Identification and characterization of insect cellulolytic systems for plant biomass degradation

A Final Report Submitted to: The Southeastern Sun Grant Center

Submitted by

Dr. Juan Luis Jurat-Fuentes

Department of Entomology and Plant Pathology University of Tennessee

2431 Joe Johnson Drive, 205 Ellington Plant Sciences Knoxville, TN 37996

Co-PIs

Dr. Cris Oppert (University of Tennessee, Knoxville, TN 37996) Dr. William Klingeman (University of Tennessee, Knoxville, TN 37996)

Dr. Brenda Oppert (USDA-ARS Center for Grain and Animal Health Research, Manhattan, KS 66502)

Project Period, July 2007-July 2011

October 31st, 2011

This project was funded by a grant from the Southeastern Sun Grant Center with funds provided by the United States Department of Energy, Office of the Biomass Program.

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Abstract Recalcitrance of lignocellulosic (plant) biomass greatly reduces cost effectiveness of ethanol biofuel production. Biotechnological improvements are expected to greatly impact lignocellulosic ethanol production and reduce production costs to compete with fossil fuels. Degradation of plant biomass to simple sugars that are then fermented to ethanol is achieved using enzymes called cellulases, which degrade plant cellulose to its glucose subunits. The goal of our project was to identify novel cellulases displaying enhanced properties for application in the lignocellulosic biofuel industry. Our work has been focused on phytophagous insects as prospecting resources because they are highly effective in degrading plant biomass and cellulases in their digestive system have been reported to function under extreme physicochemical conditions. We conducted an extensive cellulolytic activity screen of more than 100 species belonging to 9 taxonomic orders using carboxymethyl cellulose (CMC) and microcrystalline cellulose (MCC) as substrates. From this screening effort, we identified a number of insect species displaying high levels of cellulase activity compared to termites, which have been traditionally considered as examples of highly effective cellulolytic organisms. Characterization of cellulolytic systems in these selected species allowed identification of an endoglucanase displaying highest levels of activity in alkaline pH (pH 11). This unique property suggests that this enzyme may be amenable to use with ionic liquids and other alkaline-based solutions generally used in pretreatment of plant biomass. We also identified a group of insects displaying levels of activity that are 3 to 5 fold higher than any other insect species tested (including termites). To identify the enzymes responsible for this activity, we have validated an expression library screening approach. Identification of novel cellulases with unique properties amenable to biorefineries is expected to greatly reduce lignocellulosic ethanol production costs and increase its competitiveness against fossil fuels.

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Final report • Problem

Cost-efficient production of ethanol biofuel from lignocellulosic biomass is highly dependent on development of efficient hydrolysis technologies (32, 40). Until production costs can be reduced and the activities of cellulolytic enzymes increased, lignocellulosic ethanol production will remain marginalized as a practical fossil fuel alternative. Enzymatic degradation of cellulose is achieved by the sequential activity of three type of enzymatic activities: endoglucanase, exocellulase, and β-glucosidase. This degradative process is considered the hydrolysis method with the greatest potential for improvement and cost reduction (19, 39, 40). Enzymes currently used for production of cellulosic ethanol were obtained from fungal and bacterial systems (18) and present a number of limitations, including low specific activity and susceptibility to product inhibition and environmental conditions. The availability of more active cellulases with activity under stringent conditions would promote the use of these enzymes in consolidated bioprocessing. These circumstances have led to increased interest in cellulase prospecting to identify novel cellulases with increased specific activity and stability, and lower susceptibility to inhibitors (19).

Phytophagous insects are recognized as being highly effective in the degradation of plant biomass, and their digestive enzymes are known to function under stringent physicochemical conditions that can be found in biofuel refineries. Based on the characterization of cellulose degradation in insects, a new functional model of the insect digestive system as a highly effective biorefinery is emerging (7). The insect produces active endocellulases that perform random cuts to expose and release individual cellulose polysaccharide chains. Exocellulases produced by symbiotic fungi and bacteria in the insect gut then cleave two to four units from the ends of the exposed cellulose chains. The products from exocellulase digestion are hydrolyzed to monosaccharides (such as glucose) by β-glucosidases produced by the insect. This sequential and synergistic action of insect and symbiont-derived enzymes results in highly effective biodegradation of lignocellulosic biomass. However, efforts to identify insect cellulolytic systems and their potential biofuel industry applications have been limited and mostly concentrated on termite species (36). To address these limitations, the objectives of our project were to identify highly effective insect cellulolytic systems and to characterize novel insect cellulases and their potential use in lignocellulosic ethanol production. Our long-term goal is to adapt these effective cellulolytic systems to reduce costs and increase yields in lignocellulosic ethanol production.

• Approach

Information on cellulolytic activity in insects is limited to selected species in six taxonomic orders (36). Due to this limitation, our first objective was to identify insect species displaying high cellulolytic activity to allow later detailed characterization of the cellulolytic system responsible for this activity. To accomplish this goal we performed an extensive screening for cellulolytic activity in digestive fluids from more than 60 insect species in 9 taxonomic orders. We used two cellulose substrates with distinct properties in this screening: carboxymethyl cellulose (CMC) and microcrystalline cellulose (MCC). Carboxymethyl cellulose is composed of cellulose polymers modified with an extra carboxymethyl group that renders the molecule soluble in water, while MCC is a native form of tightly packed cellulose chains linked by hydrogen bonds and is water insoluble. Although CMC has traditionally been used to test for cellulase activity mainly because its high water solubility makes it amenable in assays, it is not characteristic of the solubility of native cellulose. While CMC has been used as

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proxy for the existence endoglucanase-like activity, degradation of MCC is definitive of the existence of a complete cellulolytic system. Based on data from this screen, we selected insect species with high cellulolytic activity relative to termite-derived and commercial cellulase samples for further characterization.

In characterizing these highly active cellulolytic systems, our approach was based on the isolation of active cellulases, obtaining protein sequence tags, and PCR cloning of the respective cDNAs. For isolation of active cellulases, we used chromatographic purification combined with cellulase activity assays and zymograms. Active purified cellulases detected by zymograms were sequenced to obtain tags amenable to design PCR primers and cloning of cDNA. Full-length cDNAs encoding for the specific cellulase were used for the expression of an active enzyme in heterologous systems, necessary to obtain information relevant to stability, activity, and specificity. Since the vast majority of insect species tested in our cellulase activity screening were not traditional insect models, we also planned to develop a high throughput assay based on screening of cDNA expression libraries to identify specific cellulolytic enzymes with high activity. Based on the use of yeast for fermentation of simple sugars to ethanol and the importance of glycosylation for activity of some insect cellulases (38), we planned to use yeast as our heterologous expression system. • Methodology Insect collection and dissections Insects evaluated in the cellulase screen were collected from the field in Eastern Tennessee (USA) or New Zealand, except for Tenebrio molitor, Tribolium castaneum, and Heliothis virescens. T. molitor and T. castaneum were obtained from laboratory cultures at the USDA Center for Grain and Animal Health Research (Manhattan, KS). H. virescens eggs were purchased from Benzon Research (Carlisle, PA). Spodoptera (pr. dolichos) egg masses were generously provided by Dr. D. Jenkins (USDA-ARS Tropical Agriculture Research Station, Mayaguez, PR). Insects were actively feeding on or in close proximity to plant host tissues, and were dissected the same day of collection or stored at 4°C for no more than three days until dissections could be performed. Guts and/or heads were dissected from the insects and placed in 50-500 µl molecular biology-grade water (Eppendorf). Depending on sample size, multiple guts or head samples were combined to ensure availability of sufficient material for subsequent assays. Dissected tissues were cut into small pieces, homogenized by vortexing, and centrifuged at 16,100 x g for 3 min at room temperature. Supernatants were transferred to new centrifuge tubes and stored at –80°C.

