MK and TB thanks Haitham Ayeb for statistical analyses and figure preparation, Walloon Region (European Regional Development-VERDIR) and Minister of Higher Education and Scientific Research (Taoufik Bettaieb) for funding.

1. Preparation of biomasses
<ol>
	<li>Biomass drying
	<ol>
		<li>Place the Posidonia leaves and aegagropile balls (Posidonia oceanica), harvested from Mediterranean beaches, in an oven at 40 &#176;C for 72 h.</li>
		<li>Place the almond shells (Prunus dulcis), generated from food industries, and olive pomace (Olea europaea L.), obtained from olive oil mills, in an oven at 40 &#176;C for 72 h.</li>
		<li>Place the pinecones (Pinus halepensis), collected from a forest, and alfa leaves (Stipa tenacissima), collected from the southern Mediterranean basin, in an oven at 40 &#176;C for 72 h. 
		&#8203;NOTE: If the biomass contains sand, it must be rinsed with distilled water before it is placed in the oven. The biomasses are shown in Figure 1A-F.</li>
	</ol>
	</li>
	<li>Biomass grinding
	<ol>
		<li>Place 20 g of each biomass in a hammer cutter equipped with a 1 mm sieve. Collect the resultant powder in a 0.25 L beaker and feed it to a hammer cutter equipped with a 0.5 mm sieve. Collect the powder in a 0.25 L beaker.</li>
	</ol>
	</li>
</ol>
2. Microwave-assisted, ultrafast lignin extraction
<ol>
	<li>Deep eutectic solvent (DES) preparation
	<ol>
		<li>Prepare DES1 (ChCl-oxalic acid) in a molar ratio of 1:1 by mixing 174 g of ChCl with 126 g of oxalic acid dihydrate in a 500 mL round-bottom flask and melting them in a bath at 70 &#176;C for 4 h until a homogenous and transparent liquid is formed.</li>
		<li>Prepare DES2 (ChCl-lactic acid) in a molar ratio of 1:1 by mixing 174 g of ChCl with 90 g of lactic acid in a 500 mL round-bottom flask and melting them in a bath at 70 &#176;C for 4 h until a homogenous and transparent liquid is formed.</li>
		<li>Prepare DES3 (ChCl-urea) in a molar ratio of 1:12 by mixing 174 g of ChCl with 120 g of urea in a 500 mL round-bottom flask and melting them in a bath at 70 &#176;C for 4 h until a homogenous and transparent liquid is formed. 
		NOTE: Stir these mixtures continuously with a stir bar at 500 rpm.</li>
	</ol>
	</li>
	<li>Combined microwave-DES treatment
	<ol>
		<li>Place 5 g of the feedstock in a microwave in a closed polytetrafluoroethylene reactor. Add 50 mL of DES, and place a stirring bar in the sample. Close the microwave container with an appropriate cap, and attach the temperature cap.</li>
		<li>Place the microwave container on the edge of the turntable, ensuring that it is constantly agitated. Set the microwave power at 800 W for 1 min. Using suitable gloves, take the container out of the microwave, and let the mixture cool. Repeat this treatment using the three DESs for each biomass sample. 
		&#8203;NOTE: Check and ensure that the temperature probe is correctly placed, and the microwave container has a homogenous temperature.</li>
	</ol>
	</li>
	<li>Lignin isolation
	<ol>
		<li>Prepare a homogeneous antisolvent solution by mixing ethanol:water in a 50:50 (v:v) ratio. Add 50 mL of the antisolvent solution to the treated feedstock, place the mixture in a centrifugation container (250 mL), and centrifuge for 5 min at 3,000 &#215; g.</li>
		<li>After centrifugation, filter the supernatant (lignin-rich fraction) using a glass filter crucible (porosity 4, 10-16 &#956;m, diameter 10 mm). Wash the remaining cellulose residue collected after centrifugation with 25 mL of the antisolvent solution.</li>
		<li>Centrifuge at 3,000 &#215; g for 5 min after each wash. Repeat washes 4x, and collect and filter the washes through the glass filter crucible (porosity N 4, 10-16 &#956;m, diameter 10 mm).</li>
		<li>Add the filtered lignin-rich fraction from step 2.3.2 to the filtered washes from step 2.3.3 in a 500 mL round bottom flask. Evaporate ethanol using a rotary evaporator at 50 &#176;C and 110 mbar.</li>
		<li>Add 150 mL of deionized water to the concentrated liquor (lignin-rich fraction), and precipitate the lignin by centrifugation. Collect lignin as a pellet, and wash it with 25 mL of distilled water; repeat the washes 4x. Lyophilize lignin, or dry it in an oven at 40 &#176;C. 
		&#8203;NOTE: If necessary, wash lignin &#62;4x to remove the salts from the solvents.</li>
		<li>Use the following formula to determine the yield: 
		<img alt="Equation 1" src="/files/ftp_upload/61997/61997eq01.jpg" /> 
		NOTE: Lignin extraction was also performed with two other DESs: choline chloride + resorcinol and choline chloride + butyric acid at 1 min. However, the amounts of lignin recovered using these DESs were extremely small (and unrecoverable) compared with the amounts obtained using the other three DESs.</li>
	</ol>
	</li>
</ol>
3. Purity determination of extracted lignin by Klason
<ol>
	<li>Sample preparation for Klason hydrolysis
	<ol>
		<li>Place the filter crucible (porosity 4, diameter 4.5 mm) in a muffle furnace at 550 &#176;C for 4 h (2 h ramp, from 25&#176;C). Remove the crucible when the oven cools to 150 &#176;C, place it in a desiccator to cool, and weigh.</li>
		<li>Add approximately 30 mg of lignin into a borosilicate glass tube (see the Table of Materials), and note the weight of the sample. Add 1 mL of 72% sulfuric acid (H2SO4) to the sample, place the sample in a 30 &#176;C bath for 60 min, and mix every 10 min by vortexing.</li>
		<li>Remove the sample, transfer it to a 100 mL glass bottle, and add 28 mL of distilled water to dilute the acid to a concentration of 4%. Place the glass bottle in an autoclave at 121 &#176;C for 60 min. Remove the glass bottle, and allow it to cool.</li>
	</ol>
	</li>
	<li>Analysis of acid-insoluble lignin
	<ol>
		<li>Filter the hydrolysate using a crucible under vacuum. Collect all the solids in a glass bottle containing deionized water. Rinse the crucible with 50 mL of deionized water.</li>
		<li>Dry the crucible containing the solids by placing it in an oven at 105 &#176;C for 16 h. Remove the crucible from the oven, place it in a desiccator, and allow it to cool. Weigh the sample.</li>
		<li>Place the crucible in a muffle furnace at 550 &#176;C for 4 h (2 h ramp). Remove it and place in a desiccator. Weigh the sample.</li>
		<li>Use the following formula to calculate the percentage of the acid-insoluble residue (AIR): 
		<img alt="Equation 2" src="/files/ftp_upload/61997/61997eq02v2.jpg" /> 
		WCSA: weight of crucible + sample after removing them from the oven 
		WC: weight of crucible 
		WCSMF: weight of crucible after removing it from the muffle furnace 
		ODW: oven dry weight of the sample</li>
	</ol>
	</li>
	<li>Analysis of acid-soluble lignin
	<ol>
		<li>Measure the absorbance of the filtrate obtained in step 3.2.1 with a spectrophotometer at 205 nm using quartz cuvettes. Use distilled water as blank.</li>
		<li>Use the following formula to calculate the percentage of the acid-soluble residue (ASL): 
		<img alt="Equation 3" src="/files/ftp_upload/61997/61997eq03.jpg" /> 
		NOTE: Absorbance should be between 0.2 and 0.7. Dilute the sample if necessary. 
		UVabs: absorbance at 205 nm 
		Pathlength: light path of the measuring cell (in cm) 
		&#949;:&#160;absorbency of biomass at a specific wavelength</li>
	</ol>
	</li>
</ol>
4. Nitrogen content in extracted lignin
<ol>
	<li>Preparation of alkali solution
	<ol>
		<li>In a 2.5 L volumetric flask, weigh 1 kg of sodium hydroxide (NaOH) and add deionized water up to the mark. Place a magnetic bar in the flask, and stir until the NaOH is completely dissolved.</li>
	</ol>
	</li>
	<li>Sulfuric acid solution preparation
	<ol>
		<li>Take 0.1 N H2SO4 (see the Table of Materials) in a 5 L volumetric flask, add deionized water up to the 5 L mark, place a magnetic bar, and stir until the contents dissolve.</li>
	</ol>
	</li>
	<li>Preparation of receiving solution
	<ol>
		<li>In a 5 L volumetric flask, dissolve 100 g of H3BO3 (boric acid) in deionized water, and bring up the volume to the mark.</li>
		<li>Weigh 100 mg of bromocresol green in a 100 mL volumetric flask, and add technical methanol up to the mark.</li>
		<li>Weigh 100 mg of methyl red in a 100 mL volumetric flask, and add technical methanol up to the mark.</li>
		<li>Pour the 5 L of H3BO3 solution from step 4.3.1, 100 mL of bromocresol green solution from step 4.3.2, 70 mL of the methyl red solution from step 4.3.3, and 5 L of deionized water into a container. Shake the receiving solution well for 30 min. 
		&#8203;NOTE: The final color of the solution must be green. If the color is not green, add 50 mL of 1 N NaOH solution.</li>
	</ol>
	</li>
	<li>Sample preparation
	<ol>
		<li>In a Kjeldahl tube, place 100 mg of lignin weighed on a nitrogen-free paper, add a tablet of Kjeldhal (1.5 g potassium sulfate (K2SO4) + 0.045 g copper sulfate pentahydrate (CuSO4.5H2O) + 0.045 g titanium dioxide (TiO2)), and add 7.2 mL of concentrated H2SO4. 
		NOTE: Use four tubes with only nitrogen-free paper (without the samples) as blanks.</li>
	</ol>
	</li>
