000 g for 10 minutes. The cellulosic residue was washed three times in 8 ml milliQ water and followed by a 4 ml acetone wash and vacuum dried. The cellulosic material was then completely hydrolysed in 500 μΙ 67% (w/w) sulphuric acid at 30 °C for 30 minutes. The glucose content of the samples was determined by the anthrone method as described by Leyva et al. (2008).

Pectic and hemicellulosic polysaccharides extraction of alcohol insoluble residue

AIR (100 mg ± 1 mg) was depectinated by treatment with 5 ml of 1 % (w/v) ammonium oxalate and incubation at 100°C for 1 hour. Samples were centrifuged at 3 500 g for 5 minutes, the supernatant removed, and the ammonium oxalate extraction repeated with the supernatants being combined. The remaining insoluble residue was washed twice with 5 ml milliQ water and all the supernatants pooled with the ammonium oxalate supernatant. The samples were neutralized with acetic acid to pH 7 and dialyzed for 24 hours a\ 4°C in milliQ water (water change four times) containing 0.1 % (v/v) sodium azide. The dialyzed supernatant and depectinated alcohol insoluble residue were freeze dried and stored in air tight tubes in a desiccator under vacuum until required. Extraction of the hemicellulosic fractions was conducted using depectinated alcohol insoluble material (70 mg ± 1 mg). Samples were treated sequentially with increasing amounts of potassium hydroxide with sodium tetraborate (0.1 M KOH/ 20 mM NaBH ₄ , 0.5 M KOH/ 20 mM NaBH ₄ and 4 M KOH/ 20 mM NaBH ₄ ). Samples were added to 5 ml 0.1 M KOH/ 20 mM NaBH ₄ and incubated for 24 hours at room temperature with shaking to extract hemicellulosic material. These were centrifuged at 3 500 g for 5 minutes and the alcohol insoluble residue washed twice with milliQ water. The resulting supernatants were pooled, neutralized to pH 7 with acetic acid and dialyzed for 24 hours at 4°C in milliQ water (water change four times) containing 0.1 % sodium azide. The extraction was repeated as above with 0.5 M KOH/ 20 mM NaBH ₄ and 4 M KOH/ 20 mM NaBH _4. The dialyzed supernatant fractions of KOH and the remaining cellulosic alcohol insoluble residue were freeze dried and stored in air tight tubes in a desiccator under vacuum until required.

Hydrolysis of pectic and hemicellulosic cell wall polysaccharides fractions

Two milligram (2 mg ± 0.1 mg) of freeze dried pectin and hemicellulosic fractions were hydrolyzed with 1 ml of 2 M TFA at 121 °C for 2 hours. The samples were then cooled and centrifuged at 8 000 g for 10 minutes and TFA resistant residue was washed twice with 2 M TFA. The resulting supernatants were pooled, and TFA evaporated. To remove residual TFA, monosaccharides were dissolved in 1 ml methanol and the methanol evaporated as before. This process was repeated twice and with the last wash 1 μΙ ribitol (2 mg/ml) was added as an internal standard.

Monosaccharide derivatisation and GC-MS analysis

The pectic and hemecellulosic TFA hydrolysed monosaccharides and internal standard (ribitol) were derivatised and analyzed by GC-MS according to Roessner et al., 2000. The gas chromatography was conducted on a 30 m Rtx ^® -5Sil MS column (RESTEK) with Integra guard with an inner diameter of 0.25 mm and 0.25 mm film thickness, the system consisted of an AS 2000 autosampler, trace GC and the quadropole trace MS (ThermoFinnigan). 1 μΙ of sample was injected with a splitless injection and the flow rate was 1 ml/min. The injection temperature was 230 ^q C and the ion source temperature was set at 200 °C. The temperature of the program was as follows: 5 min at 70°C, followed by a 1 'C/rnin oven ramp to 76°C and a second ramp of 6°C/min to 350 °C and the temperature equilibrated to 70 °C before injection of the next sample. The mass spectra were recorded at two scans per sec with the scanning range of 500- 600 m/z. The chromatograms and mass spectra were evaluated using Xcalibar software bundle version 1 .2 (Finnigan Corporation 1998-200) and eluting compounds were identified using the NIST library (www.nist.gov). The resulting response values were calculated by dividing the peak area of the compound with the peak area of the internal standard (ribitol) and then dividing by the mass of the tissue used.

