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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J., 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=Facile+fractionation+of+lignocelluloses+by+biomass-derived+deep+eutectic+solvent+(DES)+pretreatment+for+cellulose+enzymatic+hydrolysis+and+lignin+valorization.">Facile fractionation of lignocelluloses by biomass-derived deep eutectic solvent (DES) pretreatment for cellulose enzymatic hydrolysis and lignin valorization.</a> Green Chemistry. 21, 275-283 (2019).</li><li>Alvarez-Vasco, C., 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=Unique+low-molecular-weight+lignin+with+high+purity+extracted+from+wood+by+deep+eutectic+solvents+(DES):+a+source+of+lignin+for+valorization.">Unique low-molecular-weight lignin with high purity extracted from wood by deep eutectic solvents (DES): a source of lignin for valorization.</a> Green Chemistry. 18, 5133-5141 (2016).</li><li>Banu, J. R., 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=A+review+on+biopolymer+production+via+lignin+valorization.">A review on biopolymer production via lignin valorization.</a> Bioresource Technologie. 290, 121790 (2019).</li><li>Gordobil, O., Olaizola, P., Banales, J. M., Labidi, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lignins+from+agroindustrial+by-products+as+natural+ingredients+for+cosmetics+chemical+structure+and+in+vitro+sunscreen+and+cytotoxic+activities.">Lignins from agroindustrial by-products as natural ingredients for cosmetics chemical structure and in vitro sunscreen and cytotoxic activities.</a> Molecules. 25 (5), 1131 (2020).</li><li>Lee, C. S., Thu Tran, T. 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...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#176;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 K2SO4 + 0.045 g CuSO4.5 H2O + 0.045 g TiO2)</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 &#215; 250 mm, 5 &#181;m) preceded by a guard column (TSK gel alpha guard column 4.6 mm &#215; 50 mm, 5 &#181;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&#176;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 
			equipped with an attenuated total reflectance (ATR) module. 
			Spectra were recorded in the 4,000&#8211;400 cm&#8722;1 range with 32 scans 
			at a resolution of 4.0 cm&#8722;1</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 H2SO4</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</...

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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 ...

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6.3K Views.  University of Liege (Gembloux Agro-Bio Tech Campus). Deep eutectic solvent-based, microwave-assisted pretreatment is a green, fast, and efficient process for lignocellulosic fractionation and high-purity lignin recovery. 

Video: Ultrafast Lignin Extraction from Unusual Mediterranean Lignocellulosic Residues

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 ...

Optimization of Synthetic Proteins: Identification of Interpositional Dependencies Indicating Structurally and/or Functionally Linked Residues

Protein alignments are commonly used to evaluate the similarity of protein residues, and the derived consensus sequence used for identifying functional units (e.g., domains). Traditional consensus-building models fail to account for interpositional dependencies &#8211; functionally required covariation of residues that tend to appear simultaneously throughout evolution and across the phylogentic tree. These relationships can reveal important clues about the processes of protein folding, thermostability, and the formation of functional sites, which in turn can be used to inform the engineering of synthetic proteins. Unfortunately, these relationships essentially form sub-motifs which cannot be predicted by simple &#8220;majority rule&#8221; or even HMM-based consensus models, and the result can be a biologically invalid &#8220;consensus&#8221; which is not only never seen in nature but is less viable than any extant protein. We have developed a visual analytics tool, StickWRLD, which creates an interactive 3D representation of a protein alignment and clearly displays covarying residues. The user has the ability to pan and zoom, as well as dynamically change the statistical threshold underlying the identification of covariants. StickWRLD has previously been successfully used to identify functionally-required covarying residues in proteins such as Adenylate Kinase and in DNA sequences such as endonuclease target sites.

Synthetic protein sequences based on consensus motifs typically ignore co-evolving residues, that imply interpositional dependencies (IPDs). IPDs can be essential to activity, and designs that disregard them may result in suboptimal results. This protocol uses StickWRLD to identify IPDs and help inform rational protein design, resulting in more efficient results.

Protein alignments are commonly used to evaluate the similarity of protein residues, and the derived consensus sequence used for identifying functional ...