	<li>Sample digestion
	<ol>
		<li>Switch on the thermostat on the digester 1 h in advance at 360 &#176;C.</li>
		<li>Place the sample tubes on a rack, place the four blank tubes at the four corners of the rack, and fill in the holes (if any) of the rack with empty tubes.</li>
		<li>Place the rack in the preheated digester, cover the suction system, and open the water pump. 
		NOTE: Take care to avoid fumes; increase the flow of water if fumes appear.</li>
		<li>After 2 h, turn off the heating, remove the samples, and place them on a metal support. Allow the rack to cool for approximately 40 min with the suction system on.</li>
	</ol>
	</li>
	<li>Kjeldhal distillation procedure
	<ol>
		<li>Switch on the Kjeldahl distiller. Allow Self-tests to run until Selection appears on the screen. Switch to Manual mode, insert an empty tube, and close the sliding door.</li>
		<li>Purge the titrant burette (0.02 N H2SO4) (lift the cover) by pressing it at the bottom and top several times, and eliminate air bubbles from the pipes by squeezing the tube of the H2SO4 bottle. Close the hood.</li>
		<li>Purge the H3BO3 receiving solution 3x.</li>
		<li>Add water 3x, and switch to Active steam (10 min). Switch to the Kjeldahl 1 analysis program. Enter Blanco using the arrows at the Result line level.</li>
		<li>Insert the tube. Start with the four blanks, and calculate their averages. Enter the value in the Blanco line. 
		NOTE: After the tube is inserted, the device automatically and successively adds 30 mL of H2O, 30 mL of H3BO3, and 40 mL of 10 N NaOH.</li>
		<li>Switch to mL of titrant at the result line. Insert the tube, and note the amount of H2SO4 used. 
		NOTE: To test the Kjeldahl distiller, consider that 50 mg of glycerin correspond to 18.60% &#177; 5% of % N. At the end of each titration, the device automatically empties and cleans the tube.</li>
		<li>Calculate the percentage of N. 
		<img alt="Equation 4" src="/files/ftp_upload/61997/61997eq04.jpg" /> 
		V s.a : Volume of sulfuric acid 
		T s.a : 0.02 N H2SO4 
		S: sample mass</li>
	</ol>
	</li>
</ol>
5. Ash content in extracted lignin
<ol>
	<li>Dry the ceramic crucibles for 1 h at 105 &#176;C. Leave them to cool in a desiccator.</li>
	<li>Weigh a crucible, and note its number. Add approximately 1 g of the sample powder. Place the crucible in the muffle furnace with the following program: a 2 h ramp up to 575 &#176;C; a plateau of 4 h at 575 &#176;C.</li>
	<li>Allow the oven to cool to 100 &#176;C. Remove the crucibles, place them in the desiccator, and weigh them.</li>
</ol>
6. Carbohydrate content
<ol>
	<li>Preparation of sodium borohydride (NaBH4)/dimethylsulfoxide (DMSO) solution
	<ol>
		<li>Place 2 g of NaBH4 in a 100 mL volumetric flask, and fill to the mark with DMSO. Heat to 100 &#176;C in a Mayor&#39;s bath, and stir the solution until completely dissolved.</li>
	</ol>
	</li>
	<li>Preparation of MIX solution
	<ol>
		<li>Place 20 mg each of xylose, arabinose, rhamnose, glucose, galactose, mannose, and 2-deoxyglucose in a 100 mL volumetric flask, and fill up to the mark of 100 mL with deionized water.</li>
	</ol>
	</li>
	<li>Hydrolysis of the sample
	<ol>
		<li>Weigh a 50 mg sample of lignin in a borosilicate glass tube, add 3 mL of 1 M H2SO4, and heat the mixture for 3 h at 100 &#176;C.</li>
		<li>Cool the sample, add 1 mL of 15 M ammonium hydroxide (NH4OH), and check the pH to ensure that it is neutral or alkaline. Add exactly 1 mL of internal standard (2-deoxyglucose) to each sample. 
		NOTE: The 2-deoxyglucose added as an internal standard makes it possible to quantify the quantity of each dose present in the sample.</li>
	</ol>
	</li>
	<li>Reduction and acetylation of monosaccharides into alditol acetate
	<ol>
		<li>Take 400 &#181;L of the solution from step 6.3.2, and place it in special tubes. Take 400 &#181;L of the control MIX solution, and place it in special tubes. 
		NOTE: Using the MIX solution facilitates the calculation of response factors (RFs) and monosaccharide percentages.</li>
		<li>Add 2 mL of the NaBH4/DMSO solution prepared in section 6.1. Close the tube, and incubate for 90 min at 40 &#176;C in a water bath. Remove the tube from the water bath, and add 0.6 mL of glacial acetic acid. 
		&#8203;NOTE: As this is an exothermic reaction, bubbles and smoke will appear.</li>
		<li>Add approximately 0.4 mL of 1-methylimidazole and approximately 4 mL of acetic anhydride. After 15 min, add 10 mL of distilled water, cool, and add ~3 mL of dichloromethane (CH2Cl2).</li>
		<li>After at least 2 h, collect ~1 mL of the lower (organic) phase, and inject it into a gas chromatograph equipped with a flame ionization detector capillary column, HP1-methylsiloxane (30 m (length) x 320 &#181;m (internal diameter), 0.25 &#181;m (film thickness)). Analyze the data.</li>
		<li>Use the following formula to calculate the response factor (RF). 
		<img alt="Equation 5" src="/files/ftp_upload/61997/61997eq05v2.jpg" /> 
		A m. p: Average of the area of the monosaccharide peak in the MIX solution 
		M a. h of 2 - deoxy glucose: Mass of 2-deoxyglucose after hydrolysis 
		A 2dg. p: Average of the area of the 2-deoxyglucose peak in the MIX solution 
		M a. h of the monosaccharide: Mass of the monosaccharide after hydrolysis 
		NOTE: Anhydro correction is 0.8 for rhamnose, 0.88 for arabinose and xylose, and 0.9 for mannose, glucose, and galactose. Mass after hydrolysis = anhydro correction x mass (g) of the monosaccharide used in the MIX solution.</li>
		<li>Use the following formula to calculate the monosaccharide mass. 
		<img alt="Equation 6" src="/files/ftp_upload/61997/61997eq06.jpg" /> 
		AP.M: Monosaccharide peak area in the analyzed sample 
		M. IS: Mass of internal standard added; here, C SI=1 mg/mL 
		AP.2: Peak area of 2-deoxyglucose in the sample 
		RF: response factor</li>
		<li>Calculate the percentage of each monosaccharide using the following formula. 
		<img alt="Equation 7" src="/files/ftp_upload/61997/61997eq07v2.jpg" /></li>
	</ol>
	</li>
</ol>
7. Chemical functions in extracted lignin (Fourier-transformed infrared)
<ol>
	<li>To identify the chemical functional groups in extracted lignin, use an FT-IR spectrometer equipped with an attenuated total reflectance (ATR) module. Open the spectroscopy software, and adjust the parameters: resolution 4 cm-1, sample scan time 32, background scan time 16, save data from 4000 to 400 cm-1, result spectrum transmittance.</li>
	<li>Do not add any sample; press background single channel. Now place 1 mg of the sample on the crystal, and press sample single channel. Process the obtained spectra.</li>
</ol>
8. Molecular weight of extracted lignin (gel permeation chromatography)
<ol>
	<li>Prepare a solution of dimethylformamide (DMF) with 0.5% lithium chloride (LiCl). Take 5 g of LiCl in a 1 L volumetric flask, add DMF up to the gauge line, and mix the contents until a homogenous liquid is obtained.</li>
	<li>Dissolve 3 mg of the lignin sample in 3 mL of DMF with 0.5% LiCl. Centrifuge in a 10 mL centrifuge tube, and separate the soluble fraction into a vial.</li>
	<li>Dissolve 3 mg of polystyrene standard 1 kDa, 2 kDa, 3 kDa, 10 kDa, 20 kDa, and 30 kDa in the solution of DMF with 0.5% LiCl. Centrifuge in 10 mL borosilicate glass tubes, and transfer the soluble fraction to a vial.</li>
	<li>Prepare a high-performance liquid chromatography-ultraviolet (UV) system.
	<ol>
		<li>Open the data system, and check the UV detector.</li>
		<li>Purge the system with distilled water. Install the plunger in the eluent (DMF with 0.5% LiCl). Open the purging valve, and purge the line with a flow rate of 1 mL/min for 15 min. Stop the flow and close the purging valve.</li>
		<li>Set the flow rate to 1 mL/min for 10 min to clean the eluent pathway to the detector. Stop the flow rate.</li>
		<li>Install the column preceded by a guard column (see the Table of Materials). Switch on the column heater at 45 &#176;C, switch on the UV detector, and set the flow rate gradually until a flow rate of 0.6 mL/min is reached.</li>
		<li>Inject 30 &#181;L of each sample for 40 min at a wavelength of 270 nm. Process the obtained data, and calculate the mass distribution using the calibration line.</li>
		<li>Calculate the number average molecular weight (Mn), weight average molecular weight (Mw), and polydispersity index (PDI). 
		<img alt="Equation 8" src="/files/ftp_upload/61997/61997eq08.jpg" /> 
		<img alt="Equation 9" src="/files/ftp_upload/61997/61997eq09.jpg" /> 
		<img alt="Equation 10" src="/files/ftp_upload/61997/61997eq10.jpg" /> 
		Mi: molecular weight of a chain 
		Ni: number of chains for that molecular weight</li>
	</ol>
	</li>
</ol>
9. Data treatment and statistical analyses
<ol>
	<li>Perform all analytical experiments in triplicate and express the results as % of dry matter.</li>
	<li>Perform one-way analysis of variance (ANOVA), and compare the means using Tukey&#39;s multiple comparison test.</li>
	<li>Perform principal component analysis (PCA) .</li>
</ol>