Determination of total uronic acid

Uronic acids content was determined (5 mg ± 0.1 mg of destarched alcohol insoluble residue) as described by Filisetti-Cozzi and Carpita (1991 ). To calculate the uronic acid concentration, a galacturonic acid standard curve was obtained using known concentration amounts (0-40 μg). Total liqnin and monomer composition determination

Klason lignin was determined by a modified method described by Huntley et al. (2003). Destarched alcohol insoluble material (100 mg ± 1 mg) was added to 1 ml of 72% (w/w) sulphuric acid and incubated at 20°C for 2 hours with constant mixing. The samples were diluted with water (28 ml) and autoclave at 121 °C for 1 hour. Samples were cooled to approximately 60 °C, and vacuum filtered through a pre-weighed glass filter and the glass filter washed with milliQ water to removed residual acid and sugars. The glass filters were dried overnight at Ι Οδ'Ό and weighed to determine the Klason acid insoluble lignin. Klason acid soluble lignin was determined by measuring the filtrate absorbance at 205 nm.

Lignin monomer composition was determined according to Robinson and Mansfield (2009) using 2 mg of destarched alcohol insoluble material. The gas chromatography was conducted on a 30 m Rtx ^® -5Sil MS column (RESTEK) with Integra guard with an inner diameter of 0.25 mm and 0.25 mm film thickness. The system consisted of an AS 2000 autosampler, trace GC and the quadropole trace MS (ThermoFinnigan). One microlitre of sample was injected with a splitless injection and the flow rate was 1 .1 ml/min with a 30 min solvent delay. The temperature of the program was as follows: initial hold at 130°C for 3 min, a 3 °C/m n ramp to 250°C and hold for 1 min to allow equilibration to the initial temperature of 130^. The peaks were identified by mass spectrum ions of 299 m/z and 269 m/z for S and G respectively. The resulting response values were calculated as before. Enzymatic saccharification of sugarcane liqnocellulosic biomass

Micro-scale enzyme saccharifications were performed as described by Stork et al. (2009) with some modification. Destarched alcohol insoluble residue (40 mg ± 0.1 mg) was placed in 5 ml tubes and mixed with 2 ml of buffer consisting of 50 mM sodium citrate ph 4.8, 0.1 % (v/v) sodium azide, 60 filter paper unit Celluclast 1 .5L and 134 U/ml cellobiose (Novozyme). The samples were shaken at 100 rpm in a horizontal position at 50°C. The progress of the reaction was measured by taking 100 μΙ aliquots at 2, 18, 24 and 48 h and the reaction terminated by incubating the sample at 100°C for 15 minutes. The glucose released was quantified according to Bergmeyer and Bernt (1974). The compositions of sugars released by enzymatic hydrolysis were qualitatively identified by GC/MS as before. Extraction and determination of soluble sugars

Soluble sugars in the internodal tissues were extracted according to a modified method by Bindon and Botha (2002). Young (internodes 1 -4) and mature (internodes 14-15) internodal tissues (100 mg ± 10 mg) were added to the extraction buffer (30 mM Hepes 5 mM MgCI ₂ and 80% (v/v) ethanol) and incubated at 80 °C for 1 hour. The samples were centrifuged at 10 000 g for 10 minutes and the extraction process was repeated three times. The resulting supernatants were pooled, vacuum dried (Genevac LTD, Ipswich, England) and re-suspended in 1 ml milliQ water. The soluble sugars were determined according to Bergmeyer and Bernt (1974). All readings were monitor at 340nm on the PowerWaveX spectrophotometer plate reader (Bio-Tek Instruments, Winooski, VT, USA).

Extraction and determination of starch concentrations

The starch content was quantified from the remaining alcohol insoluble material, 50 mg ± 1 mg was added to 50 mM sodium acetate buffer pH 4.8 and incubated at ^~ \ 00 °C for 1 hour. The samples were cooled to room temperature, 5 U amyloglucosidase (AMG) from Aspergillus niger (Fluka) was added and incubated overnight (16-18 hours) at 55 °C and the reaction terminated by incubating the sample at 100°C for 15 minutes. Samples were centrifuged at 10 000 g for 10 min, washed twice with 70% ethanol and the supernatant pooled and vacuum dried. Background glucose and glucose released by AMG were quantified according to Bergmeyer and Bernt (1974) method.

