An improved phenol-sulfuric acid method for the quantitative measurement

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<p>Algal Research 70 (2023) 102986</p><p>Available online 24 January 2023</p><p>2211-9264/© 2023 Elsevier B.V. All rights reserved.</p><p>An improved phenol-sulfuric acid method for the quantitative measurement</p><p>of total carbohydrates in algal biomass</p><p>Wei Chen a,b,*,1, Lei Gao a,1, Lirong Song d, Milton Sommerfeld b, Qiang Hu b,c,*</p><p>a School of Marine Science and Engineering, Qingdao Agricultural University, PR China</p><p>b Department of Applied Sciences and Mathematics, Arizona State University, 7001 E. Williams Field Road, Mesa, AZ 85212, USA</p><p>c Instiute for Advanced Study, Shenzhen University, Nanshan District, Shenzhen, Guangdong 518060, PR China</p><p>d Institute of Hydrobiology, the Chinese Academy of Sciences, Wuhan, Hubei 430072, PR China</p><p>A R T I C L E I N F O</p><p>Keywords:</p><p>Total carbohydrates</p><p>Phenol</p><p>Sulfuric acid</p><p>Microalgae</p><p>Determination</p><p>Colorimetric measurement</p><p>A B S T R A C T</p><p>Total carbohydrate analysis with phenol‑sulfuric acid method is the simplest method among many colorimetric</p><p>and modern instrument analytical methods. It has great potential to be used as a rapid screening method for</p><p>characterization of algal biomass for algae-based biofuel research. The reproducibility and accuracy of the</p><p>conventional phenol‑sulfuric method remain to be improved when applied directly to algal biomass. In the</p><p>present study, several important factors that influenced the colorimetric determination were characterized and</p><p>methods to eliminate the influences were further developed. Results indicated that system temperature and the</p><p>addition mode of sulfuric acid and phenol were the key parameters which affected the color development and</p><p>reproducibility of the conventional methods. A new design using the premixed reagent replacing the individual</p><p>phenol and sulfuric acid solution solved this problem ideally. The color inferences caused by algal pigments</p><p>including chlorophylls and carotenoids were quantitatively analyzed. Solutions for reducing overestimation of</p><p>total carbohydrates were also addressed. In addition, hydrolysis conditions for algae biomass such as type of</p><p>acids, temperature, and time were also investigated and optimized in this study. The new modified phe-</p><p>nol‑sulfuric acid method has been shown to be more reproducible and reliable than the conventional methods for</p><p>the analysis of total carbohydrates from algal biomass.</p><p>1. Introduction</p><p>In recent years, algae biomass-derived biofuels have received</p><p>increased attention worldwide due to its high potential to be converted</p><p>into several kinds of biofuels including biodiesel, bioethanol, butanol</p><p>and so on [1–3]. For studies on the production of bioethanol using algal</p><p>biomass, determination of total carbohydrate content is helpful for</p><p>chemical characterization and for studies related to both strain selection</p><p>and downstream bioconversion processing. In the past half-century,</p><p>many methods based on the reaction of carbohydrates with aromatic</p><p>compounds using an acid medium have been reported for the spectro-</p><p>photometric analysis of total carbohydrates [4–9] and among them, the</p><p>phenol‑sulfuric acid method is used widely, because of its high sensi-</p><p>tivity and simplicity, since its introduction by Dubois et al. [6,10]. This</p><p>colorimetric method and the improved methods, based on Dubois’</p><p>original form have been employed for various kinds of samples including</p><p>environmental samples [11,12], higher plants [13–15], fungi [16,17],</p><p>human body fluid [18], and algae [19].</p><p>For carbohydrate determination in microalgae and other biological</p><p>samples, usually dissolved or purified carbohydrate extractions are used</p><p>as the samples for further colorimetric determinations [20,21]. There</p><p>are few early reports of carbohydrate measurements using whole algal</p><p>biomass after acid hydrolysis [22,23]. As algae-based biofuel studies</p><p>continue, a simple, high through-put and reliable method for rapid</p><p>measurement of total carbohydrates while using whole algae biomass</p><p>for direct analysis, without additionally time-consuming extraction and</p><p>purification steps will be very critical. However, information is still</p><p>lacking for the carbohydrate analysis using whole biomass. The phe-</p><p>nol‑sulfuric acid method also would either occasionally cause large</p><p>standard errors between replicate analyses or overestimate/underesti-</p><p>mate the total carbohydrates even after numerous improvements</p><p>[12,24,25]. This means that the key parameter affecting color</p><p>* Corresponding authors at: Department of Applied Sciences and Mathematics, Arizona State University, 7001 E. Williams Field Road, Mesa, AZ 85212, USA.