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M., Weon Choi, J., Won, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lignin+for+white+natural+sunscreens.">Lignin for white natural sunscreens.</a> International Journal of Biological Macromolecules. 122, 549-554 (2019).</li><li>Widsten, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lignin-based+sunscreens-state-of-the-art,+prospects+and+challenges.">Lignin-based sunscreens-state-of-the-art, prospects and challenges.</a> Cosmetics. 7, 85 (2020).</li><li>Qian, Y., Qiu, X., Zhu, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lignin:+a+nature-inspired+sun+blocker+for+broad-spectrum+sunscreens.">Lignin: a nature-inspired sun blocker for broad-spectrum sunscreens.</a> Royal Society of Chemistry. 17, 320-324 (2015).</li><li>Zijlstra, D. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extraction+of+lignin+with+high+&#946;-O-4+content+by+mild+ethanol+extraction+and+its+effect+on+the+depolymerization+yield.">Extraction of lignin with high &#946;-O-4 content by mild ethanol extraction and its effect on the depolymerization yield.</a> Journal of Visualized Experiments. (143), e58575 (2019).</li></ol>

The authors report no conflict of interest.

This study had many objectives; the first of which was to prepare and use low-cost green solvents with the characteristics of both ionic liquids and organic solvents. The second objective was to fractionate the biomass and extract lignin in a single step, without requiring preliminary steps such as the extraction of extractables using Soxhlet or hemicellulose using alkaline solvents, basic, or thermophysical techniques. The third aim was to recover lignin by simple filtration after the treatment, without adjustment of pH, but simply by adding distilled water. The results of the ultrafast extraction of lignin from six different sources using the microwave-assisted, DES-based process using three different DESs indicate that the extraction yield can vary depending on the biomass and nature of the DES. For example, the highest yield of lignin extraction among all three DESs was from olive pomace. This was followed by the yields from alfa leaves, pinecones, and almond shells. The extraction yields were lower for the leaves and balls of Posidonia oceanica.
The purity of lignin was evaluated using the Klason, Kjeldahl (nitrogen), carbohydrate (GC), and ash methods. As depicted in Figure 3 and Figure 5A-C, the purity of lignin decreased owing to the co-precipitation of nitrogen, carbohydrate, and ash components with lignin. The conditions for lignin extraction with DES1 ensured high purity, but a low yield, indicating that process improvements are necessary for the positive correlation between the yield and purity of lignin. The lignin yield can be improved if the duration of the treatment is longer, the microwave power is increased from 800 W to 1200 W, or the ratio of solid:solvent (1:10) is reduced. Lignin molecular weight data provide an insight into the dissociation or repolymerization of lignin fragments after treatment. An increase in the Mw of lignin for the biomasses was observed after the extraction using microwave-DES, as is evident, for example, in the case of Posidonia leaves (the Mw is 50093 for DES3 and is 70726 for DES1), which demonstrates that depolymerization occurred during the extraction of lignin and was followed by a rapid repolymerization of the carbon-carbon interunit under the action of the DES. This requires the use of a capturing agent, such as formaldehyde, to stabilize the deployment.
In DES pretreatment, lignin dissociation and condensation are the two competing reactions. The PDI of the extracted lignins is lower than that of beech lignin extracted by organic solvents (ethanol/water/H2SO4) reported in the literature17. This indicates that DES treatment improves molecular weight homogeneity in lignin compared to treatment with organic solvents. The FTIR spectra indicate that lignin functional groups are influenced by the DES solvent used. Spectra show signals at 1,730-1,702 cm-1 assigned to the stretching vibration of unconjugated carbonyl groups, whereas peaks at 1,643-1,635 cm-1 indicate the stretching vibration of conjugated carbonyl groups. These results demonstrate the possibility of extracting value-added lignin of high purity from Mediterranean biomasses (which is presently undervalued and used either as feed or as soil amendment) and can help determine the optimal DES solvent while ensuring the purity of lignin. For example, DES1 demonstrated the purest extraction of lignin, though with a lower yield than that observed using the other two DESs.
The proposed method can be applied easily because of the inexpensive and green ChCl-oxalic acid deep eutectic solvent system. Choline chloride is an organic salt and oxalic acid is available as a natural product of plants, which are abundant with low cost. This technique (an ultrafast protocol, which in one step provides biomass fractionation and high-purity lignin recovery) is applicable to any type of lignocellulosic biomass that has a chemical composition similar to that studied here on the laboratory scale using the microwave-DES process or at the pilot scale using the DES-ultrasound process or by convectional heating.

Sustainable biorefinery processes integrate biomass processing, its fractionation into molecules of interest, and their conversion to value-added products1. In second-generation biorefining, pretreatment is considered essential for fractionating biomass into its main components2. Traditional pretreatment methods utilizing chemical, physical, or biological strategies have been widely applied3. However, such pretreatment is considered the most expensive step in biorefining and has other disadvantages such as long processing time, high heat and power consumption, and solvent impurities4. Recently, DESs, whose properties are similar to those of ionic liquids3, have emerged as green solvents owing to advantages such as biodegradability, environmental-friendliness, ease of synthesis, and recovery after treatment5.
DESs are mixtures of at least one HBD, such as lactic acid, malic acid, or oxalic acid, and a hydrogen-bond acceptor (HBA) such as betaine or choline chloride (ChCl)6. HBA-HBD interactions enable a catalytic mechanism that permits cleavage of chemical bonds, causing biomass fractionation and lignin separation. Many researchers have reported the DES-based pretreatment of lignocellulosic feedstocks such as ChCl-glycerol on corn&#39;s cob and stover7,8, ChCl-urea, and ChCl-oxalic acid on wheat straw9, ChCl-lactic acid on Eucalyptus sawdust10, and ChCl-acetic acid11 and ChCl-ethylene glycol on wood11. To improve DES efficiency, the pretreatment should be combined with microwave treatment to accelerate biomass fractionation5. Many researchers have reported such a combined pretreatment (DES and microwave) of wood8 and of corn stover, switchgrass, and Miscanthus5, which provides new insight into the capacity of DESs for lignocellulosic fractionation and lignin extraction in one easy step over a short period.
Lignin is a phenolic macromolecule valorized as a raw material for the production of biopolymers and presents an alternative for the production of chemicals such as aromatic monomers and oligomers12. In addition, lignin has antioxidant and ultraviolet absorption activities13. Several studies have reported lignin applications in cosmetic products14,15. Its integration in commercial sunscreen products has improved the sun protection factor (SPF) of the product from SPF 15 to SPF 30 with the addition of only 2 wt % lignin and up to SPF 50 with the addition of 10 wt % lignin16. This paper describes an ultrafast approach for lignin-carbohydrate cleavage, assisted by combined DES-microwave pretreatment of Mediterranean biomasses. These biomasses consist of agri-food byproducts, particularly olive pomace and almond shells. Other biomasses that were investigated were those of a marine origin (Posidonia leaves and aegagropile) and those originating from a forest (pinecones and wild grasses). The focus of this study was to test low-cost green solvents to evaluate the effects of this combined pretreatment on feedstock fractionation, to investigate its influence on lignin purity and yield, and to study its effects on the molecular weights and chemical functional groups in the extracted lignin.

