Effective Microwell Plate-Based Screening Method for Microbes Producing Cellulase and Xylanase and Its Application
Effective Microwell Plate-Based Screening Method for Microbes Producing Cellulase and Xylanase and Its Application
Journal of Microbiology and Biotechnology. 2014. Nov, 24(11): 1559-1565
Copyright © 2014, The Korean Society For Microbiology And Biotechnology
  • Received : May 21, 2014
  • Accepted : July 24, 2014
  • Published : November 28, 2014
Export by style
Cited by
About the Authors
Jennifer Jooyoun Kim
School of Marine and Atmospheric Science, Stony Brook University, Stony Brook, NY 11794-5000, USA
Young-Kyung Kwon
Department of Environmental Marine Sciences, Hanyang University, Ansan 426-791, Republic of Korea
Ji Hyung Kim
Korea Institute of Ocean Science and Technology, Ansan 426-744, Republic of Korea
Soo-Jin Heo
Korea Institute of Ocean Science and Technology, Ansan 426-744, Republic of Korea
Youngdeuk Lee
Korea Institute of Ocean Science and Technology, Ansan 426-744, Republic of Korea
Su-Jin Lee
Korea Institute of Ocean Science and Technology, Ansan 426-744, Republic of Korea
Won-Bo Shim
Department of Plant Pathology and Microbiology, Texas A&M University, College Station, TX 77843-2132, USA
Won-Kyo Jung
Department of Biomedical Engineering, Pukyong National University, Busan 608-737, Republic of Korea
Jung-Ho Hyun
Department of Environmental Marine Sciences, Hanyang University, Ansan 426-791, Republic of Korea
Kae Kyoung Kwon
Korea Institute of Ocean Science and Technology, Ansan 426-744, Republic of Korea
Do-Hyung Kang
Korea Institute of Ocean Science and Technology, Ansan 426-744, Republic of Korea
Chulhong Oh
Korea Institute of Ocean Science and Technology, Ansan 426-744, Republic of Korea

Cellulase and xylanase are main hydrolysis enzymes for the degradation of cellulosic and hemicellulosic biomass, respectively. In this study, our aim was to develop and test the efficacy of a rapid, high-throughput method to screen hydrolytic-enzyme-producing microbes. To accomplish this, we modified the 3,5-dinitrosalicylic acid (DNS) method for microwell plate-based screening. Targeted microbial samples were initially cultured on agar plates with both cellulose and xylan as substrates. Then, isolated colonies were subcultured in broth media containing yeast extract and either cellulose or xylan. The supernatants of the culture broth were tested with our modified DNS screening method in a 96-microwell plate, with a 200 µl total reaction volume. In addition, the stability and reliability of glucose and xylose standards, which were used to determine the enzymatic activity, were studied at 100℃ for different time intervals in a dry oven. It was concluded that the minimum incubation time required for stable color development of the standard solution is 20 min. With this technique, we successfully screened 21 and 31 cellulase- and xylanase-producing strains, respectively, in a single experimental trial. Among the identified strains, 19 showed both cellulose and xylan hydrolyzing activities. These microbes can be applied to bioethanol production from cellulosic and hemicellulosic biomass.
Cellulase is an enzyme that hydrolyzes the β-1,4-glucosidic bond present in cellulose, which is the main component of the plant cell wall [2] . Xylanase similarly metabolizes β-1,4-xylosidic linkages found in xylan, which is the second most abundant natural polysaccharide behind cellulose [7] . Such enzymes that are capable of hydrolyzing polysaccharides have tremendous potential for numerous industrial applications, including food processing [4 , 11] , bioethanol development [5 , 9 , 18 , 25] , pulp and paper processing [15 , 23] , and animal feed manufacturing [3 , 17 , 32] . Many microorganisms are known to produce cellulases and xylanases to degrade cellulose and xylan, and ultimately utilize the hydrolyzed products as its carbon source [13 , 22] .
In previous studies, various screening methods have been suggested for the detection of polysaccharidedegrading microbes. These methods usually utilize agar plates with application of different types of stains, such as Congo red or Gram’s iodine [12 , 14 , 30 , 31] . Polysaccharidedegrading strains are distinguishable on these stained plates as clear zone forms along with the growth of the colony. Efforts were made to improve the efficiency of these test methods by adopting microwell plate-based colorimetric strategies [8 , 10 , 19 , 27 , 34 , 35] . One of the widely used quantifiable screening methods for cellulaseand xylanase-producing microbes is the 3,5-dinitrosalicylic acid (DNS) method. The DNS method was originally proposed to test the sugar contents in diabetic patients’ urine [28 , 29] , reliably identifying the presence of glucose in urine samples by color reaction. This method has also been widely used in testing the presence of different polysaccharides, such as cellulose, mannan, xylan, laminarin, agar, and sucrose, as the DNS solution reacts with reducing sugars in the sample [1 , 16 , 19 , 20 , 21 , 24] .