Extraction and determination of hexose phosphate and UDP-Glucose

Hexose phosphate and UDP-Glucose pools in the internodal tissues were extracted according to Stitt et a/ (1989). The supernatant was applied to activated Supelclean 100 mg ENVI-Carb SPE (Supelco) columns and the flow through containing hexose phosphate was retained as described by Rabina et al. (2001 ). The hexose phosphate and nucleotide sugar supernatant were freeze dried and resuspended in 250 μΙ milliQ water. The hexose phosphates (Gluc-6-P, Fruc-6-P and Glu-1 -P) were quantified as described by Lang and Michal (1974) and UDP-glucose was determined as described by Chai et al. (2004) spectrophotometrically at 340 nm. Statistical analyses

The student's t tests (two-tailed) were performed to test for significant differences between group means using Statistica version 7 (StatSoft, Inc. 2004). The term significant is used to indicate differences for which P < 0.05.

Results

Integration of C. savignyi CesA in sugarcane

Sugarcane was transformed using a vector (pCel) designed to express the CsCesA cDNA constitutively in transgenic plants. Following transformation ten independent sugarcane lines were found to contain the CsCesA sequence in the genome. These were grown in callus suspension cultures subsequently. Three of those lines were found to secrete a water soluble glucose polymer into the medium. This polymer was isolated from the suspension culture medium and shown to be cellulose by subjecting it to cellulase treatment. This led us to hypothesize that these clones were functionally expressing the CsCesA transgene, so we generated plants from them and transferred these into the greenhouse. The mature transgenic plants were morphologically similar to untransformed controls throughout the 18 months period in the glasshouse (Figure 1 a). Three lines pCel1 .1 , pCel6.2 and pCel8.3 were chosen for further investigation. Polymerase chain reaction (PCR) analysis confirmed integration of the transgene into their genome (Figure 1 b) while CsCesA mRNA accumulation was also shown using reverse transcription PCR (RT-PCR) (Figure 1 c).

Total cellulose synthase enzyme activity and cellulose content in mature transgenic sugarcane

Significant increases in the total cellulose synthase enzyme activity of between 20- 74%, in both young and mature internodes was measured in all the transgenic lines (Figure 2a). This alteration corresponded with increased cellulose contents in the transgenic lines of between 13-28% in both young and mature internodal tissues in the transgenic lines (Figure 2b).

Effects of increasing the cellulose contents on soluble sugars pools

Glucose and fructose increased slightly in both young and mature internodal tissues, while sucrose was slightly increased in younger internodes (Table 1 ). Hexose phosphate (glucose-1 -P, glucose-6-P and fructose-6-P) and UDP-glucose pools were determined. The younger internodes of all transgenic lines contained generally more hexose phosphates and UDP-glucose (Table 1 ). However, these differences disappeared in the mature internodes. Cell wall composition is altered in CsCesA expressing lines

The increased cellulose contents affected the biosynthesis of the cell wall hemicelluloses and pectic polysaccharides. Total cell wall sugars in young and mature internodal tissues of transgenic plants were significantly increased due to a rise in pectic and hemicellulosic glucose, galacturonic acid and galactose. However, there were no alterations in mannose, rhamnose, arabinose or xylose (Table 2). All transgenic plants had significant increases in total uronic acids (Figure 3).

Total lignin content of the transgenic plants was significantly reduced by between 5- 13% in both young and mature internodes (Table 3). Correspondingly the lignin monomers syringyl (S) and guaiacyl (G) were also reduced (Table 3). The drop in the lignin monomer content resulted in a higher S/G ratio of the young and mature internodes of all transgenic plants with the only exception being in mature internodes of pCel6.2M. Alterations in cell wall components improves the enzymatic saccharification of untreated biomass material