</p><p>E-mail addresses: chenwei@qau.edu.cn (W. Chen), huqiang@asu.edu (Q. Hu).</p><p>1 These authors contributed equally to this work.</p><p>Contents lists available at ScienceDirect</p><p>Algal Research</p><p>journal homepage: www.elsevier.com/locate/algal</p><p>https://doi.org/10.1016/j.algal.2023.102986</p><p>Received 14 April 2022; Received in revised form 27 August 2022; Accepted 19 January 2023</p><p>mailto:chenwei@qau.edu.cn</p><p>mailto:huqiang@asu.edu</p><p>www.sciencedirect.com/science/journal/22119264</p><p>https://www.elsevier.com/locate/algal</p><p>https://doi.org/10.1016/j.algal.2023.102986</p><p>https://doi.org/10.1016/j.algal.2023.102986</p><p>https://doi.org/10.1016/j.algal.2023.102986</p><p>http://crossmark.crossref.org/dialog/?doi=10.1016/j.algal.2023.102986&domain=pdf</p><p>Algal Research 70 (2023) 102986</p><p>2</p><p>development and/or hydrolysis still needs to be fully understood.</p><p>Moreover, very little information is available on the color interference</p><p>caused by algal pigments such as, chlorophyll, and carotene during the</p><p>course of carbohydrate analysis using the conventional phenol‑sulfuric</p><p>acid method.</p><p>One aim of this study was to systematically test the phenol‑sulfuric</p><p>acid method and identify key parameters which affect the color devel-</p><p>opment and simplify the method to allow for greater reproducibility.</p><p>Another aim of this study is to quantify the color interference effect</p><p>caused by algal pigments and make the quantification process with this</p><p>method more accurate and reliable.</p><p>2. Material and methods</p><p>2.1. Organisms and culture conditions</p><p>Algal strains were obtained from the in-house algae culture collec-</p><p>tion at the Laboratory for Algae Research and Biotechnology at the</p><p>Arizona State University Polytechnic Campus in Mesa, Arizona; Chlorella</p><p>zofingiensis LRB-AZ-1201; Scenedesmus sp. LRB-AS-0401; Nanno-</p><p>chloropsis sp.; Haematococcus pluvialis were used. Scenedesmus sp. and</p><p>C. zofingiensis were maintained in BG-11 culture medium. Chlorella</p><p>zofingiensis biomass 1 and biomass 2 refer to Chlorella zofingiensis LRB-</p><p>AZ-1201 at different growth stages and with different carbohydrate</p><p>content. Nannochloropsis sp were maintained in f/2 medium. All algal</p><p>strains except Haematococcus pluvialis were cultured in 100 cm × 100</p><p>cm × 2.5 cm vertical panel photobioreactors (containing 15 L of me-</p><p>dium) maintained at 20 ◦C while exposed to continuous illumination at a</p><p>light intensity of 300 μmol m− 2 s− 1. Culture mixing was provided by</p><p>bubbling air containing 2 % CO2 (v/v) directly into the culture. Hae-</p><p>matococcus pluvialis was grown in BG-11 culture medium at room tem-</p><p>perature using tubular photobioreactors (containing 800 mL of medium)</p><p>and aerated with 1.5 % CO2 in air at 50 mL min− 1. Light was provided by</p><p>cool white fluorescent lamps with 30 μmol photon m− 2 s− 1 of light in-</p><p>tensity. When a culture reached the stationary stage due to nitrogen</p><p>deficiency, it was transferred into fresh nitrogen-deficient BG-11 me-</p><p>dium (BG-11 without sodium nitrate) by centrifugation (10 ◦C for 5 min</p><p>at 4000 rpm) and moved to a high-light environment (220 μmol photon</p><p>m− 2 s− 1) to transform the green cells to red cysts with high astaxanthin</p><p>contents. The culture was further incubated for 10 days and then har-</p><p>vested by centrifugation at 10 ◦C for 5 min at 4000 rpm. All biomass</p><p>samples were harvested by centrifugation and stored at − 20 ◦C for</p><p>further use prior to be dried with freeze drier.</p><p>2.2. Chemicals and reagents</p><p>Analytical</p><p>grade (98 %) H2SO4 was obtained from VWR (USA) and</p><p>regent-grade phenol was from Sigma. D-glucose used for generating the</p><p>calibration curve was of analytical grade and from Fisher scientific</p><p>(USA). Pigment standards such as astaxanthin and β-carotenoid were</p><p>purchased from Sigma (USA) and all the other analytical grade chem-</p><p>icals and solvents were also purchased from Sigma or other commercial</p><p>suppliers.</p><p>2.3. Effect of incubation temperatures and time on the color development</p><p>for the phenol-H2SO4 method</p><p>The addition sequence of H2SO4 and phenol solution and the addi-</p><p>tion speed of H2SO4 may have some influence on the color development</p><p>process. In the present study, different ways of H2SO4 addition were</p><p>compared such as adding acid drop by drop to sample; adding acid all at</p><p>once; adding acid drop by drop and then incubating in boiling water for</p><p>10 min; and adding phenol followed by acid being added all at once. To</p><p>further understand the influence of incubation temperature, a D-glucose</p><p>solution was employed and H2SO4 was added drop by drop in a cold ice</p><p>bath to keep the system at low temperature, and then samples were</p><p>placed in water bath at 20 ◦C, 40 ◦C, 60 ◦C, 90 ◦C and 100 ◦C, respec-</p><p>tively. Incubation time of 0 min, 5 min, 10 min, 20 min, 30 min and 40</p><p>min were compared after the addition of H2SO4 with high and low</p><p>concentration sugar samples.