<table><tbody><tr><td>HPLC Gel Permeation Chromatography</td><td>Agilent 1200 series</td><td></td><td></td></tr><tr><td>1 methylimadazole</td><td>Acros organics</td><td></td><td></td></tr><tr><td>2-deoxy-D-glucose (internal standard)</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Acetic acid</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Acetic anhydride</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Adjustables pipettors</td><td></td><td></td><td></td></tr><tr><td>Alkali</td><td></td><td></td><td>alkali-extracted lignin</td></tr><tr><td>Arabinose (99%)</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Autoclave</td><td>CERTO CLAV (Model CV-22-VAC-Pro)</td><td></td><td></td></tr><tr><td>Water Bath at 70 &amp;deg;C</td><td></td><td></td><td></td></tr><tr><td>Boric acid</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Bromocresol</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Catalyst</td><td>CTQ (coded A22) (1.5 g K&lt;sub&gt;2&lt;/sub&gt;SO&lt;sub&gt;4&lt;/sub&gt; + 0.045 g CuSO&lt;sub&gt;4&lt;/sub&gt;.5 H&lt;sub&gt;2&lt;/sub&gt;O + 0.045 g TiO&lt;sub&gt;2&lt;/sub&gt;)</td><td>Merck</td><td></td></tr><tr><td>Centrifugation container</td><td></td><td></td><td></td></tr><tr><td>Centrifuge</td><td>BECKMAN COULTER</td><td>Avanti J-E centrifuge</td><td></td></tr><tr><td>Ceramic crucibles</td><td></td><td></td><td></td></tr><tr><td>Choline chloride 99%</td><td>Acros organics</td><td></td><td></td></tr><tr><td>Column</td><td>Agilent PLGel Mixed C (alpha 3,000 (4.6 &amp;times; 250 mm, 5 &amp;micro;m) preceded by a guard column (TSK gel alpha guard column 4.6 mm &amp;times; 50 mm, 5 &amp;micro;m)</td><td></td><td></td></tr><tr><td>Column</td><td>HP1-methylsisoxane (30 m, 0.32 mm, 0.25 mm)</td><td></td><td></td></tr><tr><td>Crucible porosity N&amp;deg;4 ( Filtering crucible)</td><td>Shott Duran Germany</td><td>boro 3.3</td><td></td></tr><tr><td>Deonized water</td><td></td><td></td><td></td></tr><tr><td>Dessicator</td><td></td><td></td><td></td></tr><tr><td>Dimethylformamide</td><td>VWR BDH Chemicals</td><td></td><td></td></tr><tr><td>Dimethylsulfoxide</td><td>Acros organics</td><td></td><td></td></tr><tr><td>Erlenmeyer flask</td><td></td><td></td><td></td></tr><tr><td>Ethanol</td><td>Merck (Darmstadtt, Germany)</td><td></td><td></td></tr><tr><td>Filtering crucibles, procelain</td><td></td><td></td><td></td></tr><tr><td>Filtration flasks</td><td></td><td></td><td></td></tr><tr><td>Fourrier Transformed Inra- Red</td><td>Vertex 70 Bruker apparatus&lt;br/&gt; equipped with an attenuated total reflectance (ATR) module.&lt;br/&gt; Spectra were recorded in the 4,000&amp;ndash;400 cm&lt;sup&gt;&amp;minus;1&lt;/sup&gt; range with 32 scans&lt;br/&gt; at a resolution of 4.0 cm&lt;sup&gt;&amp;minus;1&lt;/sup&gt;</td><td></td><td></td></tr><tr><td>Galactose (98%</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Gaz Chromatography</td><td>Agilent (7890 series)</td><td></td><td></td></tr><tr><td>Glass bottle 100 mL</td><td></td><td></td><td></td></tr><tr><td>Glass tubes ( borosilicate) with teflon caps 10 mL</td><td></td><td></td><td></td></tr><tr><td>Glucose (98%</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Golves</td><td></td><td></td><td></td></tr><tr><td>Graduated cylinder 50 mL /100 mL</td><td></td><td></td><td></td></tr><tr><td>H2SO4 Titrisol (0.1 N)</td><td>Merck (Darmstadtt, Germany)</td><td></td><td></td></tr><tr><td>H2SO4 (95-98%)</td><td>Sigma Aldrich (St. Louis, USA)</td><td>BUCHI R-114)</td><td></td></tr><tr><td>Hummer cutter equiped with 1 mm and 0.5 mm sieve</td><td>Mill Ttecator (Sweden)</td><td>Cyclotec 1093</td><td></td></tr><tr><td>Indulin</td><td></td><td></td><td>Raw lignin control</td></tr><tr><td>Kjeldahl distiller</td><td>Kjeltec 2300 (Foss)</td><td></td><td></td></tr><tr><td>Kjeldahl tube</td><td>FOSS</td><td></td><td></td></tr><tr><td>Kjeldhal rack</td><td></td><td></td><td></td></tr><tr><td>Kjeldhal digester</td><td>Kjeltec 2300 (Foss)</td><td></td><td></td></tr><tr><td>Kjeldhal suction system</td><td></td><td></td><td></td></tr><tr><td>Lab Chem station Software</td><td></td><td></td><td>GC data analysis</td></tr><tr><td>Lactic acid</td><td>Merck (Darmstadtt, Germany)</td><td></td><td></td></tr><tr><td>Lithium chloride LiCl</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Mannose (98%)</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Methyl red</td><td></td><td></td><td></td></tr><tr><td>Microwave</td><td>START SYNTH MILESTONE Microwave laboratory system</td><td></td><td></td></tr><tr><td>Microwave temperature probe</td><td></td><td></td><td></td></tr><tr><td>Microwave container</td><td></td><td></td><td></td></tr><tr><td>Muffle Furnace</td><td></td><td></td><td></td></tr><tr><td>NaOH</td><td>Merck (Darmstadtt, Germany)</td><td></td><td></td></tr><tr><td>Nitrogen free- paper</td><td></td><td></td><td></td></tr><tr><td>Opus</td><td></td><td></td><td>spectroscopy software</td></tr><tr><td>Oven</td><td>GmbH Memmert SNB100</td><td>Memmert SNB100</td><td></td></tr><tr><td>Oxalic acid</td><td>VWR BDH Chemicals</td><td></td><td></td></tr><tr><td>P 1000</td><td></td><td></td><td>Soda-processed lignin</td></tr><tr><td>pH paper</td><td></td><td></td><td></td></tr><tr><td>precision balance</td><td></td><td></td><td></td></tr><tr><td>Infrared spectroscopy</td><td></td><td></td><td></td></tr><tr><td>Quatz cuvette</td><td></td><td></td><td></td></tr><tr><td>Rhamnose (98%)</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Rotary vacuum evaporator</td><td>Bucher</td><td></td><td></td></tr><tr><td>Round-bottom flask 500 mL</td><td></td><td></td><td></td></tr><tr><td>sodium borohydride NaBH4</td><td></td><td></td><td></td></tr><tr><td>Schott bottle</td><td></td><td></td><td>glass bottle</td></tr><tr><td>Sovirel tubes</td><td>sovirel</td><td></td><td>Borosilicate glass tubes</td></tr><tr><td>Spatule</td><td></td><td></td><td></td></tr><tr><td>Special tube</td><td></td><td></td><td></td></tr><tr><td>Spectophotometer</td><td>UV-1800 Shimadzu</td><td></td><td></td></tr><tr><td>Sterilization indicator tape</td><td></td><td></td><td></td></tr><tr><td>Stir bar in teflon</td><td></td><td></td><td></td></tr><tr><td>Stirring plate</td><td></td><td></td><td></td></tr><tr><td>Syringes</td><td></td><td></td><td></td></tr><tr><td>Sodium borohydride</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr><tr><td>Titrisol</td><td>Merck</td><td>Merck 109984</td><td>0.1 N H&lt;sub&gt;2&lt;/sub&gt;SO&lt;sub&gt;4&lt;/sub&gt;</td></tr><tr><td>Urea</td><td>VWR BDH Chemicals</td><td></td><td></td></tr><tr><td>Vials</td><td></td><td></td><td></td></tr><tr><td>VolumetriC flask 2.5 L /5 L</td><td>Bucher</td><td></td><td></td></tr><tr><td>Vortex</td><td></td><td></td><td></td></tr><tr><td>Xylose (98%)</td><td>Sigma Aldrich (St. Louis, USA)</td><td></td><td></td></tr></tbody></table>

ultrafast lignin extraction from unusual mediterranean lignocellulosic residues

Pretreatment is still the most expensive step in lignocellulosic biorefinery processes. It must be made cost-effective by minimizing chemical requirements as well as power and heat consumption and by using environment-friendly solvents. Deep eutectic solvents (DESs) are key, green, and low-cost solvents in sustainable biorefineries. They are transparent mixtures characterized by low freezing points resulting from at least one hydrogen bond donor and one hydrogen bond acceptor. Although DESs are promising solvents, it is necessary to combine them with an economic heating technology, such as microwave irradiation, for competitive profitability. Microwave irradiation is a promising strategy to shorten the heating time and boost fractionation because it can rapidly attain the appropriate temperature. The aim of this study was to develop a one-step, rapid method for biomass fractionation and lignin extraction using a low-cost and biodegradable solvent. 
In this study, a microwave-assisted DES pretreatment was conducted for 60 s at 800 W, using three kinds of DESs. The DES mixtures were facilely prepared from choline chloride (ChCl) and three hydrogen-bond donors (HBDs): a monocarboxylic acid (lactic acid), a dicarboxylic acid (oxalic acid), and urea. This pretreatment was used for biomass fractionation and lignin recovery from marine residues (Posidonia leaves and aegagropile), agri-food byproducts (almond shells and olive pomace), forest residues (pinecones), and perennial lignocellulosic grasses (Stipa tenacissima). Further analyses were conducted to determine the yield, purity, and molecular weight distribution of the recovered lignin. In addition, the effect of DESs on the chemical functional groups in the extracted lignin was determined by Fourier-transform infrared (FTIR) spectroscopy. The results indicate that the ChCl-oxalic acid mixture affords the highest lignin purity and the lowest yield. The present study demonstrates that the DES-microwave process is an ultrafast, efficient, and cost-competitive technology for lignocellulosic biomass fractionation.