Our key aim was to develop and test the efficacy of a rapid, high-throughput method to screen hydrolytic-enzymeproducing microbes. The microwell plate-based DNS screening method demonstrated in this study allows efficient high-throughput screening of microbial strains that produce hydrolytic enzymes. This DNS-based screening method detects simple sugers produced by microbial enzymatic reactions. The microbes with polysaccharide hydrolyzing enzyme production identified with this procedure can be further investigated for industrial applications.
Materials and Methods
- Microbes Sampling and Culture
Microbial samples were collected on islands located on the west coast of South Korea. Sampling for microbes that produce cellulase or xylanase was done by collecting degraded plant materials and herbivorous insects. Samples were categorized as either terrestrial or marine samples based on their site of collection.
Two types of agar plates were prepared to grow microbes of interest: ground water (GW) for terrestrial samples and sea water (SW) for marine samples. The names of each plate indicated the base water used to prepare those agar plates. Plates were designed to contain 0.1% of carboxymethyl-cellulose (CMC) sodium salt (Sigma, Germany) and 0.1% of xylan from beechwood (TCI, Japan), along with 2% of Bacto agar (BD Biosciences, USA), to selectively grow cellulose- and xylan-degrading microbes, respectively.
Suspensions of minced terrestrial and marine samples were diluted and spread on the appropriate type of agar plates and incubated at 30℃ for 2-3 days. Successfully isolated colonies were cultured in two different types of liquid broth. First, 0.1% of CMC sodium salt and 0.1% of yeast extract (BD Biosciences, USA) broth was used to culture cellulase-active microbes. Microbes with xylanase activity were cultured in medium containing 0.1% of xylan and 0.1% of yeast extract. Both batches were incubated at 30℃ with agitation (200 rpm) for 48 h. Finally, crude enzyme solution was extracted by taking the supernatant of the cultured broth after centrifugation at 12,000 rpm for 1 min. The overall experimental procedure is provided as a flow chart in Fig. 1 .
PPT Slide
Lager Image
Schematic flow chart of novel screening method for cellulase- and xylanase-producing microbes. Ground water and sea water were the type of basal waters used to prepare the agar plates or broths. Cellulose, xylan, and yeast extract were used of 0.1% concentration in all media.
- DNS Solution Preparation
DNS solution was prepared by dissolving 0.25 g of 3,5-dinitrosalicyclic acid and 75 g of sodium potassium tartrate in 50 ml of 2 M sodium hydroxide solution. Then, the final volume was brought up to 250 ml using distilled water. Prepared solutions were stored in the dark.
- DNS Standard Curve Development
Sterilized polystyrene 96-well plates (SPL, Korea) were used to carry out the DNS reactions. Series of glucose and xylose standards were independently prepared for the cellulase and xylanase experiments. Standard solutions each contained 0, 0.2, 0.4, 0.6, 0.8, and 1 µmole of D(+)-glucose monohydrate (Sigma, Germany) or D(+)-xylose (Sigma, Germany) in 40 µl of distilled water. Then, 160 µl of DNS solution was added into each well, yielding a final volume of 200 µl. The microwell plate, with its top covered to minimize evaporative loss, was wrapped with aluminum foil to prevent the plate from melting from the direct contact with the dry oven. The optimal reaction time for standard curve stability was tested at 10, 20, and 30 min in the 100℃ dry oven. Finally, the test plates were analyzed with a microplate reader (Bio-TEK Instruments, USA) to determine the optical density of each sample at 570 nm.
- Screening of Cellulase- and Xylanase-Producing Microbes
CMC and xylan 1% substrate solutions were prepared to test cellulase and xylanase activity, respectively. In each well, 20 µl of crude enzyme solution and 20 µl of appropriate 1% substrate solution were added. The negative control mixture contained 20 µl of distilled water with 20 µl of 1% substrate solution. The enzymatic reaction was carried out by incubating the microplate with its cover tightly sealed at 40℃, by floating in a water bath for 1 h. The DNS reaction was carried out at 100℃ for 20 min (previously determined optimal time) in a dry oven and analyzed as described earlier.