</p><p>2.4. Effect of pigments, lipids and proteins on the phenol-H2SO4 method</p><p>The pigment standards, chlorophyll a, β-carotene, triacylglycerol and</p><p>BioRad BSA standards were added to D-glucose solutions to imitate</p><p>biomass contents for pigments, lipids and proteins of 4 %, 4 %, 50 % and</p><p>50 % (% of dry weight), respectively. The mixtures containing pigments</p><p>and different concentrations of D-glucose (imitate carbohydrate contents</p><p>of 5 %, 15 %, 25 %, 35 % and 50 % in dry algae biomass) were hy-</p><p>drolyzed in 4 M trifluoroacetic acid (TFA) for 4 h and then the hydro-</p><p>lysates were analyzed with the improved phenol-H2SO4 method. In this</p><p>method, the visualization reagent was premixed using the following</p><p>protocol. 100 mL concentrate H2SO4 was added slowly into 50 mL Mili-</p><p>Q water in an ice bath until the temperature of the solution reached</p><p>room temperature. 0.5 g phenol was then dissolved in the diluted H2SO4</p><p>to form the evenly visualization reagent. 20 μL of each hydrolysate were</p><p>introduced into a 2mL centrifuge tube with rubber seal and plastic cap</p><p>and then 0.9 mL of the visualization reagent was added to each of tube</p><p>immediately and placed in an ice bath. These centrifuge tubes were</p><p>capped and thoroughly vortexed and then incubated in a boiling water</p><p>bath for 15 to 20 min. After that tubes were taken out and cooled in an</p><p>ice bath. 300 μl samples from each tube were taken out and put into a</p><p>96-well plate for spectrophotometric determination.</p><p>2.5. Spectrophotometric determinations of carbohydrates</p><p>For the spectrophotometric determination of carbohydrates, Spec-</p><p>traMax 350 Microplate Reader (Molecular Devices, USA) was used for</p><p>the analysis of absorbance spectrum characters and absorbance at 490</p><p>nm according to the method described by Dubois et al. (1956).</p><p>2.6. Effects of algae pigments on the quantification of total carbohydrates</p><p>in algae biomass</p><p>To learn the influence of algal pigments on the on the quantification</p><p>of total carbohydrates in algae biomass with the phenol-sulfuric acid</p><p>method, algae biomass with and without pigment extractions were used</p><p>before hydrolysis and then hydrolysates were quantified with standard</p><p>curve generated with D-glucose. Moreover, the hydrolysates with pig-</p><p>ments were also used as blank samples for quantification and compared</p><p>quantification results with samples with or without pigment extractions.</p><p>For the pigment extraction, 10 mg of algal biomass were weighed into</p><p>15 mL centrifuge tubes followed by the addition of 0.5 mL acetic acid</p><p>into the bottom of tubes. Then, the tubes were capped and incubated in</p><p>water bath set at 80 ◦C for 20 min. After the incubation, 10 mL acetone</p><p>was added into the tubes and then vortexed vigorously for 1 min. The</p><p>tubes were then centrifuged for 5 min at 4000 rpm (20 ◦C) and super-</p><p>natants containing pigments were poured off carefully to obtain</p><p>pigment-free biomass.</p><p>2.7. Hydrolysis of algae biomass</p><p>For the hydrolysis experiment, different acids and hydrolyzing time</p><p>were systematically investigated. In detail, the decolorized algal samples</p><p>were added into 5 mL acid solutions, and then the centrifuge tubes</p><p>containing samples and acids were incubated in boiling water for a</p><p>certain time. The tubes were shaking every half hour by vortex during</p><p>the course of hydrolysis. As soon as the hydrolysis step was completed,</p><p>the tubes were taken out immediately and put into an ice-bath for</p><p>cooling. The detailed hydrolysis conditions were as follows: A) 4 M</p><p>trifluoroacetic acid (TFA) for 4 h at 100 ◦C; B) 2 M HCl for 4 h at 100 ◦C;</p><p>C) 4 M HCl for 2 h at 100 ◦C; D) 4 M HCl for 4 h at 100 ◦C; E) 6 M HCl for</p><p>W. Chen et al.</p><p>Algal Research 70 (2023) 102986</p><p>3</p><p>2 h at 100 ◦C; F) 6 M HCl for 4 h at 100 ◦C; G) pretreatment with 12 M</p><p>HCl for 2 h at room temperature, then dilute to 4 M and incubated at</p><p>100 ◦C for another 2 h; H) 80 % H2SO4 for 20 h at room temperature and</p><p>I) no hydrolysis process.</p><p>2.8. Statistical analysis</p><p>Comparisons of means were conducted by one-way analyses of</p><p>variance (ANOVA) followed by Bonferroni tests to identify the sources of</p><p>detected significance. In all cases, comparisons that showed a p value <</p><p>0.05 were considered significant.