This work was supported by a Natural Sciences and Engineering Research (NSERC)-USRA grant to S.J., and by an NSERC-Discovery Grant to I.M. We thank Richard Bourgault, Meghan Rains, and Amanda Fluke for technical assistance. Seeds of att1-1 and att1-2 mutants were kindly provided by Dr. Jian-Min Zhou, Institute of Genetics and Developmental Biology, Beijing, China.

Note: This protocol was adapted from Bonaventure et al. (2004), Molina et al. (2006), Li et al. (2013)7,10,11. Steps 1-5 are summarized in Figure 1.
1. Tissue Delipidation
Note: Always rinse all glassware and caps with chloroform, letting dry under fume hood, before using.
CAUTION: Perform tissue homogenization and all solvent transfer steps under fume hood; always wear lab coat, gloves and splash safety goggles to avoid direct contact with chemicals and to protect the samples from contamination.
<ol>
	<li>Preheat water bath and heat block to 85 &#176;C.</li>
	<li>Weigh approximately 0.5 g of each leaf sample into pre-weighed 20 mm x 125 mm glass test tubes with polytetrafluoroethylene&#160;(PTFE)-faced screw caps. Include four replicates per sample.</li>
	<li>Place 2-propanol in an Erlenmeyer flask (approximately 125 ml; 25 ml per g of sample). Add 2,6-di-tert-butyl-4-methylphenol (also known as butylated hydroxytoluene, BHT) to a final concentration of 0.01% (w/v). 
	Note: Add BHT from a 5% (w/v) stock solution in methanol (BHT helps minimize oxidation of unsaturated fatty acids).</li>
	<li>Preheat 2-propanol solution to 85 &#176;C in water bath.</li>
	<li>Add 12 ml hot 2-propanol solvent to each sample tube and heat for 15 min at 85 &#176;C in heat block. This step inactivates lipases that may be released from disrupted cells.</li>
	<li>Let tubes cool down to room temperature and grind tissue thoroughly with homogenizer until&#160;a homogeneous suspension is obtained.</li>
	<li>Place the samples in an orbital shaker and shake for 1-2 hours at 100 rpm and room temperature.</li>
	<li>Centrifuge for 10 min at 800 x g and discard supernatant.</li>
	<li>Add an equal volume of 2-propanol and shake for 12 hours at room temperature.</li>
	<li>Centrifuge for 10 min at 800 x g and discard supernatant.</li>
	<li>Add 12 ml CHCl3:CH3OH (2:1, v/v) to residue (25 ml per g of sample) and shake overnight at 100 rpm and room temperature.</li>
	<li>Centrifuge for 10 min at 800 x g and discard supernatant.</li>
	<li>Add 12 ml CHCl3:CH3OH (1:2, v/v) to residue and shake overnight at 100 rpm and room temperature.</li>
	<li>Centrifuge for 10 min at 800 x g and remove solvent.</li>
	<li>Let samples dry under fume hood overnight at room temperature.</li>
	<li>Air-dry the residue and then place in a vacuum desiccator over anhydrous CaCl2 or CaSO4 until constant weight is reached (3-5 days).</li>
</ol>
2. Depolymerization: Methanolysis with Sodium Methoxide (Figure 2)
CAUTION: Perform steps 2.3 &#8211; 2.4, 2.7 &#8211; 2.8, 2.10 &#8211; 2.11, 2.13 &#8211; 2.15 and 2.17 &#8211; 2.18 under fume hood; always wear lab coat, gloves and splash safety goggles.