Figure 2A-C depict the lignin yield of extraction from the six feedstocks, shown in Figure 1A-F, after the combined microwave-DES pretreatment. The results show that the lignin yield obtained with DES1 (ChCl-oxalic acid) (Figure 2A) was lower than the yields obtained with DES2 (ChCl-lactic acid) and DES3 (ChCl-urea) (Figure 2B,C). In addition, lignin yields from pinecones (PC) and olive pomace (OP) were higher at 32.31% and 26.04% for DES1 treatment and 48.72% and 43.76 for DES3, respectively. Lignin yield from alfa leaves (A) was significantly higher than the yields of all other lignins extracted with DES2. Figure 3A-C show that lignin purity exceeded 70% for the three pretreatments of the biomasses, except for DES3 pretreatment of alfa leaves (A), aegagropile (Ag), and almond shells (AS) in the DES3 (ChCl-urea) treatment, which gave a lignin purity of 65%. The highest lignin purity (&#62; 90%) was obtained with the DES1 treatment: alfa leaves (A) 94%, almond shells (AS) 93%, pinecones (PC) 90%, Posidonia leaves (PL) 92%, and olive pomace (OP) 91%.
Lignin purity and yield data were subjected to principal component analysis (PCA) by considering two parameters (yield and purity) and 18 treatments. Figure 4 shows that the correlation circle explained 100% of the total variation. The first component, PCA1, explained 58.09%, and the second component, PCA2, explained 41.91% of the total variation. Lignin purity was positively correlated with DES1 (Ox) treatment. The Pearson correlation coefficients (R) were alfa (A Ox) 0.32, olive pomace (OP Ox) 0.27, pinecones (PC Ox) 0.2, Posidonia leaves (PL Ox) 0.35, almond shells (AS Ox) 0.32, and aegagropile (Ag Ox) 0.05, respectively. However, DES3 treatment was negatively correlated with lignin yield with R-values that oscillated between &#8722;0.37 and &#8722;0.05. Thus, PCA results confirmed that lignin extracted with DES1 was the purest with the lowest yield.
Lignin was characterized for its sugar, nitrogen, and ash contents (Figure 5A-C). The total sugar content was determined by gas chromatography (GC). The carbohydrate content in lignin was extracted using DES3 (ChCl-urea) was the highest (6-15%). This was followed by lignin extracted using DES2 (ChCl-lactic acid), which had a carbohydrate content of 3-12%. However, the lowest carbohydrate content (1%) was reported for lignin extracted using DES1 (ChCl-oxalic acid). The type of sugars identified differed significantly (Figure 6A-C); D-xylose and D-glucose were the most abundant monosaccharides. These results indicate that DES1 was extremely selective in its extraction of lignin compared with the other two DESs, which extracted not only lignin, but also carbohydrates. In other words, lignin purity was lower after extraction with the lactic acid and urea DESs. 
The high selectivity of DES1 to fractionate the lignocellulosic matrix and extract pure lignin is probably because of the high acidity of its hydrogen bonds (alpha = 1.3). Choline chloride contains chloride ions that break the intramolecular interactions of hydrogen bonds, and the carboxylate groups in oxalic acid contribute to dissolve the lignin polymers. Similarly, the nitrogen content of lignin extracted using DES1 was lower than the nitrogen content of lignin extracted using DES2 and DES3, reaching up to 3% (Figure 5A-C). Lignin extracted from alfa leaves had the highest nitrogen content: 2.70, 3.84, and 3.40 for DES1, DES2, and DES3, respectively. These results prove that nitrogenous compounds were extracted and co-precipitated with lignin. Furthermore, lignin calcination in all the samples indicated that lignin extracted using DES2 and DES3 contained a higher inorganic component than lignin extracted using DES1.
These results indicate that DES1 promoted the extraction of lignin with high purity, but with low nitrogen, carbohydrate, and ash contents. In other words, lignin extracted using DES1 (ChCl-oxalic acid) was purer than that extracted using DES2 (ChCl-lactic acid) and DES3 (ChCl-urea), which possesses lower purity and high nitrogen, carbohydrate, and ash contents. Table 1 summarizes the molecular mass distribution of lignin, as analyzed by gel permeation chromatography (GPC) and represented by the number-average molecular weight (Mn), weight-average molecular weight (Mw) and polydispersity index (PDI). The Mw values ranged from 48,123 to 147,233 g mol-1. The lignin extracted by DES2 from alfa leaves, almond shells, and aegagropile had a lower PDI than the lignin extracted by DES1, DES3, and alkali, as well as raw lignin. In contrast, lignin extracted by DES2 from pinecones, olive pomace, and Posidonia leaves showed higher PDI. The lower PDI of the lignin extracted from aegagropile indicates that its molecular weight is more homogenous than that of the lignins extracted from the other biomasses.
The chemical functional groups present in extracted lignin were investigated by FTIR spectroscopy (Figure 7A-F). The strong, broad band between 3,441 and 3,198 cm-1 was attributed to OH stretching vibrations of the alcoholic and phenolic hydroxyl groups involved in hydrogen bonding. The signals in the wavenumber range 2,963-2,852 cm-1 were assigned to alkyl C-H stretching vibrations. Olive pomace, alfa leaves, and almond shells showed more intense bands than the other biomasses. No bands were observed from 2,800 to 1,800 cm-1. The lignin obtained by DES1 and DES2 treatment, had a rising band at 1,708 cm-1, which indicated the presence of unconjugated C=O groups. However, this signal was absent in the solvent spectra (Figure 8B). Lactic and oxalic acid spectra were characterized by a band in the 1,737-1,723 cm-1 range, which indicated the presence of unconjugated C=O groups, whereas the urea spectrum was characterized by two signals in the wavenumber range of 1,660 cm-1 and 1,604 cm-1 attributed to amide groups. The bands at 1,606-1,618 cm-1&#160;were observed in lignin extracted by DES1 and DES2 treatment, associated with ring-conjugated C=C stretch.
The signal at 1,640 cm-1 in lignin extracted by DES3 indicated the presence of C=O stretching vibration in conjugated carbonyl groups of lignin. The signal at 1516 cm-1 arose from the vibrations of the aromatic rings present in lignin, while the band at 1200 cm-1 indicated the presence of ether groups. Bands in the wavenumber range of 1,250-1,200 cm-1 were assigned to C-O stretching of nonaromatic alcohols. The band at 953 cm-1 was assigned to methyl substituents. The results indicate that DES-lignin fractions spectra showed signals at 1,730-1,702 cm-1 and 1,643-1,635 cm-1, assigned to the stretching vibration of unconjugated and conjugated carbonyl groups, respectively. However, these band ranges were absent in three commercial lignins: raw, soda-processed, and alkali-extracted lignins (Figure 8A).This observation indicates that during its extraction and solubilization, some functional groups of lignin were conjugated with oxalic and lactic acid.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61997/61997fig01.jpg" /> 
Figure 1: Mediterranean biomasses studied. (A) Almond shells, (B) Olive pomace, (C) Cone pines, (D) Aegagropile (Posidonia balls), (E)&#160;Posidonia leaves, (F) Alfa leaves. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61997/61997fig02.jpg" /> 