Direct Imaging of Laser-driven Ultrafast Molecular Rotation

We present a method for visualizing laser-induced, ultrafast molecular rotational wave packet dynamics. We have developed a new 2-dimensional Coulomb explosion imaging setup in which a hitherto-impractical camera angle is realized. In our imaging technique, diatomic molecules are irradiated with a circularly polarized strong laser pulse. The ejected atomic ions are accelerated perpendicularly to the laser propagation. The ions lying in the laser polarization plane are selected through the use of a mechanical slit and imaged with a high-throughput, 2-dimensional detector installed parallel to the polarization plane. Because a circularly polarized (isotropic) Coulomb exploding pulse is used, the observed angular distribution of the ejected ions directly corresponds to the squared rotational wave function at the time of the pulse irradiation. To create a real-time movie of molecular rotation, the present imaging technique is combined with a femtosecond pump-probe optical setup in which the pump pulses create unidirectionally rotating molecular ensembles. Due to the high image throughput of our detection system, the pump-probe experimental condition can be easily optimized by monitoring a real-time snapshot. As a result, the quality of the observed movie is sufficiently high for visualizing the detailed wave nature of motion. We also note that the present technique can be implemented in existing standard ion imaging setups, offering a new camera angle or viewpoint for the molecular systems without the need for extensive modification.

We present a protocol for creating a real-time movie of a molecular rotational wave packet using a high-resolution Coulomb explosion imaging setup.

We present a method for visualizing laser-induced, ultrafast molecular rotational wave packet dynamics. We have developed a new 2-dimensional Coulomb ...

Separation of Aldehydes and Reactive Ketones from Mixtures Using a Bisulfite Extraction Protocol

The purification of organic compounds is an essential component of routine synthetic operations. The ability to remove contaminants into an aqueous layer by generating a charged structure provides an opportunity to use extraction as a simple purification technique. By combining the use of a miscible organic solvent with saturated sodium bisulfite, aldehydes and reactive ketones can be successfully transformed into charged bisulfite adducts that can then be separated from other organic components of a mixture by the introduction of an immiscible organic layer. Here, we describe a simple protocol for the removal of aldehydes, including sterically-hindered neopentyl aldehydes and some ketones, from chemical mixtures. Ketones can be separated if they are sterically unhindered cyclic or methyl ketones. For aliphatic aldehydes and ketones, dimethylformamide is used as the miscible solvent to improve removal rates. The bisulfite addition reaction can be reversed by basification of the aqueous layer, allowing for the re-isolation of the reactive carbonyl component of a mixture.

Here, we present a protocol to remove aldehydes and reactive ketones from mixtures by a liquid-liquid extraction protocol directly with saturated sodium bisulfite in a miscible solvent. This combined protocol is rapid and facile to perform. The aldehyde or ketone can be re-isolated by the basification of the aqueous layer.

The purification of organic compounds is an essential component of routine synthetic operations. The ability to remove contaminants into an aqueous layer ...

Measurement of Ultrafast Vibrational Coherences in Polyatomic Radical Cations with Strong-Field Adiabatic Ionization

We present a pump-probe method for preparing vibrational coherences in polyatomic radical cations and probing their ultrafast dynamics. By shifting the wavelength of the strong-field ionizing pump pulse from the commonly used 800 nm into the near-infrared (1200-1600 nm), the contribution of adiabatic electron tunneling to the ionization process increases relative to multiphoton absorption. Adiabatic ionization results in predominant population of the ground electronic state of the ion upon electron removal, which effectively prepares a coherent vibrational state (&#34;wave packet&#34;) amenable to subsequent excitation. In our experiments, the coherent vibrational dynamics are probed with a weak-field 800 nm pulse and the time-dependent yields of dissociation products measured in a time-of-flight mass spectrometer. We present the measurements on the molecule dimethyl methylphosphonate (DMMP) to illustrate how using 1500 nm pulses for excitation enhances the amplitude of coherent oscillations in ion yields by a factor of 10 as compared to 800 nm pulses. This protocol may be implemented in existing pump-probe setups through the incorporation of an optical parametric amplifier (OPA) for wavelength conversion.

We present a protocol for probing ultrafast vibrational coherences in polyatomic radical cations that result in molecular dissociation.

We present a pump-probe method for preparing vibrational coherences in polyatomic radical cations and probing their ultrafast dynamics. By shifting the ...

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.

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

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 ...