Determination of cellulolytic activity Proteins in gut or head fluid samples were quantified using the Coomassie protein assay (Pierce) with BSA as standard (4) or a Qubit Fluorometer (Invitrogen, Carlsbad, CA). Cellulolytic activity in digestive fluid extracts was tested against microcrystalline cellulose (MCC, Acros Organics, Geel, Belgium), carboxymethyl cellulose (CMC, Sigma-Aldrich, St. Louis, MO), 4-nitrophenyl β-D-glucuronide (pNPG, Sigma-Aldrich, St. Louis, MO), and pulverized switchgrass (13) as substrates. Activity assays using cellulose polymers (MCC and CMC) or pulverized switchgrass as substrates were performed using a modified 3’-5’dinitrosalicylic acid (DNSA) assay (20, 25). Protein samples (50 µg or 150 µg for the CMC and MCC/switchgrass assays, respectively) were mixed with a 2% (w/v) substrate solution in 50 mM sodium citrate buffer (pH 6.0). Activity assays were conducted in polystryrene 96-well plates at 50°C for 1 hour (CMC) or 2 hours (MCC, and switchgrass). A modified DNSA reagent

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containing Rochelle salt (21) was added to samples to stop enzymatic activity, and color was developed at 100°C for 15 min. Microplates were cooled at room temperature for 5 min and centrifuged at 2000 x g for 2 min to precipitate any remaining substrate. Supernatants (50 µL) were transferred to new 96-well polystyrene microplates used to read absorbance at 595 nm on a Synergy HT microplate reader (BioTek, Winooski, VT) using the Gen5 software (v. 2.0, BioTek, Winooski, VT). The concentration of reducing sugars in each sample was determined using a glucose standard curve and background amounts of reducing sugars were corrected by subtracting final from initial values of the calculated reducing sugars in the sample. One unit of cellulolytic activity was defined as the amount of enzyme that produced 1 µmol of reducing sugar (glucose equivalents) per min at 50°C and pH 6.0. Specific activities were reported as units per mg of protein. All specific activities represented averages from at least three independently pooled samples (i.e. biological replicates). Determination of β-glucosidase activity was accomplished utilizing pNPG as a cellobiose analog (27). The 100 µL reaction mixture contained 25 µL of digestive fluids, 25 µL of 10 mM pNPG, and 100 µL of 50 mM sodium citrate buffer at pH 6.0. Samples were incubated for 30 min at 50°C in a dark incubator, and then the reaction was terminated by the addition of 100 µL of 400 mM NaOH-glycine buffer (pH 10.8) to allow for color development. For quantification of β-glucosidase activity, a standard curve of ρ-nitrophenol (10-100 nmol mL-1) was used (29), and absorbance of the standard curve and reaction mixture was measured at 405 nm in a Synergy HT microplate reader (BioTek, Winooski, VT) using the Gen5 software (v. 2.0, BioTek, Winooski, VT). Background amounts of reducing sugars were corrected by subtracting final from initial values of the calculated reducing sugars in the sample. Cellulase zymography Zymograms to detect cellulolytic activity bands were performed as described elsewhere (31), with minor modifications. Gels were prepared by including 0.1% CMC before polymerization in a SDS-10%PAGE resolving gel mixture. Gel mixtures were heated to 30°C while CMC was added slowly to prevent aggregation. After all CMC was dissolved, APS and TEMED were added and gels were allowed to polymerize overnight at room temperature. Commercial grade Aspergillus niger cellulase (MP Biomedicals, 1 mg) was used as a positive control. Samples (40 µl) were partially denatured at 70°C for 20 min to decrease activity band smearing. Following heating, samples were briefly centrifuged and loaded in gels. Proteins in samples were separated by SDS-10%PAGE at a constant 100 V at 4°C for approximately 4 h or until dye reached the bottom of the gel. After electrophoresis, gels were washed five times at room temperature each wash for 30 min with 50 ml of wash buffer (0.1 M sodium succinate, 10 mM DTT, pH 5.8; the last wash was at 30°C). Remaining CMC in the gel was stained by incubation in a solution of 0.1% Congo Red for 10-15 min at room temperature. Gels were destained by washing in 50 ml of 1 M NaCl until cellulase bands became visible as clear areas where CMC had been degraded due to enzymatic activity. After destaining for 20 min, 100 µl of glacial acetic acid was added to the gel wash for improved band visualization (34). Following this treatment, gels turned dark-purple in color with activity bands remaining as clear zones. Images of gels were acquired using a Versadoc 1000 Imager (Bio-Rad), and pictures were inverted and enhanced using Adobe Photoshop CS2 software (v. 9.0.2). Statistics Data from Dictyoptera, Dermaptera, Diptera, Isoptera, and Hymenoptera (head fluid only) were not used in statistical comparisons among orders due to the limited number of species