</p><p>3. Results and discussion</p><p>3.1. Causes of low reproducibility/repeatability and influence of</p><p>temperature in conventional phenol‑sulfuric acid method</p><p>Since the first introduction of phenol‑sulfuric acid method by Dubois</p><p>and his colleagues in 1956 [10], there have been numerous improve-</p><p>ments on this original method and its subsequent improved methods</p><p>regarding reproducibility, repeatability and accuracy [12,24–28].</p><p>However, large standard deviations occasionally occurred when this</p><p>method was applied to different samples including pure sugar standards</p><p>[24,25,27,29]. This suggests that some very important operation pa-</p><p>rameters for phenol‑sulfuric acid have been neglected and not fully</p><p>controlled in the course of the procedure. In original form of Dubois’s</p><p>method [10], 80 % phenol was added first into the sugar samples, and</p><p>then concentrated sulfuric acid would be added very rapidly to the</p><p>above sample-phenol mixtures. Moreover, the addition of sulfuric acid</p><p>required that the stream of acid being directed against the liquid surface</p><p>rather than against the side of the test tube in order to obtain good</p><p>mixing. After that, the tubes were allowed to stand for 10 min, and then</p><p>they were vortexed and placed for 10 to 20 min in a water bath at 25 ◦C</p><p>to 30 ◦C. In this protocol, there is no additional heating process during</p><p>the course of color development. However, many improvements on this</p><p>method were implemented in the following half century including im-</p><p>provements in the sequence of phenol and sulfuric acid [27], incubation</p><p>time and temperature [12,29], additional hot water incubation step and</p><p>temperatures [25,30].</p><p>In the present study, the first parameter that we tested was the</p><p>addition mode and sequence of sulfuric acid and phenol. It was pre-</p><p>sented in Fig. 1 that addition mode and sequence of sulfuric acid and</p><p>phenol will significantly (p < 0.05) influence the color development</p><p>process report numbers. When the sulfuric acid was added very slowly,</p><p>there was no dramatic increase in system temperature and the absor-</p><p>bance was very low. As a comparison, the absorbance and temperature</p><p>were very high when sulfuric acid was added rapidly no matter the</p><p>addition sequence of sulfuric acid and phenol. Interestingly, the absor-</p><p>bance could be significantly increased with further incubation in boiling</p><p>water for those tubes with sulfuric acid being added drop by drop. The</p><p>above phenomenon clearly indicated that the addition of sulfuric acid</p><p>could affect the system temperature and temperature might be a key</p><p>parameter in controlling the color development. To confirm this</p><p>assumption, sulfuric acid was added drop by drop and cooled in ice bath</p><p>to keep the system temperature below 20 ◦C. Then the tubes were vor-</p><p>texed to mix sulfuric acid and phenol thoroughly and then further</p><p>incubate the mixture in boiling water to develop the color. Data from</p><p>Fig. 2 indicated that absorbance dramatically increased with the in-</p><p>crease of temperature. The addition mode and sequence of sulfuric acid</p><p>and phenol are the likely cause of the differentiation in system</p><p>Fig. 1. The influence of addition of H2SO4 on the carbohydrate determination</p><p>A) drop by drop; B) all at one time; C) drop by drop, then incubated in boiling</p><p>water for 10 min; D) all at one time, add sulfuric acid prior to the addition</p><p>of phenol.</p><p>Fig. 2. Effect of temperatures on the absorbance at 490 nm for carbohydrate</p><p>analysis H2SO4-phenol protocol.</p><p>Fig. 3. Temperature was controlled by an ice bath for all treatments when</p><p>adding sulfuric acid, and then the mixtures were incubated in boiling water. A)</p><p>phenol was added firstly then sulfuric acid was added drop by drop; B) sulfuric</p><p>acid was added drop by drop followed by the addition of phenol C) sulfuric acid</p><p>and phenol were premixed in an ice bath, then add the reagent to samples drop</p><p>by drop; D) sulfuric acid and phenol were premixed in an ice bath, then add the</p><p>reagent to samples all at one time.</p><p>W. Chen et al.</p><p>Algal Research 70 (2023) 102986</p><p>4</p><p>temperature and further lead to the differentiation in developed colors.</p><p>Although Dubois et al. [10] already realized that the good mixing and</p><p>addition of sulfuric acid from the liquid surface was very important to</p><p>get reproducible results, the key parameter–temperature is still not</p><p>controllable in the subsequent improved methods even until now.