Ultrafast Time-resolved Near-IR Stimulated Raman Measurements of Functional &#960;-conjugate Systems

Femtosecond time-resolved stimulated Raman spectroscopy is a promising method of observing the structural dynamics of short-lived transients with near infrared (near-IR) transitions, because it can overcome the low sensitivity of spontaneous Raman spectrometers in the near-IR region. Here, we describe technical details of a femtosecond time-resolved near-IR multiplex stimulated Raman spectrometer that we have recently developed. A description of signal generation and optimization, measurement, data acquisition, and calibration and correction of recorded data is provided as well. We present an application of our spectrometer to analyze the excited-state dynamics of &#946;-carotene in toluene solution. A C=C stretch band of &#946;-carotene in the second lowest excited singlet (S2) state and the lowest excited singlet (S1) state is clearly observed in the recorded time-resolved stimulated Raman spectra. The femtosecond time-resolved near-IR stimulated Raman spectrometer is applicable to the structural dynamics of &#960;-conjugate systems from simple molecules to complex materials.

Details of signal generation and optimization, measurement, data acquisition, and data handling for a femtosecond time-resolved near-IR stimulated Raman spectrometer are described. A near infrared stimulated Raman study on the excited-state dynamics of &#946;-carotene in toluene is shown as a representative application.

Femtosecond time-resolved stimulated Raman spectroscopy is a promising method of observing the structural dynamics of short-lived transients with near ...

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 ...

Designed for Molecular Recycling: A Lignin-Derived Semi-aromatic Biobased Polymer

The development of chemically recyclable biopolymers offers opportunities within the pursuit of a circular economy. Chemically recyclable biopolymers make a positive effort to solve the issue of polymer materials in the disposal phase after the use phase. In this paper, the production of biobased semi-aromatic polyesters, which can be extracted entirely from biomass such as lignin, is described and visualized. The polymer poly-S described in this paper has thermal properties similar to certain commonly used plastics, such as PET. We developed a Green Knoevenagel reaction, which can efficiently produce monomers from aromatic aldehydes and malonic acid. This reaction has been proven to be scalable and has a remarkably low calculated E-factor. These polyesters with ligno-phytochemicals as a starting point show an efficient molecular recycling with minimal losses. The polyester poly(dihydrosinapinic acid) (poly-S) is presented as an example of these semi-aromatic polyesters, and the polymerization, depolymerization, and re-polymerization are described.

An example of a closed-loop approach towards a circular materials economy is described here. A whole sustainable cycle is presented where biobased semi-aromatic polyesters are designed by polymerization, depolymerization, and then re-polymerized with only slight changes in their yields or final properties.

The development of chemically recyclable biopolymers offers opportunities within the pursuit of a circular economy. Chemically recyclable biopolymers make ...

Extraction of Non-Protein Amino Acids from Cyanobacteria for Liquid Chromatography-Tandem Mass Spectrometry Analysis

Non-protein amino acids (NPAAs) are a large class of amino acids (AAs) that are not genetically encoded for translation into proteins. The analysis of NPAAs can provide crucial information about cellular uptake and/or function, metabolic pathways, and potential toxicity. &#946;-methylamino-L-alanine (BMAA) is a neurotoxic NPAA produced by various algae species and is associated with an increased risk for neurodegenerative diseases, which has led to significant research interest. There are numerous ways to extract AAs for analysis, with liquid chromatography-tandem mass spectrometry being the most common, requiring protein precipitation followed by acid hydrolysis of the protein pellet. Studies on the presence of BMAA in algal species provide contradictory results, with the use of unvalidated sample preparation/extraction and analysis a primary cause. Like most NPAAs, protein precipitation in 10% aqueous TCA and hydrolysis with fuming HCl is the most appropriate form of extraction for BMAA and its isomers aminoethylglycine (AEG) and 2,4-diaminobutyric acid (2,4-DAB). The present protocol describes the steps in a validated NPAA extraction method commonly used in research and teaching laboratories.

The present protocol describes the extraction of non-protein amino acids from biological matrices via trichloroacetic acid (TCA) protein precipitation and acid hydrolysis before analysis using liquid chromatography-tandem mass spectrometry.

Non-protein amino acids (NPAAs) are a large class of amino acids (AAs) that are not genetically encoded for translation into proteins. The analysis of ...