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tested from these orders. Enzyme activity data from Coleoptera, Hymenoptera (gut fluid only), Lepidoptera, and Orthoptera were factored by Order and substrate type, or by Order and tissue type, using two-way ANOVAs with the Holm-Sidak method for post-hoc pairwise multiple comparisons (overall α=0.05). Non-normal data were analyzed using the Kruskal-Wallis test. Comparisons between enzymatic activities from diverse species were made using one-way ANOVAs with either the Holm-Sidak (H-test) or Dunn’s (F-test) method for post-hoc pairwise multiple comparisons (overall α=0.05; Gardiner and Gettinby, 1998). Statistical analyses were performed using SigmaPlot for Windows (v. 11.0; Systat Software). Data are presented as means +/- standard error of the mean (SEM), unless otherwise indicated. Cloning and construction of TcEG1 expression cassette Sixth-instar T. castaneum larvae were collected and anesthetized on ice for 10 min before dissection. Head capsules, which contain salivary and labial glands were isolated and placed into individual micro-centrifuge tubes (10 per tube) with 100 µl of RNAlater (Ambion). Tubes containing head tissues were incubated at 4 °C overnight to allow RNAlater to infiltrate and preserve the RNA. Tubes were quick-spun and RNAlater removed, followed by flash freezing and mortar pestle grinding. RNA was extracted using the RNeasy kit (Qiagen) following manufacturer’s instructions, and the product was evaluated on a 1% agarose gel and stored at -80 °C until used. Concentrations of RNA were estimated using the Quant-iT RNA assay kit (Invitrogen) following manufacturer’s instructions. Superscript III reverse transcriptase (Invitrogen) was used to synthesize cDNA from RNA collected from head. Primers for TcEG-1 were designed for amplification of the full-length cDNA sequence and to assist with proper insertion into the pIZT/V5-His (Invitrogen) expression vector so that a 6xHis tag was added to the C-terminus of the expressed protein. Cloning was facilitated by EcoRI and NotI restriction enzyme sites engineered in primers used for amplification (underlined): TcpIZTFwd 5’-GGAATTCGATGTTCTACTCATTGTGGGTGCTACTATTT-3’ and TcpIZTRev 5’- ATAGTTTAGCGGCCGCCAATTTATTCTCATTTTCAATATAAAT-3’. Accuprime pfx supermix (Invitrogen) was used for PCR reactions containing 5% DMSO, 20 mmol of each primer, and 1 µl of cDNA. The PCR protocol consisted of an initial 5 min at 95 °C, followed by 30 cycles of 30 sec at 95 °C, 30 sec at 60 °C, 90 sec at 68 °C, and a final 10 min extension at 68 °C. The PCR products were cloned into TOPO TA (Invitrogen) and constructs transformed into E. coli TOP10 competent cells (Invitrogen). Positive clones were determined by sequencing transformants in forward and reverse directions (University of Tennessee Sequencing Facility). Both the native and His-tagged TcEG1 inserts in the TOPO plasmid were excised using EcoRI and NotI digestion and gel-purified (QIAquick Gel Extraction Kit, Qiagen). Purified inserts were ligated using T4 DNA ligase (Invitrogen) into pIZT/V5-His to generate the pIZT/V5/TcEG1-His expression cassette. Constructs were transformed into E. coli TOP10 cells (Invitrogen) and positive transformants as well as reading frames were confirmed by sequencing in forward and reverse directions . Plasmid for insect cell transfection was produced and purified using a HiPure Plasmid Maxiprep kit (Invitrogen). Sequence alignment and phylogenetic analysis The TcEG1 sequence (Glean 15370) was acquired from the T. castaneum genome (22) and was translated by ExPASy Translate and analyzed with the Compute pI/MW tool to predict isoelectric point (pI) and molecular weight (MW) of the predicted protein (http://expasy.org/tools). DictyOGlyc 1.1 (http://www.cbs.dtu.dk/services/DictyOGlyc) and NetNGlyc 1.0 (http://www.cbs.dtu.dk/services/NetNGlyc) software packages were used to

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predict O- and N- glycosylation sites, respectively. Signal peptide prediction was performed using SignalP 3.0 (2) (http://www.cbs.dtu.dk). For phylogenetic comparisons, we performed a BLASTP search (1) of the NCBInr database (accessed September 15, 2010) using the TcEG1 protein sequence as query, limiting by organism group to the Insecta Class to find insect homologs of this protein. Sequences with e values <e-100 (corresponding to about 54% sequence identity) were selected for alignment and tree construction. Predicted endogenous cellulase sequences from coleopteran species in the NCBInr database that were not returned by the BLASTp search (Psacothea hilaris, Apriona germari and Phaedon cochlearia) were also included in sequence alignments for phylogenetic and molecular evolutionary analyses. Retrieved sequences were aligned with the predicted TcEG1 protein sequence for construction of a maximum parsimony phylogenetic tree using MEGA version 4 (33). For prediction of the three dimensional (3D) structure of TcEG1, we search structural databases using the Phyre server (http://www.sbg.bio.ic.ac.uk/~phyre/index.cgi) to identify accurate models (11). The cellulase from Nasutitermes takasagoensis (Accession number BAA33708.1) was identified as optimum model and used to determine alterations in protein structure due to a specific Gly to Ser change observed in the TcEG1 sequence compared to other GHF9 members. Prediction of phosphorylation of the 70Ser residue in TcEG1 was performed using Phos 2.0 (www.cbs.dtu.dk/services/NetPhos/). Transient expression of TcEG1 in Drosophila S2 cell cultures Cultured Drosophila S2 cells were maintained in serum-free insect cell medium (HyQ SFX-Insect, Hyclone) and transfected as previously described (9). Briefly, approximately 2×106 S2 cells from a confluent culture were suspended in 2 ml fresh media and allowed to adhere overnight to surface-treated 60 × 15 mm polystyrene dishes (Falcon). Plasmid transfection mixtures were prepared by mixing either 4 µg of pIZT, or 5 µg of pIZT/V5/TcEG1-His- plasmids with 1 ml of serum-free insect medium and 20 µl of Cellfectin reagent (Invitrogen). Transfection mixtures were incubated with cells at 20 °C for 4 h, after which fresh media (3 ml final volume) was added to the plates. Cell cultures were incubated at 26 °C for three days before collection and centrifugation at room temperature (6,000 x g for 3min). Cell pellets and supernatant media were retained to test for TcEG1 expression. Supernatant media was concentrated approximately 10-fold using a SpeedVac (Thermo Scientific). The S2 cell pellets were lysed by resuspension in native purification buffer (25 mM NaH2PO4 pH 8.0, 250 mM NaCl) and two freeze-thaw cycles. After lysis, samples were cleared by centrifugation as above and supernatants collected and stored at -80 °C until used. Detection of TcEG1-His expression Expression of His-tagged TcEG1 in S2 cell cultures was detected by Western blotting using antisera against the 6xHis tag. Protein concentrations in S2 cell lysate and media supernatant were estimated using the Coomassie Protein Assay Reagent (Pierce) following manufacturer’s instructions using bovine serum albumin (BSA) as standard. Proteins (1 µg) in S2 cell lysate or media supernatant were solubilized in 2X sample buffer (15) and heat denatured for 20 min. Proteins were separated by electrophoresis on SDS-10%PAGE gels, then electro-transferred to polyvinylidene difluoride (PVDF) filters overnight at 4 °C under constant voltage (20 V). Filters were blocked with 3% BSA in PBST (135 mM NaCl, 2 mM KCl, 10 mM Na2HPO4, 1.7 mM KH2PO4, pH 7.5, 0.1% Tween-20) for 1 h at room temperature. After blocking, filters were probed with a 1:10,000 dilution of antisera (Novus Biologicals) against 6xHis tag conjugated to horseradish peroxidase (HRP) for 1 h in blocking buffer. After several

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washes with PBST and 0.1% BSA, blots were developed with the Supersignal West Pico chemiluminescence substrate (Pierce). Enzymatic activity: thermostability and pH optima Cellulase activity in digestive fluids or media and cell pellets from S2 cell cultures expressing TcEG1 and mock-transfected cell cultures under diverse temperature and pH conditions was determined using the DNSA assays as described above. Protein samples (50 µg and 150 µg for CMC and MCC assays, respectively) or glucose standards (ranging from 0 to 313 µg) were added to substrate solutions (2% w/v in 50 mM sodium citrate buffer, pH 6.0) and incubated for 1 h (CMC) at 50 °C. For thermal stability assays samples (25 µg total protein) were incubated at 30 °C, 40 °C, 50 °C, 60 °C, or 70 °C for 10 min before testing in DNSA assays as described above using CMC as substrate. The effect of pH on cellulase activity was measured by incubating samples (10 µg) in 2% (w/v) CMC dissolved in buffer at pH 2.0 (50 mM maleic acid), pH 5.0 (50 mM citrate buffer), pH 7.0 (50 mM phosphate buffer), pH 8.5 (50 mM glycine buffer), pH 9.0 (50 mM Tris-HCl), or pH 12 (50 mM sodium phosphate). Degradation of CMC at 50 °C for one hour was detected and measured as above. All absorbance readings represent averages from at least three experiments with material from independent recombinant enzyme or larval batches (biological replicates) performed in triplicate (technical replicates). Statistical significance was tested through one-way analysis of variance (ANOVA) and pairwise multiple comparison procedures (Holm-Sidak method, overall α = 0.05) using the SigmaPlot v11.0 software (Systat Software IN., IL).