</p><p>Moreover, the additional sequences of sulfuric acid and phenol also have</p><p>significant influence on the developed color according to the previous</p><p>studies. In the investigation by Rao and Pattabiraman [27], it was re-</p><p>ported that phenol underwent sulfonation in suit and phenol-sulfonic</p><p>acid formed decreased the absorbance of mixtures for many hexoses</p><p>and pentoses, and suggested that the addition of concentrated sulfuric</p><p>acid to the samples followed by phenol solution could yield better re-</p><p>sults. However, our study also indicated that there is almost no color for</p><p>the mixtures, if the system temperatures were controlled during the</p><p>course of adding sulfuric acid and phenol (This implies that no chemical</p><p>reactions occurred in this step). After the sample - sulfuric acid-phenol</p><p>mixture was incubated in boiling water bath, color was fully devel-</p><p>oped and there was no difference on color intensities for the treatments</p><p>with either sulfuric acid being added prior to phenol or phenol being</p><p>added prior to sulfuric acid (Fig. 3). Our results also suggested that</p><p>addition mode of sulfuric acid has also no influence on the developed</p><p>color intensities if the temperature could be controlled properly. At</p><p>beginning, we controlled the system temperature by adding sulfuric acid</p><p>very slowly and cooling test tubes in ice bath in a time-consuming</p><p>manner. To improve efficiency, we designed a premix reagent by</p><p>mixing sulfuric acid with water in ice bath followed by the addition of</p><p>pure phenol into the cool acid solution. After one-step addition of pre-</p><p>mix reagent into the sample, the mixtures were incubated in boiling</p><p>water bath to develop the color. With this method, the color intensities</p><p>were very reproducible and were comparable with those by adding</p><p>sulfuric acid and phenol separately or with different speeds (Fig. 3).</p><p>In the conventional phenol‑sulfuric acid method, the sample-phe-</p><p>nol‑sulfuric acid mixture usually needs to be incubated for 10 to 20 min</p><p>in a water bath at 25 ◦C to 30 ◦C or room temperature [10], or incubated</p><p>for 30 min in 80 ◦C or 100 ◦C water bath [25,30,31], or incubated for 15</p><p>min at higher temperature-110 ◦C [25] to obtain reproducible mea-</p><p>surements. In order to identify the influence of incubation time and</p><p>further optimize this parameter, we also tested the color changes for</p><p>different incubation times with both conventional addition mode</p><p>described by Dubois et al. [10] and our developed new one-step addition</p><p>method. With the conventional sulfuric acid addition method, the color</p><p>intensity significantly (p < 0.05) decreased in the first 5 min and then</p><p>maintained at a relatively constant value for both high-sugar and low-</p><p>sugar content samples (Fig. 4 A and B). With our new method, the</p><p>color intensity significantly (p < 0.05) increased in first 5 mins and then</p><p>also maintained at a constant value after a 10min incubation (Fig. 4 C</p><p>and D). Therefore, 15 to 20 min was selected as the optimal incubation</p><p>time.</p><p>Fig. 4. Effect of incubation time on the carbohydrate analysis of sugar samples A) low sugar content samples and B) high sugar content samples with conventional</p><p>phenol - sulfuric acid protocol; C) low sugar content samples and D) high sugar content samples with our new method.</p><p>W. Chen et al.</p><p>Algal Research 70 (2023) 102986</p><p>5</p><p>3.2. Causes of low accuracy and influence of algae pigments on the</p><p>quantification of carbohydrate with phenol‑sulfuric acid method</p><p>For the analysis of carbohydrate content in algae and other organ-</p><p>isms, usually carbohydrates were extracted and purified first, and then</p><p>were determined with the phenol‑sulfuric acid method [25,32]. As a</p><p>result, very little information is available on the influence of pigments.</p><p>With the development of algae-derived biofuel research, direct analysis</p><p>of carbohydrates with whole algal biomass became more and more</p><p>necessary. In the literature, several studies used whole algal biomass for</p><p>direct hydrolysis and carbohydrate determinations without considering</p><p>the color interference. However, our preliminary results indicated the</p><p>overestimations of the total carbohydrate content with direct determi-</p><p>nation and quantification. To quantify the potential influences of major</p><p>components of algae cells (lipids, proteins, pigments) on the determi-</p><p>nation of carbohydrates with the phenol‑sulfuric acid method, chloro-</p><p>phyll a, β-carotene, triacylglycerol and BSA were added to D-glucose</p><p>solutions to imitate carbohydrate contents for each composition of 4 %,</p><p>4 %, 50 % and 50 % (% of dry weight), respectively, in algae biomass. At</p><p>the same time, pigments standards without the addition of D-glucose</p><p>were mixed with phenol‑sulfuric acid premix reagent and incubated in</p><p>boiling water for 20 mins, and then spectrometric characters were</p><p>scanned from 300 nm to 750 nm. Results demonstrated that there were</p><p>still some peaks around 490 nm after sulfuric acid treatment for astax-</p><p>anthin, β-carotene and chlorophyll a, but the absorption intensities</p><p>decreased to a great extent compared with original astaxanthin,</p><p>β-carotene standards. For chlorophyll a, there was no absorption peaks</p><p>around 490 nm before sulfuric acid treatment, while small peaks</p><p>occurred at around 480 nm after conducting phenol‑sulfuric acid</p><p>method. These results indicated that the influence of pigments cannot be</p><p>ignored in carbohydrates quantification</p><p>with this colorimetric method,</p><p>as algae biomass usually contains a high content of chlorophylls and</p><p>other pigments.