Based on alterations in the transgenic plants cell wall composition, the lignocellulosic biomass was examined for its susceptibility to saccharification by cellulosic enzymes. Internodal tissue from all transgenic lines showed significant improvement in their saccharification efficiencies. The highest was observed in the younger internodal tissues of line pCeM .I Y with 39% with pCel6.2Y and pCel8.3Y having 28% and 32% respectively (Figure 4a). In the mature internodes pCel8.3M had the highest saccharification efficiency of 28%, with pCeM .I M and pCel6.2M having 13% and 5% respectively in comparison to the wild type (Figure 4b). To confirm that cellulose was hydrolyzed to its monomer glucose, the fermentable sugars released by the enzymatic hydrolysis were analyzed by GC-MS. As expected, the main sugar released by cellulase and cellobiase hydrolysis was glucose, at more than 93%, followed by trace amounts of xylose and arabinose in all transgenic internodal tissues. There was a positive correlation between cellulose content and saccharification efficiency (Figure 4c), and accordingly glucose released after saccharification clearly correlated negatively with the lignin content (Figures 4d and e).

Table 1. Analysis of soluble sugars in the internodal tissues of transgenic and wild type sugarcane plants. WT; wild-type; pCel; Transgenic sugarcane lines Y; Younger internodes; M; Mature internodes. Values are means ± SE (n=3). Values set in bold type were determined by the t test to be significantly different (P<0.05) from therespective wild type internodes.

Concentration (μΓηοΙ/g FW) Concentration (nmol/g FW)

Plant Line Glue Fruc Sue Total Gluc-6-P Gluc-1-P Fruc-6-P UDP-Gluc

WTY 128 ±1.1 71 ± 1 .3 150 ±2.1 349 ±4.5 2.8 ±0.04 1.5 ±0.03 1.0 ±0.05 0.006 ± 0.004 pCeM.IY 147 ±0.9 81 ± 1 .1 190 ±1.6 418 ±3.5 5.1 ± 0.05 3.5 ± 0.04 1.8 ±0.03 0.018 ±0.003 pCel6.2Y 145 ± 1.3 89 ± 1 .0 169 ±2.1 385 ±4.4 3.5 ± 0.01 2.9 ± 0.03 1.1 ±0.02 0.011 ±0.002 pCel8.3Y 170 ± 1.5 98 ± 1 .5 195 ± 1.2 443 ±4.2 4.1 ± 0.03 3.0 ± 0.03 1.9 ±0.04 0.017 ±0.003

WTM 98.1 ±2.6 52 ± 1 .3 389 ±2.2 539 ±6.1 0.8 ±0.02 0.5 ±0.02 0.3 ±0.03 0.008 ± 0.004 pCeM.IM 159 ± 1.5 68 ± 1 .0 391 ±1.9 618 ±4.4 0.7 ±0.03 0.6 ±0.03 0.4 ±0.07 0.010 ±0.006 pCel6.2M 198 ±2.4 70 ± 1 .1 399 ±2.3 667 ±5.8 1.0 ±0.03 0.7 ±0.01 0.3 ±0.05 0.009 ± 0.005 pCel8.3M 161 ± 1.6 78 ± 1 .5 393 ±2.1 632 ±5.2 0.9 ±0.03 1.0 ±0.02 1.3 ±0.05 0.008 ± 0.005

Glu, Glucose; Fruc, Fructose; Sue, Sucrose; Gluc-6-P, Glucose-6-phosphate; Gluc-1-P, Glucose-1 -phosphate; Fruc-6-P, Fructose -6-phosphate; UDP-Gluc, Uridine-diphosphate glucose.

Table 2. Composition of hemicelluloses and pectin sugars monomers in internodal tissues of transgenic in comparison with the wild type sugarcane plants cell wall. WT; Wild type sugarcane. pCel; Transgenic sugarcane lines. Y; Young internodes. M; Mature internodes. Values are means ± SE (n=3). Values set in bold type were determined by the t test to be significantly different (P<0.05) from the respective wild type internodes. (Relative abundance/mg DW)

Plant Line Mannose Rhamnose Galacturonic acid Galactose Glucose Arabinose Xylose Total sugars