Purification of cellulolytic enzymes We used a two-step liquid chromatography strategy to purify cellulolytic enzymes. Pooled fluids (25-45 ml) were diluted to 50 ml with 0.01 M sodium acetate buffer pH 4.9 (buffer A). Samples were cleared by centrifugation (5 min at 21,000 x g) and filtered (0.22 µm) before loading on a HiLoad 16/60 Superdex 200 prep grade gel filtration column previously equilibrated with buffer A and connected to an AKTA FPLC system (GE Healthcare). Proteins in the sample were eluted with 240 ml of buffer A at constant flow (1 ml min-1), with continuous collection of 2 ml fractions. To test for presence of cellulolytic activity in the collected FPLC fractions, we used 1% agarose plates containing 0.2% CMC. An aliquot (20 µl) of each fraction was spotted on the plate and allowed to digest the CMC substrate at 30°C for 2 h. After incubation, plates were stained with 0.1% Congo red and destained with 1 M NaCl. Cleared spots revealed fractions containing active cellulase protein. For gut fluids, cellulase activity was detected in the 65 to 120 mL elution volume, while for head-derived fluids cellulase activity was detected in the 75 to 92 mL elution volume. Fractions within the detected peak of activity were tested for activity in zymograms or pooled and used for anion exchange chromatography with a Mono Q 5/50 GL anion exchange column equilibrated with buffer C (50 mM Bis-Tris, pH 4.8). Prior to anion exchange chromatography, pooled fractions were dialyzed (10,000 MWCO) overnight against buffer C. After loading the pooled sample in the column, proteins were eluted using a 50% gradient over five column volumes of buffer D (buffer C plus 1 M NaCl). Fractions were tested for cellulolytic activity using the CMC-agarose plate assay, as described above. Under these conditions, cellulases from gut and head-derived fluids were eluted from the column with less than 10% NaCl. Fractions containing cellulolytic activity were diluted three-fold in buffer A, then used for a second anion exchange run using a 50% gradient of buffer D, but over two

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column volumes. Fractions with cellulolytic activity, as determined by the CMC-agarose plate assay, were pooled and concentrated approximately 100-fold using a speed vacuum (Savant, Farmingdale, NY). Enzyme sequencing and identification Purified cellulase samples were analyzed using zymograms to test for the presence of active cellulase (SDS-8%PAGE, 0.2% CMC). For N-terminal sequence tags, purified cellulase bands were separated by SDS-10%PAGE and electrotransferred to a polyvinylidene fluoride (PVDF) membrane overnight. After transfer, the membrane was stained for total protein using Coomassie brilliant blue, and bands corresponding to active cellulases were excised and submitted for N-terminal protein sequencing (Iowa State Protein Facility, Ames, IA). Amino acid sequences were queried to the NCBInr and Swiss-Prot databases (March 24, 2010) using BLASTp.

For protein identification using nano liquid chromatography tandem mass spectrometry analysis (LC/MS/MS), partially purified cellulase protein samples from fractions displaying activity were subjected to zymography (SDS-10%PAGE, 0.2% CMC). Following zymography, gels stained for total protein using ProtoBlue Safe coomassie stain (National Diagnostics, Atlanta, GA) were aligned to zymograms by comparing prestained molecular size marker ladders, allowing for the excision of gel regions containing protein bands corresponding to cellulolytic activity. Three gel regions were removed and submitted to MS Bioworks (Ann Arbor. MI) for LC/MS/MS analysis. Briefly, proteins in the gel slices were extracted and digested with trypsin (Promega) at 37°C for 4h, then the generated peptides analyzed in a LTQ Orbitrap Velos mass spectrometer (ThermoFisher). Mass spectrometry data was searched using the Mascot software (Matrix Science, London, UK) to query against the NCBInr database (accessed February 19th, 2011). Parameters used in the Mascot searches included allowance for a maximum of 2 missed cleavages; iodoacetamide derivative of cysteine as a fixed modification; S-carbamoylmethylcysteine cyclization (N-terminus) of the n-terminus, deamidation of asparagine and glutamine, oxidation of methionine, and acetylation of the n-terminus were specified as variable modifications. Parent ion tolerance was set to 10.0 ppm and fragment ion mass tolerance to 0.50 Da. Scaffold (version Scaffold_3_00_07, Proteome Software Inc., Portland, OR) was used to validate MS/MS based peptide and protein identifications. Peptide identifications were accepted if they could be established at greater than 50.0% probability as specified by the Peptide Prophet algorithm (10). Protein identifications were accepted if they could be established at greater than 80.0% probability and contained at least 1 identified peptide. Protein probabilities were assigned by the Protein Prophet algorithm (24). Proteins that contained similar peptides and could not be differentiated based on MS/MS analysis alone were grouped to satisfy the principles of parsimony. Cloning of TcEG1 into expression plasmid p426-GPD The full length cDNA encoding TcEG1 was cloned from the existing insect cell expression construct, using the polymerase chain reaction (PCR) protocol described below. To include BamH1 and EcoR1 restriction sites at the 5’ and 3’ ends, respectively, we used forward primer 5’-CGGGATCCATGTTCTACTCATTGTGGGTGCTAC-3’ and reverse primer 5’- CGGAATTCTCACAATTTATTCTCATTTTCAATA-3’, wherein the restriction sites are underlined. The optimized PCR protocol included an initial denaturation step of 2 minutes at 92°C followed by 30 cycles of 1 minute denaturation at 92°C, annealing for 30 seconds at 63°C, and elongation for 1 minute at 72°C. Resulting PCR products were separated on a 1% agarose gel, purified, and subjected to restriction digestion using BamH1 and EcoR1 restriction enzymes