Figure 2: Lignin yield. (A) Choline chloride + Oxalic acid (DES1), (B) Choline chloride + Lactic acid (DES2), (C) Choline chloride + Urea (DES3). Significant differences were determined with one-way ANOVA and Fisher&#39;s post-hoc test (*P &#60; 0.05; **P &#60; 0.01; ***P &#60; 0.001). Abbreviations: A = Alfa leaves, AS = Almond shells, PC = Pinecones, PL = Posidonia leaves, OP = Olive pomace, Ag = Aegagropile; ns = not significant. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61997/61997fig03.jpg" /> 
Figure 3: Lignin (%). (A) Choline chloride + Oxalic acid (DES1), (B) Choline chloride + Lactic acid (DES2), (C) Choline chloride + Urea (DES3). Significant differences were determined with one-way ANOVA and Fisher&#39;s post hoc test (*P &#60; 0.05; **P &#60; 0.01; ***P &#60; 0.001). Abbreviations: A = Alfa leaves, AS = Almond shells, PC = Pinecones, PL = Posidonia leaves, OP = Olive pomace, Ag = Aegagropile; ns = not significant. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61997/61997fig04.jpg" /> 
Figure 4: Principal component analysis of yield and purity of lignin extracted from Mediterranean biomasses. Hydrogen-bond acceptor (HBA) is choline chloride (ChCl) and hydrogen-bond donors (HBD) are Ox = oxalic acid, Lac : lactic acid, and Urea.&#160;PCA = principal component analysis; A = Alfa leaves, AS = Almond shells, PC = Pinecones, PL = Posidonia leaves, OP = Olive pomace, Ag = Aegagropile. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61997/61997fig05.jpg" /> 
Figure 5: Carbohydrate (%), nitrogen (%), and ash content (%) in lignin samples. (A) Choline chloride + Oxalic acid (DES1), (B) Choline chloride + Lactic acid (DES2), (C) Choline chloride + Urea (DES3). Significant differences were determined with one-way ANOVA and Fisher&#39;s post-hoc test (*P &#60; 0.05; **P &#60; 0.01; ***P &#60; 0.001). Abbreviations: A = Alfa leaves, AS = Almond shells, PC = Pinecones, PL = Posidonia leaves, OP = Olive pomace, Ag = Aegagropile; ns = not significant. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61997/61997fig06.jpg" /> 
Figure 6: Identification of monosaccharides in lignin samples (%).&#160;(A) Choline chloride + Oxalic acid (DES1), (B) Choline chloride + Lactic acid (DES2), (C) Choline chloride + Urea (DES3). Significant differences were determined with one-way ANOVA and Fisher&#39;s post-hoc test (*P &#60; 0.05; **P &#60; 0.01; ***P &#60; 0.001). Abbreviations: A = Alfa leaves, AS = Almond shells, PC = Pinecones, PL = Posidonia leaves, OP = Olive pomace, Ag = Aegagropile; ns = not significant. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61997/61997fig07.jpg" /> 
Figure 7: Fourier-transform infrared spectra of lignin samples. (A) Alfa leaves, (B) Almond shells, (C) Pinecones, (D) Posidonia leaves, (E) Olive pomace, (F) Aegagropile. Abbreviations: DES1 = Choline chloride + Oxalic acid, DES2 = Choline chloride + Lactic acid, DES3 = Choline chloride + Urea. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/61997/61997fig08.jpg" /> 
Figure 8: Fourier-transform infrared spectra. (A)&#160;Lignin controls, (B) hydrogen bond donors. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/61997fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample</td>
			<td>Treatment</td>
			<td>Mn</td>
			<td>Mw</td>
			<td>PDI</td>
		</tr>
		<tr>
			<td rowspan="3">A</td>
			<td>Urea</td>
			<td>47558</td>
			<td>120141</td>
			<td>2.5</td>
		</tr>
		<tr>
			<td>Lac</td>
			<td>35241</td>
			<td>73665</td>
			<td>2.1</td>
		</tr>
		<tr>
			<td>Ox</td>
			<td>35793</td>
			<td>84312</td>
			<td>2.4</td>
		</tr>
		<tr>
			<td rowspan="3">AS</td>
			<td>Urea</td>
			<td>50181</td>
			<td>105817</td>
			<td>2.1</td>
		</tr>
		<tr>
			<td>Lac</td>
			<td>60409</td>
			<td>104915</td>
			<td>1.7</td>
		</tr>
		<tr>
			<td>Ox</td>
			<td>83112</td>
			<td>147233</td>
			<td>1.8</td>
		</tr>
		<tr>
			<td rowspan="3">PC</td>
			<td>Urea</td>
			<td>34013</td>
			<td>65181</td>
			<td>1.9</td>
		</tr>
		<tr>
			<td>Lac</td>
			<td>55513</td>
			<td>145963</td>
			<td>2.6</td>
		</tr>
		<tr>
			<td>Ox</td>
			<td>46409</td>
			<td>102298</td>
			<td>2.2</td>
		</tr>
		<tr>
			<td rowspan="3">PL</td>
			<td>Urea</td>
			<td>25696</td>
			<td>50093</td>
			<td>1.9</td>
		</tr>
		<tr>
			<td>Lac</td>
			<td>45530</td>
			<td>122900</td>
			<td>2.7</td>
		</tr>
		<tr>
			<td>Ox</td>
			<td>28427</td>
			<td>70726</td>
			<td>2.5</td>
		</tr>
		<tr>
			<td rowspan="3">OP</td>
			<td>Urea</td>
			<td>29669</td>
			<td>70424</td>
			<td>2.4</td>
		</tr>
		<tr>
			<td>Lac</td>
			<td>26735</td>
			<td>66743</td>
			<td>2.5</td>
		</tr>
		<tr>
			<td>Ox</td>
			<td>34161</td>
			<td>75509</td>
			<td>2.2</td>
		</tr>
		<tr>
			<td rowspan="3">Ag</td>
			<td>Urea</td>
			<td>30184</td>
			<td>48123</td>
			<td>1.6</td>
		</tr>
		<tr>
			<td>Lac</td>
			<td>33835</td>
			<td>52123</td>
			<td>1.5</td>
		</tr>
		<tr>
			<td>Ox</td>
			<td>30025</td>
			<td>49808</td>
			<td>1.7</td>
		</tr>
		<tr>
			<td rowspan="2">Control</td>
			<td>Raw lignin</td>
			<td>23275.3</td>
			<td>36496.5</td>
			<td>1.6</td>
		</tr>
		<tr>
			<td>Alkali-extracted lignin</td>
			<td>22792.6</td>
			<td>43014.3</td>
			<td>1.9</td>
		</tr>
	</tbody>
</table>
Table 1: Molecular weights of the lignins. Abbreviations: A = Alfa leaves, AS = Almond shells, PC = Pinecones, PL = Posidonia leaves, OP = Olive pomace, Ag = Aegagropile; Mn = number-average molecular weight; Mw = weight-averaged molecular weight; PDI = polydispersity index; Ox =oxalic acid; Lac = lactic acid.
Figure S1: Lignin.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/Figure S1.pdf" target="_blank">Please click here to download this File.</a>
Figure S2: Samples after being autoclaved (30 mg of lignin + 1 mL of 72% sulfuric acid + 28 mL of distilled water).&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/Figure S2.pdf" target="_blank">Please click here to download this File.</a>
Figure S3: Lignin pellets.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/Figure S3.pdf" target="_blank">Please click here to download this File.</a>
Figure S4: Solid residue washed four times to recover maximum lignin content.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/Figure S4.pdf" target="_blank">Please click here to download this File.</a>
Figure S5: Gel permeation chromatograms of lignin controls, raw and alkali-extracted lignins.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/Figure S5.pdf" target="_blank">Please click here to download this File.</a>
Figure S6: Gel permeation chromatograms of lignin samples. Abbreviations: A = Alfa leaves, AS = Almond shells, PC = Pinecones, PL = Posidonia leaves, OP = Olive pomace, Ag = Aegagropile; DES1 = Choline chloride + Oxalic acid, DES2 = Choline chloride + Lactic acid, DES3 = Choline chloride + Urea.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/Figure S6.pdf" target="_blank">Please click here to download this File.</a>
Figure S7: Flowsheet of the deep eutectic solvent (DES)-microwave process for lignin extraction.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61997/Figure S7.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Ultrafast Lignin Extraction from Unusual Mediterranean Lignocellulosic Residues at JoVE.com