WTY 0.26 ± 0.03 0.36 ± 0.05 0.92 ± 0.04 5.65 ± 0.50 12.0 ±1.02 28.8 ±1.71 79.7 ±2.50 128 ±5.40 pCeM.IY 0.29 ± 0.07 0.38 ± 0.04 1.27 ±0.03 5.29 ± 0.61 27.5 ±2.10 29.2 ±1.02 83.9 ±3.11 148 ±6.98 pCel6.2Y 0.22 ± 0.02 0.36 ± 0.07 1.22 ±0.02 5.64 ± 0.52 24.4 ± 1.31 28.2 ±1.53 79.9 ± 2.93 140 ±6.40 pCel8.3Y 0.20 ±0.06 0.36 ± 0.09 1.97 ±0.04 5.68 ± 0.62 23.2 ± 1.42 29.1 ±1.53 79.7 ±1.54 135 ±5.30

WTM 0.24 ± 0.04 0.20 ± 0.07 0.76 ± 0.05 4.44 ± 0.45 19.8 ±1.33 21.1 ±1.72 79.9 ±1.96 126 ±5.62 pCeM.IM 0.19 ±0.05 0.18 ±0.08 0.98 ± 0.04 4.96 ± 0.41 23.9 ± 1.4 22.5 ±1.14 82.1 ±2.23 135 ±5.35 pCel6.2M 0.19 ±0.04 0.18 ±0.02 0.92 ± 0.03 5.71 ± 0.35 22.8 ± 1.50 21.8 ±1.62 78.9 ±1.54 130 ±5.10 pCel8.3M 0.21 ±0.08 0.19 ±0.07 0.98 ± 0.05 5.74 ±0.12 22.4 ± 0.9 22.9 ± 1.9 78.6 ±1.52 131 ±4.64

Table 3. Analysis of total lignin content and monomer unit composition in transgenic and wild type sugarcane plants. WT ; Wild type; pCel; Transgenic sugarcane lines. Y; Younger internodes; M; Mature internodes. Values are means ± SE (n=3). Values set in bold type were determined by the t test to be significantly different (P<0.05) from the respective wild type.

Klason Lignin (mg/g DW) Thioacidolysis (ug/mg DW)

Plant Line Acid insoluble Acid soluble Total Lignin G Lignin S Lignin S/G Total yield

WTY 0.16 ±0.04 126 ±3.5 127 ±3.6 1.08 ±0.07 0.70 ± 0.04 0.65 1.79 ±0.11 pCeM.IY 0.10 ±0.02 115 ±3.3 115 ±3.3 0.27 ± 0.06 0.21 ± 0.03 0.77 0.48 ± 0.09 pCel6.2Y 0.12 ±0.04 120 ±3.5 120 ±3.5 0.61 ± 0.04 0.49 ± 0.05 0.81 1.10 ±0.09 pCel8.3Y 0.09 ± 0.03 110 ±4.2 110 ±4.2 0.43 ± 0.05 0.29 ± 0.02 0.66 0.72 ± 0.07

WTM 0.03 ± 0.00 170 ±5.7 170 ±5.7 1.09 ±0.09 1.41 ± 0.34 1.28 2.49 ± 0.43 pCel1.1M 0.01 ± 0.00 140 ±3.8 159 ±3.8 0.81 ± 0.02 1.13 ±0.40 1.40 1.94 ±0.42

_P Cel6.2M 0.02 ± 0.00 146 ±6.7 161 ±6.7 0.89 ± 0.07 1.01 ±0.51 1.13 1.91 ±0.58

_P Cel8.3M 0.01 ± 0.00 154 ±4.7 155 ±4.7 0.81 ± 0.07 1.07 ±0.35 1.32 1.88 ±0.42

Discussion

Expression of CsCesA mRNA resulted in increased total cellulose synthase (CesA) activity in both young (29-74%) and mature (20-33%) internodal tissues. This led to concomitant rises in the cellulose contents of between 17-31 % in young and 13-28% in mature internodes. There are a number of pleiotropic alterations in metabolite concentrations in the transgenic plants. As sucrose is thought to be the main source for the production of UDP-glucose, the substrate of the CesA complex, then increases in cellulose might be expected to decrease sucrose concentrations. In fact these were slightly increased in young tissue, along with glucose and fructose. This indicates that sucrose concentrations are maintained in sugarcane culm, most likely through increased sucrose cycling in the transgenic plants, something supported by the increased UDP-glucose and hexose phosphates concentrations which are allosteric activators of sucrose phosphate synthase (Reimhoiz et al., 1994; van der Merwe et al., 2010).