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(Fermentas, Vilnius, Lithuania). These restriction sites allowed easy, unidirectional insertion of amplified cDNA sequences into the multiple cloning site of the yeast expression vector p426-GPD (23). After digestion for 1 h, cDNA and p426-GPD were run on a 1% Agarose/EtBr gel and purified using a QIAquick gel extraction kit (Qiagen, Hilden, Germany). Purified cDNA was then ligated into the digested p426-GPD plasmid using T4 DNA ligase (Fisher Scientific, Atlanta, GA) to yield the p426-GPD-TcEG1 construct. This construct was amplified in E. coli using the QIAprep spin maxiprep plasmid purification (Qiagen, Hilden, Germany). Purified p426-GPD-TcEG1 plasmid was the quantified using a Qubit fluorometer (Invitrogen, Carlsbad, CA) and stored at -20⁰C until needed for yeast transformation. Transformation of yeast with expression plasmid p426-GPD-TcEG1 Purified plasmid p426-GDP-TcEG1 was used to transform the TRY127 yeast strain using an established lithium acetate transformation protocol (8). A single yeast colony was selected and grown in 5 mL of yeast peptone dextrose (YPD) broth overnight at 30°C with shaking (125 rpm). After 24 hours, cultures were diluted into 25 mL YPD to an OD600 of 0.1, and then grown at 30°C for 4 h while shaken at 150 rpm. Cells were harvested by centrifugation (5,000 x g for 2 min at 4⁰C), and the pellet washed with 25 mL of sterile distilled water. Washed pellets were then resuspended in 500 µL of LiOAc/TE buffer (10X TE [1M Tris, 0.5M EDTA, pH 8.0], 1M LiOAc), transferred to a microfuge tube, and then re-pelleted at 13,000 x g for 1 minute. The supernatant was removed and the pellet resuspended by vortexing in 225 µL of fresh LiOAc/TE buffer. An aliquot (50 µL) of the resuspended cell sample was transferred to a microcentrifuge tube, and 5 µL of single stranded DNA (Herring Sperm DNA, Promega) was added as a carrier DNA strand. Next, 15 µL of the expression plasmid p426-GPD-TcEG1 was added to the tube along with 300 µL of LiOAc/TE/PEG buffer (10X TE, 1M LiOAc, 20g PEG). Following vortexing, the tube was incubated for 30 min at 30°C before being transferred to a 42°C water bath for 20 min. The reaction mixture was then pelleted at 8,000 x g for 1 min, and the supernatant aspirated. The cell pellet was then resuspended in 200 µL of sterile distilled water and plated on minimal media agar lacking uracil (MLU agar). To select successful transformants MLU agar plates were used, as only transformant yeast cells would be capable of growing in the absence of uracil. To additionally test for the expression of an active TcEG1 cellulase, glucose was substituted in the MLU agar composition with 0.2% carboxymethyl cellulose (MLU-CMC). Thus, only successful yeast transformants would be able to grow by hydrolyzing CMC as the only available carbon source. As negative control, we used yeast transformed with the empty expression vector p426-GPD. After streaking, plates where inverted and incubated at 30ºC for 7 d. Plates were removed at the end of the 7 d growth period and photographed against an illuminated white background using a Canon Rebel Ti DSLR camera (Canon, Tokyo, Japan), to document growth. Expression and partial purification of TcEG1 from yeast cultures A single isolated colony picked from an MLU-CMC plate was inoculated into an Erlenmeyer flask containing 50 mL of sterile YPD broth. As negative control, we used a single colony of TRY127 transformed with the empty p426-GPD expression vector. Inoculated flasks were incubated at 30°C for 7 d in a shaking incubator (175 rpm). Following incubation, cellular material and debris were cleared by centrifugation (5,000 x g for 5 minutes at 4⁰C). The supernatant (~50-mL) was placed in a 30K MWCO protein concentrator (Omnicron, Billerica, MA) and used to concentrate protein following manufacturer’s protocols. The concentrated sample (~2 mL) was quantified using a Qubit Fluorometer (Invitrogen, Carlsbad, CA) before storage at 4°C until used (no longer than one wk).

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• Findings Screening for cellulase activity in insect digestive fluids

Structural differences of the two substrates used to detect cellulolytic activity revealed different levels of enzymatic activity in head and gut fluid samples from the insect species tested. Degradation of CMC functions as a proxy for EG activity, while MCC conversion to glucose approximates total cellulolytic activity (EG, CBH, and β-glucosidases). Cellulolytic activity from gut fluids on CMC and MCC substrates was detected for a total of 63 and 56 species, respectively (Fig. 1). MCC activity was determined for a lower number of species due to sample size limitations. Activity was found in all species tested, except for Epargyreus clarus and Spodoptera (pr. dolichos), which yielded no detectable activity against CMC. There was a high degree of variability among samples, which may be due to differences in feeding activity or asynchrony at time of dissection. The highest gut fluid activity against MCC (i.e. greater than 0.05 U per mg of protein) was observed for species among the Coleoptera, Hymenoptera, Lepidoptera and Orthoptera orders (Fig. 1A). In contrast, the highest gut fluid cellulolytic activities with the CMC substrate (i.e., greater than 0.5 U per mg of protein) were obtained for species among the Dictyoptera, Coleoptera, Isoptera, and Orthoptera orders (Fig. 1B). Lepidopteran activity against CMC in gut fluids was significantly lower than in Coleoptera or Orthoptera (Fig. 1B; F3,113=21.78, P<0.001). Gut fluid samples from the representative termite and cockroach samples (Reticulitermes hageni and Cryptocercus ssp. respectively) displayed high activity against CMC, but significantly lower activity against MCC (R. hageni, H1=7.53, P=0.004; Cryptocercus sp., F1,5=9.25, P=0.038). We detected a number of species with activity against both CMC and MCC comparable to samples from termites and cockroaches. Scolytus (pr. rugulosus) and Anisota virginiensis had the highest gut fluid activities for CMC and MCC, respectively. The only dermapteran species measured, Forficula auricularia, had low activity against both CMC and MCC.

Only 22 (CMC) or 18 (MCC) species were measured for head fluid activity due to limitations of sample size. When comparing among orders, the relative levels of cellulolytic activity for head fluids were slightly different to the relative activity level patterns observed with gut fluids. Enzymatic activities in head fluids against CMC and MCC were not significantly different among orders (F5,32=0.44, P=0.815). Even though gut fluid activities in larval Synanthedon scitula were relatively low (Fig. 1) the highest measured activities in our study were from head fluids of this insect, although this level of activity was not statistically different from other head samples (P>0.05 for all pairwise comparisons). The highest cellulolytic activities in R. hageni were detected in gut fluids against CMC, which were significantly higher than head fluids on either CMC or MCC as well as gut fluids on MCC (F1,16=9.56, P=0.01). Using zymograms with CMC as substrate we were able to confirm diversity in size and number of cellulases in digestive fluids from diverse insect species. This diversity was also observed for species within taxonomic orders. However, we were unable to determine the origin (endogenous versus symbionts) of these enzymes. The data from this study represents the most comprehensive survey of cellulase activity in insects and is a valuable resource for future studies of insect enzymes as well as biorefinery applications. Characterization of cellulolytic activity in specific insect species

Among the screened insect species Dissosteira carolina was selected for further analysis based on its local abundance and important cellulase activity levels. We first determined that D. carolina cellulases were expressed during all the life stages of the insect, and that they were

12

localized to the foregut and midgut regions, suggesting an endogenous origin. Using liquid chromatography we purified a cellulase activity band which in proteomic analyses was found to contain peptides matching to insect cellulases. One of the enzymes had high identity to a cricket endoglucanase, suggesting conservation of cellulolytic enzymes within taxonomic Orders.

We also found two additional D. carolina cellulases of 43-kDa (from head-derived fluid extracts) or 45 kDa (from gut fluid extracts) that were not homologous to database proteins, suggesting they may represent novel cellulases. Interestingly, all the enzymes matching the sequence tags obtained from purified D. carolina cellulases belong to GHF9. Based on this homology and the localization of these enzymes to the anterior gut regions, which in Orthoptera contain very limited microbial flora (5), we concluded that the identified cellulases are probably endogenous enzymes. This conclusion is further supported by the homology observed for this enzyme to endogenous insect cellulases, and absence of this enzyme from the hindgut region, which is a reservoir for symbiotic flora and enzymes (5). These data represent the first report identifying specific cellulases in the gut from an insect in the Acrididae Family. Acrididae contains numerous grasshopper pest species that confound management of crop commodities worldwide, including many being considered as feedstock for lignocellulosic ethanol biofuel (14, 26). In this regard, we advanced D. carolina as a model insect exhibiting effective cellulolytic activity and will use this model to prospect for novel cellulolytic enzymes leading to more efficient biofuel production.