</p><p>Fig. 5 A describes the influence of chlorophylls on the carbohydrate</p><p>determinations in algae biomass with different carbohydrate content. If</p><p>the algal biomass contains 4 % of chlorophylls (% of dry weight), it will</p><p>lead to overestimations of 5 % to ~300 % when carbohydrate content</p><p>ranged from 5 % to 50 % dry weight (Fig. 5). Higher color interferences</p><p>were observed with astaxanthin and β-carotene at even higher carbo-</p><p>hydrate contents, with overestimations ranging between 20 % and ~</p><p>400 % (Fig. 5 B and C). If the algal biomass contains 50 % carbohydrate,</p><p>there is no significant influence on the quantification when β-carotene</p><p>content is below 1.0 %, but higher β-carotene will lead to significant</p><p>overestimations (p < 0.05, Fig. 5 D). Effects of lipids and proteins on the</p><p>quantification of carbohydrates in algae biomass were also investigated</p><p>similarly in this study, but no obvious interferences were observed with</p><p>very high lipid and protein contents (50 % in dry biomass was imitated).</p><p>Many microalgal strains used for biofuel production, could potentially</p><p>accumulate high contents of pigments such as β-carotene, astaxanthin,</p><p>lutein, chlorophylls and so on besides lipids and carbohydrates [3,33].</p><p>The influences of these pigments should be taken into consideration in</p><p>order to obtain accurate results with the phenol‑sulfuric acid method.</p><p>Fig. 5. The influence of chlorophyll (A), β-carotene (B and D), astaxanthin (C) on the carbohydrate determination with phenol-sulfuric acid method. Different</p><p>concentration of D-glucose was used as samples to imitate the algae biomass hydrolysate containing different concentration of total carbohydrates.</p><p>W. Chen et al.</p><p>Algal Research 70 (2023) 102986</p><p>6</p><p>3.3. Hydrolysis of total carbohydrates in whole algal biomass into simple</p><p>sugars for subsequent phenol‑sulfuric acid determination</p><p>For total carbohydrate analysis in algae, algae biomass is usually</p><p>extracted and purified, and is followed by the determination using</p><p>phenol‑sulfuric acid method [34]. Several early papers reported the</p><p>determination of carbohydrates in whole algae biomass using acid hy-</p><p>drolysis [35–37], and other recent investigations simply cited the hy-</p><p>drolysis protocols from these references [19]. For the hydrolysis of</p><p>carbohydrates into simple sugars from alga and other organisms, several</p><p>acids with different concentrations and different hydrolysis tempera-</p><p>tures or time were involved, including 0.05 M–12 M sulfuric acid</p><p>incubated at 0–100 ◦C for 2 h to 24 h [35–38], hydrochloric acids, tri-</p><p>chloroacetic acids, acetic acid and trifluoroacetic acid (TFA) varied in</p><p>different concentrations and incubated in boiling water for 1 h to 4 h</p><p>[12,39–41]. In our preliminary studies, we conducted algal hydrolysis</p><p>with trifluoroacetic acid and hydrochloric acid in boiling water for 4 h.</p><p>Unfortunately, there were significant variations in color intensity in the</p><p>subsequent carbohydrate determination with the phenol‑sulfuric acid</p><p>method. In order to better understand the influence of acid species,</p><p>concentrations and hydrolysis time on the hydrolysis process with whole</p><p>algae biomass, the commonly-used acids and hydrolysis protocols were</p><p>further compared and optimized. Results from Fig. 6 indicated that</p><p>hydrolyzing algal biomass at 100 ◦C for 4 h and 2 h, respectively, with 4</p><p>M TFA and 4 M HCl gave the highest color intensities. No hydrolysis step</p><p>or hydrolyzing algae biomass in sulfuric acid gave lower absorbance.</p><p>Moreover, hydrolysis with too low or too high concentration (less or</p><p>over than 4 M) of HCl also decreased the color intensity significantly (p</p><p>< 0.05) and determination of carbohydrates with TFA could improve</p><p>reproducibility greatly. Microalgae, especially green algae which usu-</p><p>ally has thick and rigid cell walls, using too weak or diluted acids may</p><p>not destroy the cell walls and cell membranes completely and hydrolyze</p><p>all the intracellular carbohydrates into simple sugar. Therefore, the first</p><p>requirement for the selection of hydrolysis acid is the strong acidity.