The recalcitrant nature of lignocellulosic biomass to saccharification is associated with a number of factors including the lignin content, the lignin monomer unit composition, the hemicelluloses and the cellulose microfibril crystallinity (Chen and Dixon, 2007, Himmel, 2007; Dien et al., 2009; Fu et al., 201 1 ; Jung et al., 2012). Genetic modification and mutations of the cellulose, hemicelluloses, pectic and lignin biosynthetic genes in Arabidopsis, sorghum and alfalfa plants were shown to have improved saccharification efficiency leading to as much as twice the amount of sugar being released from the lignocellulosic feedstock (Dien et al., 2009; Harris et al., 2009; Lionetti et al., 2010; Fu et al., 201 1 ). However, the feedstock biomass was pre-treated with high temperatures and acidic or basic solutions that produce inhibitory products (furfural and hydroxymethylfurfural) which can subsequently inhibit hexose fermenting microorganisms and reduce the ethanol yield (Gamez et al., 2004; Klinke et al., 2004). Transgenic sugarcane lines where the lignin content was reduced through repression of the COMT gene had improved saccharification efficiencies without pre-treatment avoiding inhibitory compound accumulation (Jung et al., 2012). Due to the cellulose increase seen in the transgenic lines investigated in this study, whether this was accompanied by alterations in other cell wall components which may improve lignocellulose saccharification was examined. When sugarcane internodal lignocelluloses where fractionated with ammonium oxalate and alkali solution, glucose, galactose, arabinose and xylose were the major monosaccharides (62%, 15%, 9% and 9% respectively) in the oxalate fractions, while xylose was dominant in the 0.1 M and 1 M NaOH fractions and glucose was the main sugar in the 4 M NaOH fraction (Souza et al., 2012). The total sugar quantities of the hemicellulosic and pectic fractions in the oxalate and KOH extracts were altered in the transgenic lines, with an increase in the levels of glucose (12-56%), galacturonic acid (17-53%) and, in two lines, galactose (22%; pCel6.2M and pCel8.3M). Mannose, rhamnose, arabinose and xylose were unchanged. The increased glucose levels of the non-cellulosic fractions may be due to a rise in mixed (1 ,3;1 ,4)^-glucans, that are part of the type II wall hemicellulose fraction (Vergara and Carpita, 2001 ; Souza et al., 2012). Total uronic acids (galacturonic and glucuronic acid) quantity were also elevated in the internodes of the transgenic lines in comparison to the untransformed control plants. Determination of both the amount of lignin and its monomer composition in the transgenic lines showed that soluble and insoluble Klason lignin contents were reduced by between 6-13% and 5-9% in young and mature internodes respectively (Table 3). The reduction was accompanied by a decrease in the lignin's monomer (S and G units) composition lines similar to previous reports where total lignin was reduced (Fu et al., 201 1 ; Ambavaram et al., 201 1 ; Jung et al., 2012).

Taken together, the cell wall composition is greatly altered, with increased cellulose leading to a decrease in the lignin content. This suggests that the cell wall may be more easily degraded by cellulosic enzymes. Untreated lignocellulose biomass of young and mature internodes was exposed to a combination of cellulase and cellobiase. Younger internodal tissues showed the highest saccharification efficiency of between 28-39% and the mature internodes 5-28% in comparison to the wild type control plants. As the transgenic lines in this study contain both increased cellulose as well as decreased lignin, the improved saccharification efficiency might be due to the cellulose being more available, to the decreased lignin allowing improved cellulases contact to cellulose, or to a combination of both. Pairwise linear regression analysis between saccharification efficiency and Klason lignin content of the transgenic lines revealed a negative correlation (R ² = 0.69). An even stronger negative correlation was observed between the saccharification efficiency and the total thioacidolysis lignin content (R ² = 0.93), agreeing with previous studies of alfalfa and switchgrass (Grabber et al., 2002; Chen and Dixon, 2007; Fu et al, 201 1 ). Second generation bioethanol production aims to use non food or feed plants with high yields in biomass and reduced recalcitrance, while limiting the use of toxic chemicals. The increased biomass, reduced lignin content and improved saccharification efficiency of the transgenic sugarcane plants reported here promises to add value to bagasse after sucrose juice extraction. The development of improved sugarcane as an energy source may encourage the application of this technology, which could ultimately lead to an environmentally friendly renewable energy source.

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