13

0.00.10.20.3Cryptocercus (pr. punctulatus)Cerambycid sp.

Elaphidion mucronatumGraphognathus leucolomaLeptinotarsa decemlineataLyctus (pr. planicollis)Neoclytus acuminatusPopillia japonicaPhyllophaga sp.Scolytus (pr. rugulosus)Tenebrio molitorTribolium castaneumForficula auriculariaMonarthopalpus flavusAllantus cinctusCladius difformisMacremphytus tarsatusNeodiprion leconteiReticulitermes (pr. flavipes)Agraulis vanillae

Anisota virginiensisAtteva punctellaBattus philenorDanaus plexippusDatana contractaDatana integerrima

Fascista cercerisellaHalysidota tessellarisHeliothis virescensHemaris diffinisHomadaula anisocentraHyphantria cuneaJunonia coeniaMalacosoma americanaNorape ovinaOmphalocera munroeiPsilocorsis cryptochiellaSaucrobotys futilalisSpodoptera (pr. dolichos)Synanthedon exitiosaSynanthedon scitulaThyridopteryx ephemeraeformis

Allonemobius (pr. socius)Chortophaga viridifasciata

Dicromorpha viridisGryllus (pr. pennsylvannicus)Hippiscus oceloteMelanoplus differentialisMelanoplus femurrubrum

Neoconocephalus triopsOrchelimum vulgareSchistocerca americanaSchistocerca damnificaScudderia (pr. curvicauda)Scudderia furcata

Syrbula admirabilisColeopteraHymenopteraLepidopteraOrthoptera

Di ct yo pt er a

Col

eopt

era

D er m ap ter a

Di

pt er aH

ymen

opt

era

Is op ter a

Lepi

dopt

era

Orth

opte

raO

rder

M

eans

U mg-1

0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8Cryptocercus (pr. punctulatus)

Cerambycid sp.Chrysobothris sp.

Elaphidion mucronatumGraphognathus leucolomaLeptinotarsa decemlineata

Lyctus (pr. planicollis)Neoclytus acuminatus

Phyllophaga sp.Popillia japonica

Scolytus (pr. rugulosus)Tenebrio molitor

Tribolium castaneumForficula auricularia

Monarthopalpus flavusAllantus cinctus

Cladius difformisMacremphytus tarsatus

Neodiprion leconteiReticulitermes (pr. flavipes)

Agraulis vanillaeAnisota senatoria

Anisota virginiensisAtteva punctellaBattus philenor

Danaus plexippusDatana contracta

Datana integerrimaEpargyreus clarus

Fascista cercerisellaHalysidota tessellaris

Heliothis virescensHemaris diffinis

Homadaula anisocentraHyphantria cunea

Junonia coeniaMalacosoma americana

Norape ovinaOmphalocera munroei

Psilocorsis cryptochiellaSaucrobotys futilalis

Spodoptera (pr. dolichos)Synanthedon exitiosa

Synanthedon scitulaThyridopteryx ephemeraeformis

Tortricid (pr. Archips sp.)Allonemobius (pr. socius)

Chortophaga viridifasciataConocephalus strictus

Dicromorpha viridisGryllus (pr. pennsylvannicus)

Hippiscus oceloteMelanoplus differentialis

Melanoplus femurrubrumMicrocentrum retinerve

Neoconocephalus triopsOrchelimum vulgare

Schistocerca americanaSchistocerca damnifica

Scudderia (pr. curvicauda)Scudderia furcata

Spharagemon bolliSyrbula admirabilis

ColeopteraHymenoptera

LepidopteraOrthoptera

Di ct yo pt er a

Col

eopt

era

D er m a pt er a

Di

pt er aH

ymen

opt

era

Is o pt er aLe

pido

pter

aO

rthop

tera

Ord

er

Mea

ns

U mg-1

Dictyoptera

Coleoptera

DipteraHymenoptera

Isoptera

Lepidoptera

Orthoptera

*

*

*

*

*

A B

Order Means

Cryptocercus (pr. punctulatus)Cerambycid sp.

Chrysobothris sp.Elaphidion mucronatum

Graphognathus leucolomaLeptinotarsa decemlineata

Lyctus (pr. planicollis)Neoclytus acuminatus

Phyllophaga sp.Popillia japonica

Scolytus (pr. rugulosus)Tenebrio molitor

Tribolium castaneumForficula auricularia

Monarthopalpus flavusAllantus cinctusCladius difformis

Macremphytus tarsatusNeodiprion lecontei

Reticulitermes hageniAgraulis vanillaeAnisota senatoria

Anisota virginiensisAtteva punctellaBattus philenor

Danaus plexippusDatana contracta

Datana integerrimaEpargyreus clarus

Fascista cercerisellaHalysidota tessellarisHeliothis virescens

Hemaris diffinisHomadaula anisocentra

Hyphantria cuneaJunonia coenia

Malacosoma americanaNorape ovina

Omphalocera munroeiPsilocorsis cryptochiella

Saucrobotys futilalisSpodoptera (pr. dolichos)

Synanthedon exitiosaSynanthedon scitula

Thyridopteryx ephemeraeformisTortricid (pr. Archips sp.)Allonemobius (pr. socius)Chortophaga viridifasciata

Conocephalus strictusDicromorpha viridis

Gryllus (pr. pennsylvannicus)Hippiscus ocelote

Melanoplus differentialisMelanoplus femurrubrumMicrocentrum retinerveNeoconocephalus triops

Orchelimum vulgareSchistocerca americanaSchistocerca damnifica

Scudderia (pr. curvicauda)Scudderia furcataSpharagemon bolliSyrbula admirabilis

ColeopteraHymenopteraLepidopteraOrthoptera

Dermaptera

*

*

Figure 1. Average specific cellulolytic activities (units per mg protein with standard error bars) of gut fluids using (A) MCC and (B) CMC as substrates. Asterisks denote missing data due to low sample size.