</p><p>Only strong acids, such as sulfuric acid, HCl and TFA work well in hy-</p><p>drolyzing the whole algal biomass. On the other hand, another impor-</p><p>tant aspect – oxidizability of the hydrolysis acid should also be taken</p><p>into consideration. As the produced aldopentose and aldohexose has</p><p>strong reductibility and is easy to be oxidized into corresponding aldonic</p><p>acids, there was significantly decreased color intensity in the subsequent</p><p>carbohydrates measurements. TFA has no oxidizability and HCl has</p><p>weak oxidizability, but sulfuric acid has very strong oxidizability. In</p><p>addition, oxidizability increased dramatically with the increase of</p><p>temperature and acid concentrations. Therefore, the hydrolysis of algal</p><p>biomass with concentrated sulfuric acid in early literatures was always</p><p>performed at room temperature or around 0 ◦C [22,36–38]. Based on the</p><p>above reasons, TFA was the best choice and HCl could also be used for</p><p>the hydrolysis of algal biomass, but hydrolysis time and acid concen-</p><p>tration should be controlled strictly.</p><p>3.4. Quantification of carbohydrates and performance of the improved</p><p>phenol‑sulfuric acid method</p><p>As the algal pigments have significant influence on carbohydrate</p><p>determination with the phenol‑sulfuric acid method [32,42], color in-</p><p>terferences caused by algal pigments should be taken into account in the</p><p>course of quantification with standard curves. In the early literatures,</p><p>determination and quantification were conducted directly after algal</p><p>biomass was hydrolyzed [36,37]. According to our results, this will</p><p>significantly overestimate total carbohydrate content. To eliminate or</p><p>reduce the potential overestimation and obtain more accurate/reliable</p><p>results, we investigated strategies related to elimination of color in-</p><p>ferences by pre-extraction algal pigments and using algal hydrolysates</p><p>as control in the quantification process. It was indicated from Fig. 7 that</p><p>direct quantification with algal hydrolysates containing pigments yiel-</p><p>ded significantly (p < 0.05) higher carbohydrate contents than the pre-</p><p>extracted pigment groups for all five green algal species which varied in</p><p>total carbohydrate contents, and pigment species and concentrations.</p><p>There are no visible differences in total carbohydrate content between</p><p>pigment pre-extraction groups and groups using algal hydrolysates as a</p><p>Fig. 6. Hydrolysis of total carbohydrates in algal whole biomass into simple</p><p>sugars with different acids and hydrolysis conditions A) 4 M TFA 4 h 100 ◦C; B)</p><p>2 M HCl 4 h 100 ◦C; C) 4 M HCl 2 h 100 ◦C; D) 4 M HCl 4 h 100 ◦C; E) 6 M HCl</p><p>2 h 100 ◦C; F) 6 M HCl 4 h 100 ◦C; G) pretreatment with 12 M HCl for 2 h at</p><p>room temperature, then dilute to 4 M and incubated at 100 ◦C for another 2 h;</p><p>H) 80 % H2SO4 for 20 h at room temperature; I) no hydrolysis process.</p><p>Fig. 7. Quantification of total carbohydrates in five algae biomass A) Chlorella</p><p>zofingiensis 1; B) Chlorella zofingiensis 2; C) Scenedesmus sp.; D) Nannochloropsis</p><p>sp.; E) Haematococcus pluvialis. With pigments: biomass was hydrolyzed and</p><p>quantified directly; without pigments: biomass was pre-extracted to remove</p><p>pigments and then use the pigment-free biomass for hydrolysis and quantifi-</p><p>cation; quantification using control: biomass was hydrolyzed directly, but hy-</p><p>drolysates containing pigments was used as control in the following total</p><p>carbohydrate analysis and quantification with phenol‑sulfuric acid method.</p><p>Table 1</p><p>Correlation coefficient, repeatability, reproducibility and limit of detection</p><p>(LOD) the improved method.</p><p>Correlation</p><p>coefficient</p><p>(R [2])</p><p>Repeatability</p><p>(RSD %)</p><p>Reproducibility</p><p>(RSD %)</p><p>Detectable range</p><p>(nmol/well)</p><p>150 μg</p><p>mL− 1</p><p>15 μg</p><p>mL− 1</p><p>150 μg</p><p>mL− 1</p><p>15 μg</p><p>mL− 1</p><p>0.9998 1.6 3.0 1.6 5.0 5–150</p><p>n = 10.</p><p>W. Chen et al.</p><p>Algal Research 70 (2023) 102986</p><p>7</p><p>control in the quantification process. Therefore, both these two methods</p><p>could eliminate or reduce the potential overestimation of carbohydrate</p><p>content. To assess the performance of the improved method, the corre-</p><p>lation coefficient, repeatability, reproducibility and linear range were</p><p>further investigated and results are indicated in Table 1. A correlation</p><p>coefficient of R2 = 0.9998 (Table 1) between the absorbance intensities</p><p>at 490 nm and total carbohydrate contents was obtained using the</p><p>improved method. Based on the standard curve generated, the linearity</p><p>ranged from 1 μg mL− 1 to 150 μg mL− 1 (equal to 5 nmol/well to 150</p><p>nmol in 96 well plates) for D-glucose. Repeated measurements for</p><p>different algae strains or the same strain with different carbohydrate</p><p>concentrations, gave very reproducible results, with relative standard</p><p>errors of 1.6, 3.0 and 1.6, 5.0 % for repeatability and reproducibility at</p><p>two concentration levels (150 μg mL− 1 and 15 μg mL− 1), respectively.