- Strain Identification
Genomic DNA was isolated from microbial strains identified to express cellulase and xylanase. The 16S ribosomal DNA was amplified with a pair of universal primers (16S-27F: 5’-AGAGTTTGATCMTGGCTCAG-3’; and 16S-1492R: 5’-TACGGYTACCTTGTTACGACTT-3’) for bacterial identification. Fungal identification was performed by amplifying the internal transcribed spacer (ITS) region using a set of primers (ITS1: 5’-TCCGTAGGTGAACCTGCGG-3’; and ITS4: 5’-TCCTCCGCTTATTGATATGC-3’). Amplified products were sequenced (Macrogen Inc., Seoul, Korea) and analyzed using the BLASTn algorithm of National Center for Biotechnology Information (NCBI, ).
- Microwell Plate-Based Cellulase and Xylanase Assay
The minimum reaction time required for DNS standard curve stability at 100℃ was determined to be 20 min ( Fig. 2 ). Both glucose and xylose standard curves showed instability when reacted for only 10 min, whereas little variation when reacted for 20 min or longer. Consequently, we concluded that 20 min is the most effective time period to stabilize the color development of the standard curve.
PPT Slide
Lager Image
Standard curves of glucose (A) and xylose (B). The optimal condition test for the DNS reaction was performed at 100℃ in a dry oven for 10, 20, and 30 min. The actual color gradation that developed in the series of standard solutions resulting from the DNS reaction is shown at the bottom right corner of each graph.
As the DNS method detects the presence of reducing sugars in samples by colorimetric response, microbes producing extracellular cellulase and xylanase were easily detected even with the naked eye when the DNS reaction was completed ( Fig. 3 ). Among 116 strains tested, 21 showed some degree of cellulase activity: J16, J29, J30, J42, J46, J48, J49, J53, J58, J59, J64, J65, J78, J81, J82, J90, J103, J104, J109, J113, and J115. For xylanase, the following 31 strains showed a detectable level of enzymatic activity: J8, J16, J19, J27, J29, J30, J34, J42, J46, J47, J48, J49, J52, J53, J57, J58, J59, J61, J64, J65, J78, J85, J89, J90, J92, J96, J103, J104, J109, J113, and J115.
PPT Slide
Lager Image
Conspicuous color changes developed in the microwell plates after the DNS reaction had been carreid out for (A) cellulase activity and (B) xylanase activity. The color intensity of each sample indicates the relative enzymatic activity under the given condition in this experiment. Some strains, such as J48, J49, J103, J04, and J113, showed both active cellulase and xylanase activities.
A greater number of microbes produced xylanase than cellulase in general. In addition, the overall enzymatic activities of xylanase-producing isolates were higher than cellulase-producing isolates. Notably, strain J113 exhibited the strongest cellulase activity as well as the xylanase activity; it hydrolyzed 17.90 µmol/ml of cellulose and 61.49 µmol/ml of xylan per 1 h. Other samples, such as J48, J49, J103, and J104, also displayed relatively strong cellulase and xylanase activities, resulting in noticeable color change in DNS analyses.
The relative enzymatic activity of each strain determined by our screening method is summarized in Fig. 4 . From 116 strains screened, we found 19 strains exhibiting both cellulase and xylanase activities. There were 12 strains solely secreting xylan-degrading enzyme, and 2 strains with cellulase production only.
PPT Slide
Lager Image
Comparison of relative cellulase and xylanase activities. J113 processed 17.90 µmol/ml of cellulose and 61.49 µmol/ml of xylan in an hour, establishing the strongest relative enzymatic activity among the strains. Cellulase activity was weaker than xylanase activity overall. A total of 9 strains displayed xylanase activity greater than 20 µmol/ml/h, whereas none expressed cellulase activity greater than 20 µmol/ml/h.
- Strain Identification
Based on 16S rDNA and ITS sequence analyses, we were able to identify 10 fungi and 21 bacteria strains. Unfortunately, two strains (J8 and J89) were not identifiable as they were unable to be re-cultured. Table 1 summarizes the successfully identified strains that secrete cellulase or xylanase. Vibrio, Bacillus, Alternaria, Graphium, Planococcus, Nectria, Trichoderma, Streptomyces, Ochrobactrum, Frondihabitans, Cryptococcus, Cladosporium , and Cellvibrio produced both extracellular cellulase and xylanase. Pseudoalteromonas, Glaciecola, Staphylococcus, Xanthomonas , and Rhodococcus were determined to solely secrete xylanase, and Psychrobacter only cellulase.