14

Figure 2. Localization of activity against CMC in gut regions of adult Dissosteira carolina. (A) Dissection of adult D. carolina digestive system with labeled regions of interest. (B) Quantitative determination of activity against CMC using a DNSA assay as described. Proteins from fluids derived from each region of the digestive tract (foregut, midgut, and hindgut, 50 µg of protein per sample) were allowed to digest CMC for 60 minutes. Means (±S.E.) are provided from three biological replicates. Columns labeled with different letters indicate levels that are significantly different (Tukey’s MRT P<0.05). (C) Active cellulase detection in fluids derived from the three regions of the digestive tract of D. carolina adults. Proteins in digestive fluids (100 µg per lane) were separated using zymograms (SDS-12% PAGE gels containing 0.2% CMC in the resolving phase). After electrophoresis cellulases were detected by staining for CMC. Clear bands were detected in areas were CMC was digested (active cellulase activity). An alternative species displaying relatively high levels of cellulolytic activity was the firebrat (Thermobia domestica). Characterization of cellulolytic enzymes from this insect through chromatographic purification and mass spectrometry resulted in identification of a number of cellulases with high homology to insect cellulases, especially enzymes from termite species. Firebrats are avid consumers of paper and other cellulose-rich materials, and the levels of activity detected in our screen demonstrate that the cellulolytic systems in these insects may contain endoglucanases that may be of use in degradation of cellulosic biomass. Through collaboration with Dr. Sean Marshall and Dr. Trevor Jackson of AgResearch New Zealand, we were able to include digestive fluids from larvae of five beetle species feeding on lignocellulosic material. These beetle larvae (grubs) present a highly specialized hindgut region in which cellulolytic and fermentative activities are localized (7). No activity against MCC or filter paper was detected for any of the samples tested. In contrast, activity against CMC was detected in all samples and in some cases at relatively high levels. In agreement with the role of the hindgut as fermentation chamber, the highest levels of activity in most tested samples were localized in the hindgut region. Only the huhu grub (Prionoplus reticularis), presented high levels of activity in all three gut regions (foregut, midgut, and hindgut), with the midgut containing the highest levels. This species presented the highest levels of cellulase activity among the species from New Zealand. This coleopteran is the largest endemic beetle found in New Zealand, and belongs to the longhorn beetle family (Cerambycidae). Members of this family have been reported to contain diverse cellulolytic enzymes, as well as enzymes that digest lignin (6, 43). This observation agrees with the feeding habits of huhu beetle grubs, which

15

feed exclusively on decaying logs. Our ongoing collaboration with the AgResearch group is focused at identifying enzymes involved in lignin and cellulose degradation in these grubs using next generation sequencing. Cloning, expression, and characterization of TcEG1

The availability of sequenced insect genomes has allowed for discovery and functional characterization of novel genes and proteins. Using the Tribolium castaneum (red flour beetle) genome, we identified, cloned, expressed, and characterized a novel endo-β-1,4-glucanase we named TcEG1 (T. castaneum EndoGlucanase 1). This enzyme has homology to enzymes in glycosyl hydrolase family 9 (GHF9), although it contains a change (Gly for Ser) in the conserved catalytic domain for GHF9 cellulases. Heterologous expression of TcEG1 in Drosophila S2 cell cultures resulted in secretion of a 51 kDa protein, as determined by Western blotting (Fig. 3), which we used to characterize TcEG1 enzymatic activity against CMC and MCC.

Figure 3. Expression of TcEG1 in Drosophila S2 cell cultures and detection of cellulolytic activity. A) Detection of TcEG1 in media supernatant (Media) but not cell pellets (Cells) from transfected S2 cell cultures. Lanes 1 are mock transfected cultures, lanes 2 are transfected with expression vector containing TcEG1. Expressed TcEG1 was detected by probing with antisera against the C-terminal 6xHis tag on TcEG1. Molecular weight markers (in kDa) are presented on the side of the image for TcEG1 size estimation. B) Comparison of activity against CMC or MCC cellulose substrates in same samples used for A. Data for each experiment represents the mean and standard deviation from three independent determinations of CMC degradation from three independent biological samples. Bars display mean values and standard errors calculated from three biological replicates. Different letters on top of columns represent significant differences (t-test, P < 0.05).

After detecting activity against CMC in TcEG1 secreted from S2 cells, we characterized thermostability and pH optima of this activity. In the thermostability assays, activity against CMC in TcEG1 was highest at 50 °C, then sharply decreased as temperatures were increased (Fig. 4A). Similar thermostability data have been reported for other cellulases from coleopteran (16, 17, 30, 37) and orthopteran (12) species, while lower thermostability has been reported for some termite cellulases (41). Although TcEG1 activity against CMC was almost negligible at pH 2, the activity increased from pH 5 to 8.5, the highest pH tested for these samples (Fig. 4B). By contrast, insect cellulases characterized in other reports have displayed highest activities at more acidic pH levels, often between pH 4 and 6 (12, 16, 17, 37). An endoglucanase from

A B

16

Aulacophora foveicollis (Lucas) had a slightly alkaline (pH 7.8) pH optimum (30), and a cellulase from C. formosanus retained 70% activity at pH 9 (41). To our knowledge, the high relative activity at pH 8.5 found for TcEG1 is unique among described insect cellulases, which may indicate greater molecular stability (3). This feature of TcEG1 may have potential utility for development of industrial enzymes that would efficiently hydrolyze lignocellulosic biomass at alkaline pH (28, 35). For example, ionic liquids with high pH conditions have been suggested for accelerated lignocellulose degradation (42), but currently available cellulolytic enzymes are not stable under these conditions. Further testing is needed to determine TcEG1 functionality under different biorefinery conditions, as well as to identify specific regions of this enzyme amenable to engineering for optimal performance in biorefineries.

Temperature (oC)20 30 40 50 60 70 80

Abso

rban

ce (5

95 n

m)

0.05

0.10

0.15

0.20

0.25

0.30

0.35

0.40A

pH0 2 4 6 8 10

Abs

orba

nce

(595

nm

)

0.00

0.02

0.04

0.06

0.08

0.10

0.12

0.14B

Figure 4. Determination of thermostability and pH optima for TcEG1. A) Thermostability of activity against CMC substrate in crude extracts of native or His-tagged TcEG1 (TcEG1 and TcEG1-His, respectively). B) Dependence of activity of TcEG1 toward CMC on pH of the solution. Activity was tested using buffers of different pH, as indicated in the figure. Data on the Y-axis represents absorbance at 595 nm corrected for background signal of cell culture media at each pH tested.

Development of a high throughput method in yeast to identify active cellulases from insects

While our screening data supports that insects are a potentially important prospecting resource for novel cellulolytic enzymes to facilitate production of lignocellulosic ethanol, identification and characterization of these enzymes is hindered by the lack of necessary genomic resources. In fact, there is a lack of genetic sequences for most of the insect species we detected with high cellulolytic activity. To overcome this limitation, we proposed developing a high-throughput functional assay to screen cDNA expression libraries for novel insect cellulases. We expect that this method would be more cost-effective in the identification of novel enzymes with applications in biofuel production. In addition, we chose yeast as our heterologous expression system and as a model for expression of cellulolytic enzymes in single-step ethanol production strategies. However, there are no published data supporting functional expression of insect cellulases in yeast. To address this knowledge gap and provide “proof-of-concept” for our high throughput screening method, we transformed Saccharomyces cerevisiae yeast to express the full-length cDNA encoding the TcEG1 endoglucanase from T. castaneum. In collaboration with Dr. Todd Reynolds (UT Microbiology), we transformed yeast with an expression vector containing the TcEG1 cDNA under the control of the glyceraldehyde-3 phosphate

17

dehydrogenase (GPD) promoter. Transformed yeast secreted the TcEG1 protein as a functional β 1,4-endoglucanase, which allowed transformants to survive on solid media with cellulose as the only carbon source available. These results demonstrate the potential for expression of functional insect cellulases in yeast cultures, supporting the potential screening of cDNA expression libraries in yeast to detect clones expressing functional cellulases. In addition, we were able to demonstrate that the secreted TcEG1 retained full activity at pH 9, as previously found with the same enzyme expressed in insect cell cultures, and that increased levels of activity were detected up to the highest pH tested (pH 12, Fig. 5).