</p><p>The improved method is reproducible/repeatable and accurate/reliable</p><p>in comparison to the original phenol‑sulfuric acid method described by</p><p>Dubois and his colleagues and its improved methods in the past-half</p><p>century.</p><p>3.5. Established standard operation protocols for carbohydrate analysis</p><p>in algal biomass</p><p>Based on investigations from this study, the detailed established</p><p>standard operation protocols (SOP) for analyzing carbohydrates in algal</p><p>biomass were proposed in Fig. 8. The SOP method will provide useful</p><p>information for the determination of carbohydrates, not only to scien-</p><p>tists in the area of algal studies but the areas of other type of carbohy-</p><p>drates as well.</p><p>4. Conclusion</p><p>Temperature is the key parameter that affects the color development</p><p>process. Using a premixed reagent instead of the conventional mode for</p><p>the addition of sulfuric acid and phenol could greatly improve the var-</p><p>iations in color intensities. Color interference is another crucial factor</p><p>that influenced the accuracy of quantification. Potential overestimation</p><p>could be reduced or eliminated by using pigment pre-extraction or using</p><p>the algal hydrolysates as a control during the quantification process. In</p><p>addition, selections of an appropriate acid, acid concentration and hy-</p><p>drolysis temperature/time are also very important to ensure accurate</p><p>measurements of total carbohydrates in algal biomass. The modified</p><p>phenol‑sulfuric acid method has been shown to be more reproducible</p><p>and reliable than the conventional methods.</p><p>CRediT authorship contribution statement</p><p>Wei Chen: Conceptualization, Methodology, Formal analysis,</p><p>Investigation, Supervision, Writing – original draft, Writing – review &</p><p>editing, Funding acquisition. Lei Gao: Software, Methodology, Formal</p><p>analysis, Investigation, Writing – original draft. Lirong Song: Valida-</p><p>tion, Resources, Visualization. Milton Sommerfeld: Writing – original</p><p>draft, Writing – review & editing, Project administration, Validation.</p><p>Qiang Hu: Conceptualization, Data curation, Supervision, Resources,</p><p>Project administration, Writing – original draft, Writing – review &</p><p>editing.</p><p>Declaration of competing interest</p><p>The authors declare that they have no known competing financial</p><p>interests or personal relationships that could have appeared to influence</p><p>the work reported in this paper.</p><p>Data availability</p><p>Data will be made available on request.</p><p>Acknowledgments</p><p>The research was supported by Shandong Provincial Natural Science</p><p>Foundation (No. ZR2020MD086), High level Talents Project of Qingdao</p><p>Agricultural University (No.665/1119022) and National Natural Sci-</p><p>ence Foundation of China (NSFC 41276150).</p><p>Fig. 8. Established standard operation protocols for analyzing carbohydrates in algal biomass.</p><p>W. 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improved phenol-sulfuric acid method for the quantitative measurement of total carbohydrates in algal biomass</p><p>1 Introduction</p><p>2 Material and methods</p><p>2.1 Organisms and culture conditions</p><p>2.2 Chemicals and reagents</p><p>2.3 Effect of incubation temperatures and time on the color development for the phenol-H2SO4 method</p><p>2.4 Effect of pigments, lipids and proteins on the phenol-H2SO4 method</p><p>2.5 Spectrophotometric determinations of carbohydrates</p><p>2.6 Effects of algae pigments on the quantification of total carbohydrates in algae biomass</p><p>2.7 Hydrolysis of algae biomass</p><p>2.8 Statistical analysis</p><p>3 Results and discussion</p><p>3.1 Causes of low reproducibility/repeatability and influence of temperature in conventional phenol‑sulfuric acid method</p><p>3.2 Causes of low accuracy and influence of algae pigments on the quantification of carbohydrate with phenol‑sulfuric acid ...</p><p>3.3 Hydrolysis of total carbohydrates in whole algal biomass into simple sugars for subsequent phenol‑sulfuric acid determi ...</p><p>3.4 Quantification of carbohydrates and performance of the improved phenol‑sulfuric acid method</p><p>3.5 Established standard operation protocols for carbohydrate analysis in algal biomass</p><p>4 Conclusion</p><p>CRediT authorship contribution statement</p><p>Declaration of competing interest</p><p>Data availability</p><p>Acknowledgments</p><p>References</p>