Description of the bacteria and fungi used for the novel screening method.
PPT Slide
Lager Image
The strain number, GenBank accession number, similar strain identified along with its similarity percentage match, type of enzyme produced, and classification are category displayed. As a whole, 21 bacterial strains and 10 fungal strains were identified. Two strains (J8 and J89) were unable to be identified. UI, unidentified; X, xylanase; C, cellulase.
Traditionally, cellulase- or xylanase-producing microbes were screened on agar plates with Congo red or Gram’s iodine stains [12 , 14 , 30 , 31] . These methods require more effort and time compared with our new method, particularly when there are a large number of samples requiring highthroughput screening. Our study demonstrates a method that allows efficient screening and detection of microbes that produce extracellular saccharification enzymes by modifying the customary DNS method. Instead of using reaction tubes and a convential spectrophotometric analysis, we tested the efficacy of 96-well microplates and a microplate reader to carry out the experimental procedures. It requires fewer reaction steps and chemicals compared with the conventional method, reducing the labor input, time, and cost needed for these assays. Microwell platebased DNS experiments have been criticized for their longer incubation time than the traditional DNS method [34] and for poor representation of the stoichiometric amount of sugar [6 , 26] . Despite these criticisms, we successfully demonstrated that our microwell plate-based method can detect the presence of extracellularly secreted cellulase and xylanase in samples.
We initially cultured our microbial isolates on agar media with both CMC and xylan to support efficient growth. Subsequently, isolated colonies were subcultured in CMC and yeast extract broth or xylan and yeast extract broth to selectively grow enzyme-specific strains. Every microbial strains used in this study (a total of 116 strains) were cultured based on this procedure. CMC and xylan included in culture broths are expected to induce the production of cellulase and xylanase in microbes. Yeast extract was added as a complex nitrogen source to facilitate microbial growth.
We additionally studied the optimal reaction time for this microwell plate-based DNS method. The traditional DNS method suggests a 10 min reaction time at 100℃, as the reaction chambers are directly heated in boiling water. However, in our new method, the DNS reaction is instead carried out in a dry oven using air as the heating medium. This allows the plate to react at 100 ℃ without having to boil water, but we do recognize that air has a lower heat conductivity than water. Adjustment of the reaction time was necessary as a result, and the optimal incubation time was determined. We concluded that 10 min was insufficient to fully develop the coloration in the samples and that minimum of 20 min is required. However, extending the incubation time to 30 min was unnecessary as there were subtle difference in color development beyond a 20 min incubation time. As this DNS assay procedure depends on the absorbance value to determine the amount of reduced sugars in each sample, it is advised that microwell plates be handled with care, avoiding contamination or damage to the bottom surface in particular.
Our key aim of this study, establishing a high-throughput colorimetric screening of strains of interest, was successful. However, as the optimal reaction condition for each enzyme was not determined, we recognize that the color intensity of the samples itself does not indicate the absolute magnitude of enzymatic activity of each sample. Enzymatic reactions are sensitive to a number of environmental factors, such as temperature and pH, and the given condition in this experiment is most likely not the optimum. Thus, color changes observed after the DNS reaction simply indicate the presence of hydrolyzed sugars produced by saccharification enzymes. The intensity of color and calculated crude enzyme activity described in this study are not the absolute standard for determination of the enzymatic activity level.
Our high-throughput method will be beneficial in identifying industrially useful strains. For example, cellulase and xylanase, the enzymes of interest in this research, are both widely used in many applications, including food processing and pulp treatments [4 , 15 , 23] . Moreover, such saccharifying enzymes are expected to be valuable in industrial biofuel research that is increasingly gaining popularity. Polysaccharide hydrolyzing enzymes play a crucial role in processing raw biomass to produce biofuel [33] . Although we applied the method only to search cellulaseand xylanase-yielding microbes, we expect this method to be effective in screening other types of saccharification enzyme-producing microbes as well. Agarase, amylase, laminarinase, or mannanase may also be studied by using this method when the respective saccharide substrate solutions are readily available.