Figure 5. Quantitative measure of activity under diverse pH for TcEG1 secreted by yeast clones. In each instance, 50 µg of concentrated protein was allowed to digest CMC for 1 hour. Each experiment represents one biological replicate (N=3) except for activity at pH 12 which was tested with two biological replicates (N=6). Background reducing sugars were corrected for by subtracting final from initial values of the calculated reducing sugars in the sample. Data is presented as the mean and corresponding standard error calculated from each experiment. Values are presented as units per gram of protein (U/g), with a unit (U) of cellulolytic activity being defined as the amount of enzyme required to produce 1 µmol of reducing sugar per minute.

• Conclusions

1) Digestive fluids from diverse insects contain cellulolytic enzymes. Levels of cellulolytic activity vary among taxonomic orders, although a number of species displayed levels of activity higher than those of termites.

2) Endoglucanase is the dominant activity in digestive fluids from insects. All detected cellulases in our work belong to GHF9.

3) Basal insect groups and coleopteran larvae feeding on decaying wood displayed the highest levels of cellulolytic activity, supporting the existence of effective enzymes that may have applications in the biofuel industry.

4) The TcEG1 cellulase represents the endoglucanase with the highest stability in alkaline pH reported. Highest levels of TcEG1 activity against CMC were detected at pH 12. This stability at high pH suggests this enzyme may be amenable to use in alkaline ionic liquids during pretreatment of lignocellulosic biomass.

5) Insect cellulases retained activity when expressed heterologously in yeast.

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Publications and presentations Publications in peer-reviewed journals Shirley, D., Oppert, C., Reynolds, T., Miracle, B., Oppert, B., Klingeman, B., and J. L. Jurat-Fuentes (In preparation) “Functional characterization of an endoglucanase from Tribolium castaneum (TcEG1) in Saccharomyces cerevisiae”. Shirley, D., Oppert, C., Klingeman, W., and J L. Jurat-Fuentes (In preparation) “Characterization of high cellulolytic activity in digestive fluids from Thermobia domestica”. Willis, J. D., Oppert, B., Oppert, C., Klingeman, W. E., and J. L. Jurat-Fuentes (2011) “Cloning, expression, and characterization of a GHF9 cellulase from Tribolium castaneum (Coleoptera: Tenebrionidae)” J. Insect Physiol. 57(2): 300-306. Willis, J.D., Oppert, C., and J. L. Jurat-Fuentes (2010) "Characterization of cellulolytic activity from digestive fluids of Dissosteira carolina (Orthoptera: Acrididae)" Comp. Biochem. Physiol. Part B Biochem. Mol. Biol. 157(3): 267-272. Willis, J.D., Oppert, C., and J. L. Jurat-Fuentes (2010) "Methods for discovery and characterization of cellulolytic enzymes from insects" Insect Sci. 17(3): 184-198.

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Oppert, C., Klingeman, W.E., Willis, J.D., Oppert, B., and J. L. Jurat-Fuentes (2010) "Prospecting for cellulolytic activity in insect digestive fluids" Comp. Biochem. Physiol. Part B, 155: 145-154. Invited presentations Invited speaker: Jurat-Fuentes, J. L. “Prospecting insects for cellulases to optimize biofuel production”. Invited seminar at the Biofuels Institute, Jiangsu University, Zhanjiang (China), June 2010. Invited speaker: Jurat-Fuentes, J. L. “Prospecting insects for enzymes to improve lignocellulosic biofuel production”. Invited seminar at the Department of Entomology, University of Illinois. Urbana/Champlain (IL), October 2009. Invited speaker: Jurat-Fuentes, J. L. “Insects and biofuels: got cellulases?”. Program Symposium: Metamorphosis: The Development of Entomology into an Interdisciplinary Science. 56th Annual ESA meeting, Reno (NV), November 2008. Contributing presentations Poster display: “Functional characterization of an endoglucanase from Tribolium castaneum in Saccharomyces cerevisae”. Shirley, D., Miracle, B., Reynolds, T., Klingeman, W., and Jurat-Fuentes, J. L. 58th Annual meeting of the Entomological Society of America (ESA) at San Diego CA), December 2010. Note: This poster won the award to the best graduate student presentation in its category. Oral presentation: “Screening insect digestive fluids for cellulolytic activity”. J. L. Jurat-Fuentes. Sun Grant Initiative Energy Conference, Washington (DC), March 2009. Poster display: “Identification and characterization of novel cellulolytic activity from salivary and digestive fluids of Dissosteira carolina (Orthoptera: Acrididae)”. Willis, J. D., Klingeman, W., Oppert, C., and J. L. Jurat-Fuentes. 56th Annual Meeting of the ESA, Reno (NV), November 2008. Oral presentation: “Identification and characterization of novel cellulase activity from digestive fluids of Dissosteira carolina (Orthoptera: Acrididae)”. Willis, J. D., Klingeman, W., Oppert, C., and J. L. Jurat-Fuentes. 35th Annual Meeting of the Tennessee Entomological Society, Nashville (TN), October 2008. Note: This presentation won the award to the best graduate student presentation.

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Acknowledgements The research team is grateful to a number of enthusiastic and very helpful collaborators that provided expertise or materials for the completion of the research described. The list of these collaborators includes Dr. Sean Marshall and Dr. Trevor Jackson (AgResearch, New Zealand), Dr. Todd Reynolds (University of Tennessee, Microbiology), and Dr. Damon J. Crook (USDA APHIS). Support for this research was provided by a grant from the Southern Sun Grant Center with funds provided by the U.S. Department of Energy, Office of the Biomass Program.

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Table of contents Page Cover page 1 Abstract 2 Full report -Problem 3 -Approach 3 -Methodology 4 -Findings -Screening for cellulase activity in insect digestive fluids 11 -Characterization of cellulolytic activity in specific insect species 12 -Cloning, expression, and characterization of TcEG1 15 -Development of a high throughput method in yeast to identify Active cellulases from insects 16 -Conclusions 17 -References 18 Publications and Presentations -Publications in peer-reviewed journals 20 -Invited presentations 21 -Contributing presentations 21 Acknowledgements 22 Table of contents 24 List of figures and tables 24 Executive summary 25 List of figures and tables Page Figure 1.-Results from cellulase screening 13

Figure 2.-Cellulases from D. carolina 14 Figure 3.-Expression of TcEG1 in S2 cells 15 Figure 4.-Thermostability and pH optimum of TcEG1 16 Figure 5.-Stability of TcEG1 from yeast cultures in alkaline pH 17

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Executive summary Production of ethanol from plant biomass is not cost effective when compared to ethanol

made from starch or with fossil fuels due to the difficulty of degrading the plant biomass into its basic sugar components that can then be fermented to ethanol. It is estimated that the discovery of novel cellulases, enzymes capable of digesting the plant material, could greatly reduce costs and increase yield of ethanol production from plant biomass. While insects are highly effective in degrading plant material for energy, research projects have concentrated on a limited number of species. The goal of our project was to screen an extensive collection of digestive fluids from diverse insects to identify species containing highly effective cellulolytic systems. Once species of interest were identified, we purified and cloned the cellulases involved in the activity. Production of these cellulases in bacteria or yeast allowed us to characterize their activity and stability to evaluate potential applications in biofuel industry. Our data demonstrate that insects are a valid resource for prospecting for new cellulases that may greatly impact cost effectivity and increase competiveness of biofuel ethanol.