Ultrafast Lignin Extraction from Unusual Mediterranean Lignocellulosic Residues

Deep eutectic solvent-based, microwave-assisted pretreatment is a green, fast, and efficient process for lignocellulosic fractionation and high-purity lignin recovery.

Pretreatment is still the most expensive step in lignocellulosic biorefinery processes. It must be made cost-effective by minimizing chemical requirements ...

ultrafast-lignin-extraction-from-unusual-mediterranean

Nanyang Technological University

Research

JoVE Journal

Chemistry

6.5K Views.  University of Liege (Gembloux Agro-Bio Tech Campus). Het doel van ons protocol is om de biomassa te fractioneren en lignine in één stap te extraheren. Met behulp van deze methode kan lignine worden teruggewonnen door eenvoudige filtratie na de behandeling zonder aanpassing van de pH, maar eenvoudig door toevoeging van gedestilleerd water. De focus van deze studie is om het effect van deze gecombineerde behandeling van grondstoffractie, de invloed ervan op de zuiverheid en opbrengst van lignine en het effect ervan op de moleculaire gewichten e...

Video: Ultrafast Lignin Extraction from Unusual Mediterranean Lignocellulosic Residues

Methods for Facilitating Microbial Growth on Pulp Mill Waste Streams and Characterization of the Biodegradation Potential of Cultured Microbes

The kraft process is applied to wood chips for separation of lignin from the polysaccharides within lignocellulose for pulp that will produce a high quality paper. Black liquor is a pulping waste generated by the kraft process that has potential for downstream bioconversion. However, the recalcitrant nature of the lignocellulose resources, its chemical derivatives that constitute the majority of available organic carbon within black liquor, and its basic pH present challenges to microbial biodegradation of this waste material. Methods for the collection and modification of black liquor for microbial growth are aimed at utilization of this pulp waste to convert the lignin, organic acids, and polysaccharide degradation byproducts into valuable chemicals. The lignocellulose extraction techniques presented provide a reproducible method for preparation of lignocellulose growth substrates for understanding metabolic capacities of cultured microorganisms. Use of gas chromatography-mass spectrometry enables the identification and quantification of the fermentation products resulting from the growth of microorganisms on pulping waste. These methods when used together can facilitate the determination of the metabolic activity of microorganisms with potential to produce fermentation products that would provide greater value to the pulping system and reduce effluent waste, thereby increasing potential paper milling profits and offering additional uses for black liquor.

Industrial wastes can be collected and modified to analyze microbial growth. Lignocellulose extraction techniques provide components to analyze specific biodegradation ability. Gas chromatography-mass spectrometry identifies fermentation products of microorganisms grown on pulping waste. These methods determine the metabolic capacity of microorganisms to degrade pulping waste.

The kraft process is applied to wood chips for separation of lignin from the polysaccharides within lignocellulose for pulp that will produce a high ...

Environment

Towards Biomimicking Wood: Fabricated Free-standing Films of Nanocellulose, Lignin, and a Synthetic Polycation

Woody materials are comprised of plant cell walls that contain a layered secondary cell wall composed of structural polymers of polysaccharides and lignin. Layer-by-layer (LbL) assembly process which relies on the assembly of oppositely charged molecules from aqueous solutions was used to build a freestanding composite film of isolated wood polymers of lignin and oxidized nanofibril cellulose (NFC). To facilitate the assembly of these negatively charged polymers, a positively charged polyelectrolyte, poly(diallyldimethylammomium chloride) (PDDA), was used as a linking layer to create this simplified model cell wall. The layered adsorption process was studied quantitatively using quartz crystal microbalance with dissipation monitoring (QCM-D) and ellipsometry. The results showed that layer mass/thickness per adsorbed layer increased as a function of total number of layers. The surface coverage of the adsorbed layers was studied with atomic force microscopy (AFM). Complete coverage of the surface with lignin in all the deposition cycles was found for the system, however, surface coverage by NFC increased with the number of layers. The adsorption process was carried out for 250 cycles (500 bilayers) on a cellulose acetate (CA) substrate. Transparent free-standing LBL assembled nanocomposite films were obtained when the CA substrate was later dissolved in acetone. Scanning electron microscopy (SEM) of the fractured cross-sections showed a lamellar structure, and the thickness per adsorption cycle (PDDA-Lignin-PDDA-NC) was estimated to be 17 nm for two different lignin types used in the study. The data indicates a film with highly controlled architecture where nanocellulose and lignin are spatially deposited on the nanoscale (a polymer-polymer nanocomposites), similar to what is observed in the native cell wall.

The objective of this research was to form synthetic plant cell wall tissue using layer-by-layer assembly of nanocellulose fibrils and isolated lignin assembled from dilute aqueous suspensions.&#160; Surface measurement techniques of quartz crystal microbalance and atomic force microscopy were used to monitor the formation of the polymer-polymer nanocomposite material.

Woody materials are comprised of plant cell walls that contain a layered secondary cell wall composed of structural polymers of polysaccharides and ...

Preparing Silica Aerogel Monoliths via a Rapid Supercritical Extraction Method

A procedure for the fabrication of monolithic silica aerogels in eight hours or less via a rapid supercritical extraction process is described. The procedure requires 15-20 min of preparation time, during which a liquid precursor mixture is prepared and poured into wells of a metal mold that is placed between the platens of a hydraulic hot press, followed by several hours of processing within the hot press. The precursor solution consists of a 1.0:12.0:3.6:3.5 x 10-3 molar ratio of tetramethylorthosilicate (TMOS):methanol:water:ammonia. In each well of the mold, a porous silica sol-gel matrix forms. As the temperature of the mold and its contents is increased, the pressure within the mold rises. After the temperature/pressure conditions surpass the supercritical point for the solvent within the pores of the matrix (in this case, a methanol/water mixture), the supercritical fluid is released, and monolithic aerogel remains within the wells of the mold. With the mold used in this procedure, cylindrical monoliths of 2.2 cm diameter and 1.9 cm height are produced. Aerogels formed by this rapid method have comparable properties (low bulk and skeletal density, high surface area, mesoporous morphology) to those prepared by other methods that involve either additional reaction steps or solvent extractions (lengthier processes that generate more chemical waste).The rapid supercritical extraction method can also be applied to the fabrication of aerogels based on other precursor recipes.

This article describes a rapid supercritical extraction method for fabricating silica aerogels. By utilizing a confined mold and hydraulic hot press, monolithic aerogels can be made in eight hours or less.

A procedure for the fabrication of monolithic silica aerogels in eight hours or less via a rapid supercritical extraction process is described. The ...

High-throughput Screening of Recalcitrance Variations in Lignocellulosic Biomass: Total Lignin, Lignin Monomers, and Enzymatic Sugar Release

The conversion of lignocellulosic biomass to fuels, chemicals, and other commodities has been explored as one possible pathway toward reductions in the use of non-renewable energy sources. In order to identify which plants, out of a diverse pool, have the desired chemical traits for downstream applications, attributes, such as cellulose and lignin content, or monomeric sugar release following an enzymatic saccharification, must be compared. The experimental and data analysis protocols of the standard methods of analysis can be time-consuming, thereby limiting the number of samples that can be measured. High-throughput (HTP) methods alleviate the shortcomings of the standard methods, and permit the rapid screening of available samples to isolate those possessing the desired traits. This study illustrates the HTP sugar release and pyrolysis-molecular beam mass spectrometry pipelines employed at the National Renewable Energy Lab. These pipelines have enabled the efficient assessment of thousands of plants while decreasing experimental time and costs through reductions in labor and consumables.

Plant cell wall structure and chemistry traits are evaluated to identify ideal feedstocks for biofuels and bio-materials. Standard methods have limitations when applied to large data sets. These high-throughput pretreatment, enzyme saccharification, and pyrolysis-molecular beam mass spectrometry methods compare large numbers of biomass samples with decreased experimental time and cost.

The conversion of lignocellulosic biomass to fuels, chemicals, and other commodities has been explored as one possible pathway toward reductions in the ...

Pretreatment of Lignocellulosic Biomass with Low-cost Ionic Liquids

A number of ionic liquids (ILs) with economically attractive production costs have recently received growing interest as media for the delignification of a variety of lignocellulosic feedstocks. Here we demonstrate the use of these low-cost protic ILs in the deconstruction of lignocellulosic biomass (Ionosolv pretreatment), yielding cellulose and a purified lignin. In the most generic process, the protic ionic liquid is synthesized by accurate combination of aqueous acid and amine base. The water content is adjusted subsequently. For the delignification, the biomass is placed into a vessel with IL solution at elevated temperatures to dissolve the lignin and hemicellulose, leaving a cellulose-rich pulp ready for saccharification (hydrolysis to fermentable sugars). The lignin is later precipitated from the IL by the addition of water and recovered as a solid. The removal of the added water regenerates the ionic liquid, which can be reused multiple times. This protocol is useful to investigate the significant potential of protic ILs for use in commercial biomass pretreatment/lignin fractionation for producing biofuels or renewable chemicals and materials.

The pretreatment of lignocellulosic biomass with protic low-cost ionic liquids is shown, resulting in a delignified cellulose-rich pulp and a purified lignin. The pulp gives rise to high glucose yields after enzymatic saccharification.

A number of ionic liquids (ILs) with economically attractive production costs have recently received growing interest as media for the delignification of ...

Lignin Down-regulation of Zea mays via dsRNAi and Klason Lignin Analysis

To facilitate the use of lignocellulosic biomass as an alternative bioenergy resource, during biological conversion processes, a pretreatment step is needed to open up the structure of the plant cell wall, increasing the accessibility of the cell wall carbohydrates. Lignin, a polyphenolic material present in many cell wall types, is known to be a significant hindrance to enzyme access. Reduction in lignin content to a level that does not interfere with the structural integrity and defense system of the plant might be a valuable step to reduce the costs of bioethanol production. In this study, we have genetically down-regulated one of the lignin biosynthesis-related genes, cinnamoyl-CoA reductase (ZmCCR1) via a double stranded RNA interference technique. The ZmCCR1_RNAi construct was integrated into the maize genome using the particle bombardment method. Transgenic maize plants grew normally as compared to the wild-type control plants without interfering with biomass growth or defense mechanisms, with the exception of displaying of brown-coloration in transgenic plants leaf mid-ribs, husks, and stems. The microscopic analyses, in conjunction with the histological assay, revealed that the leaf sclerenchyma fibers were thinned but the structure and size of other major vascular system components was not altered. The lignin content in the transgenic maize was reduced by 7-8.7%, the crystalline cellulose content was increased in response to lignin reduction, and hemicelluloses remained unchanged. The analyses may indicate that carbon flow might have been shifted from lignin biosynthesis to cellulose biosynthesis. This article delineates the procedures used to down-regulate the lignin content in maize via RNAi technology, and the cell wall compositional analyses used to verify the effect of the modifications on the cell wall structure.