This research was financially supported by research grants from the Korea Institute of Ocean Science and Technology (KIOST; PE99213, PE99214) and “Creative Allied Project (CAP)” of Korea Research Council of Fundamental Science and Technology, Korea Institute of Science and Technology, and KIOST (PE99272). Also this paper was studied with the support of the MSIP (Ministry of Science, Ict & future Planning).
View Fulltext  
Agrawal M , Pradeep S , Chandraraj K , Gummadi SN 2005 Hydrolysis of starch by amylase fromBacillussp. KCA102: a statistical approach. Process. Biochem. 40 2499 - 2507    DOI : 10.1016/j.procbio.2004.10.006
Bayer EA , Chanzy H , Lamed R , Shoham Y 1998 Cellulose, cellulases and cellulosomes. Curr. Opin. Struct. Biol. 8 548 - 557    DOI : 10.1016/S0959-440X(98)80143-7
Beauchemin KA , Rode LM , Sewalt VJH 1995 Fibrolytic enzymes increase fiber digestibility and growth rate of steers fed dry forages. Can. J. Anim. Sci. 75 641 - 644    DOI : 10.4141/cjas95-096
Camacho NA , Aguilar OG 2003 Production, purification, and characterization of a low-molecular-mass xylanase fromAspergillussp. and its application in baking. Appl. Biochem. Biotechnol. 104 159 - 171    DOI : 10.1385/ABAB:104:3:159
Chang MCY 2007 Harnessing energy from plant biomass. Curr. Opin. Chem. Biol. 11 677 - 684    DOI : 10.1016/j.cbpa.2007.08.039
Chundawat SPS , Balan V , Dale BE 2008 High-throughput microplate technique for enzymatic hydrolysis of lignocellulosic biomass. Biotechnol. Bioeng. 99 1281 - 1294    DOI : 10.1002/bit.21805
Collins T , Gerday C , Feller G 2005 Xylanases, xylanase families and extremophilic xylanases. FEMS Microbiol. Rev. 29 3 - 23    DOI : 10.1016/j.femsre.2004.06.005
Feng ZH , Wang YS , Zheng YG 2011 A new microtiter plate-based screening method for microorganisms producing alpha-amylase inhibitors. Biotechnol. Bioprocess Eng. 16 894 - 900    DOI : 10.1007/s12257-011-0033-7
Fujita Y , Takahashi S , Ueda M , Tanaka A , Okada H , Morikawa Y 2002 Direct and efficient production of ethanol from cellulosic material with a yeast strain displaying cellulolytic enzymes. Appl. Environ. Microbiol. 68 5136 - 5141    DOI : 10.1128/AEM.68.10.5136-5141.2002
Goncalves C , Rodriguez-Jasso RM , Gomes N , Teixeira JA , Belo I 2010 Adaptation of dinitrosalicylic acid method to microtiter plates. Anal. Methods 2 2046 - 2048    DOI : 10.1039/c0ay00525h
Harbak L , Thygesen HV 2002 Safety evaluation of a xylanase expressed inBacillus subtilis. Food Chem. Toxicol. 40 1 - 8    DOI : 10.1016/S0278-6915(01)00092-8
Hendricks CW , Doyle JD , Hugley B 1995 A new solid medium for enumerating cellulose-utilizing bacteria in soil. Appl. Environ. Microbiol. 61 2016 - 2019
Henrissat B 1994 Cellulases and their interaction with cellulose. Cellulose 1 169 - 196    DOI : 10.1007/BF00813506
Kasana R , Salwan R , Dhar H , Dutt S , Gulati A 2008 A rapid and easy method for the detection of microbial cellulases on agar plates using Gram’s iodine. Curr. Microbiol. 57 503 - 507    DOI : 10.1007/s00284-008-9276-8
Kirk O , Borchert TV , Fuglsang CC 2002 Industrial enzyme applications. Curr. Opin. Biotechnol. 13 345 - 351    DOI : 10.1016/S0958-1669(02)00328-2
Kurakake M , Komaki T 2001 Production of β-mannanase and β-mannosidase fromAspergillus awamoriK4 and their properties. Curr. Microbiol. 42 377 - 380    DOI : 10.1007/s002840010233
Lewis GE , Hunt CW , Sanchez WK , Treacher R , Pritchard GT , Feng P 1996 Effect of direct-fed fibrolytic enzymes on the digestive characteristics of a forage-based diet fed to beef steers. Anim. Sci. 74 3020 - 3028