Lignin Down-regulation of Zea mays via dsRNAi and Klason Lignin Analysis

A double stranded RNA interference (dsRNAi) technique is employed to down-regulate the maize cinnamoyl coenzyme A reductase (ZmCCR1) gene to lower plant lignin content. Lignin down-regulation from the cell wall is visualized by microscopic analyses and quantified by the Klason method. Compositional changes in hemicellulose and crystalline cellulose are analyzed.

To facilitate the use of lignocellulosic biomass as an alternative bioenergy resource, during biological conversion processes, a pretreatment step is ...

Bioengineering

Extraction of Lignin with High &#946;-O-4 Content by Mild Ethanol Extraction and Its Effect on the Depolymerization Yield

Lignin valorization strategies are a key factor for achieving more economically competitive biorefineries based on lignocellulosic biomass. Most of the emerging elegant procedures to obtain specific aromatic products rely on the lignin substrate having a high content of the readily cleavable &#946;-O-4 linkage as present in the native lignin structure. This provides a miss-match with typical technical lignins that are highly degraded and therefore are low in &#946;-O-4 linkages. Therefore, the extraction yields, and the quality of the obtained lignin are of utmost importance to access new lignin valorization pathways. In this manuscript, a simple protocol is presented to obtain lignins with high &#946;-O-4 content by relatively mild ethanol extraction that can be applied to different lignocellulose sources. Furthermore, analysis procedures to determine the quality of the lignins are presented together with a depolymerization protocol that yields specific phenolic 2-arylmethyl-1,3-dioxolanes, which can be used to evaluate the obtained lignins. The presented results demonstrate the link between lignin quality and potential for the lignins to be depolymerized into specific monomeric aromatic chemicals. Overall, the extraction and depolymerization demonstrates a trade-off between the lignin extraction yield and the retention of the native aryl-ether structure and thus the potential of the lignin to be used as the substrate for the production of chemicals for higher-value applications.

Here, we present a protocol to perform ethanol extraction of lignin from several biomass sources. The effect of the extraction conditions on the lignin yield and &#946;-O-4 content are presented. Selective depolymerization is performed on the obtained lignins to obtain high aromatic monomer products.

Lignin valorization strategies are a key factor for achieving more economically competitive biorefineries based on lignocellulosic biomass. Most of the ...

Comprehensive Compositional Analysis of Plant Cell Walls (Lignocellulosic biomass) Part I: Lignin

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st century. This has rekindled interest in the use of plant products as industrial raw materials for the production of liquid fuels for transportation1 and other products such as biocomposite materials7. Plant biomass remains one of the greatest untapped reserves on the planet4. It is mostly comprised of cell walls that are composed of energy rich polymers including cellulose, various hemicelluloses (matrix polysaccharides, and the polyphenol lignin6 and thus sometimes termed lignocellulosics. However, plant cell walls have evolved to be recalcitrant to degradation as walls provide tensile strength to cells and the entire plants, ward off pathogens, and allow water to be transported throughout the plant; in the case of trees up to more the 100 m above ground level. Due to the various functions of walls, there is an immense structural diversity within the walls of different plant species and cell types within a single plant4. Hence, depending of what crop species, crop variety, or plant tissue is used for a biorefinery, the processing steps for depolymerization by chemical/enzymatic processes and subsequent fermentation of the various sugars to liquid biofuels need to be adjusted and optimized. This fact underpins the need for a thorough characterization of plant biomass feedstocks. Here we describe a comprehensive analytical methodology that enables the determination of the composition of lignocellulosics and is amenable to a medium to high-throughput analysis. In this first part we focus on the analysis of the polyphenol lignin (Figure 1). The method starts of with preparing destarched cell wall material. The resulting lignocellulosics are then split up to determine its lignin content by acetylbromide solubilization3, and its lignin composition in terms of its syringyl, guaiacyl- and p-hydroxyphenyl units5. The protocol for analyzing the carbohydrates in lignocellulosic biomass including cellulose content and matrix polysaccharide composition is discussed in Part II2.

Comprehensive Compositional Analysis of Plant Cell Walls Lignocellulosic biomass Part I: Lignin

Plant biomass is a major carbon-neutral renewable resource that could be used for the production of biofuels. Plant biomass consists mainly of cell walls, a structurally complex composite material termed lignocellulosics. Here we describe a protocol for a comprehensive analysis of the content and composition of the polyphenolic lignin.

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st ...

Biology

Fractionation of Lignocellulosic Biomass using the OrganoCat Process

The shift from a petroleum-based to a more sustainable and bio-based economy requires the development of new refinery concepts to maintain the supply of raw materials and energy. For these novel and sustainable biorefinery concepts, it is important to use catalysts and solvents that are aligned with the principles of Green Chemistry. Therefore, the implementation of biogenic alternatives can be a promising solution. The lignocellulose pretreatment and fractionation process presented herein-OrganoCat-is an integrated fractionation of lignocellulose into its main components using biogenic acids such as 2,5-furandicarboxylic acid as catalyst. Hemicelluloses and other non-cellulosic polysaccharides are selectively depolymerized by the diluted acid and dissolved, while the crystalline cellulose remains in the solid pulp. In the presence of a second organic phase consisting of biogenic 2-methyltetrahydrofuran, disentangled lignin is extracted in situ. The process allows for the efficient fractionation of the three main components-lignin, cellulose, and non-cellulosic sugars. This helps to focus on the quality of the lignin, the improvement of enzymatic hydrolysis of the cellulose-enriched pulp, and the mild non-cellulosic sugar extraction with low degradation.

OrganoCat is a method for the pretreatment and fractionation of lignocellulose under mild conditions into lignin, fermentable sugars, and cellulose pulp. In a biogenic, biphasic solvent system of water and 2-methyltetrahydrofuran with 2,5-furancarboxylic acid as catalyst, the OrganoCat products are separated in situ for straightforward product recovery.

The shift from a petroleum-based to a more sustainable and bio-based economy requires the development of new refinery concepts to maintain the supply of ...

Label-free in situ Imaging of Lignification in Plant Cell Walls

Meeting growing energy demands safely and efficiently is a pressing global challenge. Therefore, research into biofuels production that seeks to find cost-effective and sustainable solutions has become a topical and critical task. Lignocellulosic biomass is poised to become the primary source of biomass for the conversion to liquid biofuels1-6. However, the recalcitrance of these plant cell wall materials to cost-effective and efficient degradation presents a major impediment for their use in the production of biofuels and chemicals4. In particular, lignin, a complex and irregular poly-phenylpropanoid heteropolymer, becomes problematic to the postharvest deconstruction of lignocellulosic biomass. For example in biomass conversion for biofuels, it inhibits saccharification in processes aimed at producing simple sugars for fermentation7. The effective use of plant biomass for industrial purposes is in fact largely dependent on the extent to which the plant cell wall is lignified. The removal of lignin is a costly and limiting factor8 and lignin has therefore become a key plant breeding and genetic engineering target in order to improve cell wall conversion.
Analytical tools that permit the accurate rapid characterization of lignification of plant cell walls become increasingly important for evaluating a large number of breeding populations. Extractive procedures for the isolation of native components such as lignin are inevitably destructive, bringing about significant chemical and structural modifications9-11. Analytical chemical in situ methods are thus invaluable tools for the compositional and structural characterization of lignocellulosic materials. Raman microscopy is a technique that relies on inelastic or Raman scattering of monochromatic light, like that from a laser, where the shift in energy of the laser photons is related to molecular vibrations and presents an intrinsic label-free molecular "fingerprint" of the sample. Raman microscopy can afford non-destructive and comparatively inexpensive measurements with minimal sample preparation, giving insights into chemical composition and molecular structure in a close to native state. Chemical imaging by confocal Raman microscopy has been previously used for the visualization of the spatial distribution of cellulose and lignin in wood cell walls12-14. Based on these earlier results, we have recently adopted this method to compare lignification in wild type and lignin-deficient transgenic Populus trichocarpa (black cottonwood) stem wood15. Analyzing the lignin Raman bands16,17 in the spectral region between 1,600 and 1,700 cm-1, lignin signal intensity and localization were mapped in situ. Our approach visualized differences in lignin content, localization, and chemical composition. Most recently, we demonstrated Raman imaging of cell wall polymers in Arabidopsis thaliana with lateral resolution that is sub-&mu;m18. Here, this method is presented affording visualization of lignin in plant cell walls and comparison of lignification in different tissues, samples or species without staining or labeling of the tissues.

Label-free in situ Imaging of Lignification in Plant Cell Walls

A method based on confocal Raman microscopy is presented that affords label-free visualization of lignin in plant cell walls and comparison of lignification in different tissues, samples or species.

Meeting growing energy demands safely and efficiently is a pressing global challenge. Therefore, research into biofuels production that seeks to find ...

Ultrafast Lignin Extraction from Unusual Mediterranean Lignocellulosic Residues

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