Menon V , Prakash G , Prabhune A , Rao M 2010 Biocatalytic approach for the utilization of hemicellulose for ethanol production from agricultural residue using thermostable xylanase and thermotolerant yeast. Bioresour. Technol. 101 5366 - 5373    DOI : 10.1016/j.biortech.2010.01.150
Miyazaki K , Takenouchi M , Kondo H , Noro N , Suzuki M , Tsuda S 2006 Thermal stabilization ofBacillus subtilisfamily-11 xylanase by directed evolution. J. Biol. Chem. 281 10236 - 10242    DOI : 10.1074/jbc.M511948200
Nichols EJ , Beckman JM , Hadwiger LA 1980 G lycosidic enzyme activity in pea tissue and pea-Fusarium solaniinteractions. Plant Physiol. 66 199 - 204    DOI : 10.1104/pp.66.2.199
Oh C , Nikapitiya C , Lee Y , Whang I , Kim SJ , Kang DH , Lee J 2010 Cloning, purification and biochemical characterization of beta agarase from the marine bacteriumPseudoalteromonassp. AG4. J. Ind. Microbiol. Biotechnol. 37 483 - 494    DOI : 10.1007/s10295-010-0694-9
Prade RA 1996 Xylanases: from biology to biotechnology. Biotechnol. Genet. Eng. 13 101 - 131    DOI : 10.1080/02648725.1996.10647925
Rahkamo L , Siika-Aho M , Vehviläinen M , Dolk M , Viikari L , Nousiainen P , Buchert J 1996 Modification of hardwood dissolving pulp with purifiedTrichoderma reeseicellulases. Cellulose 3 153 - 163    DOI : 10.1007/BF02228798
Reczey K , Szengyel Zs , Eklund R , Zacchi G 1996 Cellulase production byT. reesei. Bioresour. Technol. 57 25 - 30    DOI : 10.1016/0960-8524(96)00038-7
Rubin EM 2008 Genomics of cellulosic biofuels. Nature 454 841 - 845    DOI : 10.1038/nature07190
Schwald W , Chan M , Breuil C , Saddler JN 1988 Comparison of HPLC and colorimetric methods for measuring cellulolytic activity. Appl. Microbiol. Biotechnol. 28 398 - 403    DOI : 10.1007/BF00268203
Shankar M , Priyadharshini R , Gunasekaran P 2009 Quantitative digital image analysis of chromogenic assays for high throughput screening of α-amylase mutant libraries. Biotechnol. Lett. 31 1197 - 1201    DOI : 10.1007/s10529-009-9999-z
Sumner JB 1921 Dinitrosalicyclic acid: a reagent for the estimation of sugar in normal and diabetic urine. J. Biol. Chem. 47 5 - 9
Sumner JB 1924 The estimation of sugar in diabetic urine, using dinitrosalicylic acid. J. Biol. Chem. 62 287 - 290
Teather RM , Wood PJ 1982 Use of Congo red-polysaccharide interactions in enumeration and characterization of cellulolytic bacteria from the bovine rumen. Appl. Environ. Microbiol. 43 777 - 780
Ten LN , Im WT , Kim MK , Kang MS , Lee ST 2004 Development of a plate technique for screening of polysaccharidedegrading microorganisms by using a mixture of insoluble chromogenic substrates. J. Microbiol. Methods 56 375 - 382    DOI : 10.1016/j.mimet.2003.11.008
Twomey LN , Pluske JR , Rowe JB , Choct M , Brown W , McConnell MF , Pethick DW 2003 The effects of increasing levels of soluble non-starch polysaccharides and inclusion of feed enzymes in dog diets on faecal quality and digestibility. Anim. Feed Sci. Tech. 108 71 - 82    DOI : 10.1016/S0377-8401(03)00161-5
Wilson DB 2009 Cellulases and biofuels. Curr. Opin. Biotechnol. 20 295 - 299    DOI : 10.1016/j.copbio.2009.05.007
Wood IP , Elliston A , Ryden P , Bancroft I , Roberts IN , Waldron KW 2012 Rapid quantification of reducing sugars in biomass hydrolysates: improving the speed and precision of the dinitrosalicylic acid assay. Biomass Bioenergy 44 117 - 121    DOI : 10.1016/j.biombioe.2012.05.003
Xiao Z , Storms R , Tsang A 2006 A quantitative starchiodine method for measuring alpha-amylase and glucoamylase activities. Anal. Biochem. 351 146 - 148    DOI : 10.1016/j.ab.2006.01.036