<ol>
	<li>Preheat heat block to 60 &#176;C.</li>
	<li>Weigh test tubes containing the dry residue (important for further calculations).</li>
	<li>Add internal standards to each tube: 25 &#969;L methyl heptadecanoate (1 mg/ml stock) and 25 &#181;l &#969;-pentadecalactone (1 mg/ml stock).</li>
	<li>Add 0.9 ml methyl acetate, 1.5 ml sodium methoxide, and 3.6 ml methanol to each tube and cap them. Alternatively, prepare a reaction mix with these three reagents and add 6-ml aliquots to each sample.</li>
	<li>Heat samples for 2 hours at 60 &#176;C and vortex periodically in 15 min intervals.</li>
	<li>Let samples cool down to room temperature.</li>
	<li>Add 10 ml of methylene dichloride (CH2Cl2) and 1.5 ml of glacial acetic acid to extract the fatty acid methyl esters.</li>
	<li>Add saline solution (0.5 M NaCl) to fill each tube and cap.</li>
	<li>Vortex samples for 1 min and centrifuge for 10 min at 800 x g.</li>
	<li>Transfer the organic phase (lower) to clean medium size tubes (16 x 125 mm glass test tubes with polytetrafluoroethylene&#160;(PTFE)-faced screw cap).</li>
	<li>Add saline solution (0.5 M NaCl) to fill each tube and cap.</li>
	<li>Vortex samples for 1 min and centrifuge for 10 min at 800 x g.</li>
	<li>Remove aqueous (upper) phase and repeat steps 2.11-2.12.</li>
	<li>Remove all aqueous (upper) phase.</li>
	<li>Add anhydrous sodium sulfate (Na2SO4) to the solvent, cap the tubes, and vortex for 1 min; samples may now be left until the following day. At this point samples can be left overnight under the fume hood.</li>
	<li>Centrifuge for 2 min at 800 x g to compact the Na2SO4 salt in the bottom.</li>
	<li>Transfer the organic phase to a small glass disposable tube (13 x 100 mm glass test tubes with polytetrafluoroethylene&#160;(PTFE)-faced screw cap).</li>
	<li>Evaporate the solvent to dryness under nitrogen and proceed to the derivatization step. If not processed immediately, store evaporated samples at -20 &#176;C (samples can be also stored before the evaporation step).</li>
</ol>
3. Preparation of Derivatives for Gas Chromatography
CAUTION: Perform steps 3.1.2 and 3.1.5 &#8211; 3.1.8 under a fume hood; always wear lab coat, gloves and splash safety goggles.
<ol>
	<li>Trimethylsilyl derivatives
	<ol>
		<li>Preheat heat block to 100 &#176;C.</li>
		<li>Add 100 &#181;l of pyridine and 100 &#181;l of BSTFA (N,O Bis-trimethylsilyl-trifluoroacetamide) to each tube and cap them.</li>
		<li>Heat samples at 100 &#176;C for 10 min.</li>
		<li>Let samples cool down to room temperature.</li>
		<li>Evaporate samples under nitrogen at room temperature. Avoid applying heat to samples, monomers are highly volatile at this stage.</li>
		<li>Add 500 &#181;l of 1:1 (v/v) heptane:toluene.</li>
		<li>Vortex samples for 1 min and centrifuge for 2 min at 800 x g.</li>
		<li>Add samples to GC vials and proceed to GC/MS analysis.</li>
	</ol>
	</li>
	<li>Acetyl derivatives 
	Note: For acetylation, modify steps 3.1.1-3.1.3 above as follows (not shown in video); steps 3.1.4-3.1.8 are the same:
	<ol>
		<li>Preheat heat block to 60 &#176;C.</li>
		<li>Add 100 &#181;l of pyridine and 100 &#181;l of acetic anhydride (Ac2O) to the tubes.</li>
		<li>Heat samples 1 hour at 60 &#176;C.</li>
	</ol>
	</li>
</ol>
4. GC/MS Analysis
<ol>
	<li>Use a HP-5 capillary column (30 m x 0.25 mm x 0.25 &#181;m film thickness) or equivalent (i.e., 5% diphenyl,&#160;95% dimethyl polysiloxane). Program the GC with helium carrier gas flow set at 1.5 ml/min and oven temperature programmed from 150 to 300 &#176;C at 3 &#176;C/min.</li>
	<li>Use split injection (split ratio 1:10) and set mass spectrometer to scan mode over 40-600 amu (electron impact ionization).</li>
	<li>Create a sequence table including solvent (blank), WT and mutant replicates, each using the indicated cutin analysis method.</li>
	<li>Load solvent and sample vials on the carrousel, add hexane to syringe-rinsing vials in the autosampler if necessary, and start the sequence. After the sequence is completed, total ion chromatogram traces are available for all samples.</li>
</ol>
5. Data Analysis
<ol>
	<li>Identify lipid polyester monomers by comparing each peak&#8217;s mass spectrum to published mass spectra or by searching a commercial library if available.</li>
	<li>For each identified peak in the total ion current chromatogram, use their respective retention times to find the areas on the integration results table from the GC/MS software (Supplemental Figure 1).</li>
	<li>For each monomer (columns A-B), add the area values found in the integration table (Supplemental Figure 1, column D) to the corresponding column for every replicate sample in the Excel table of monomers (Supplemental File 2, columns C-F), which is a spreadsheet to quantify Arabidopsis cutin TMSi derivatives. Use Supplemental File 3 if acetylated derivatives are prepared instead.</li>
	<li>Add the areas of the internal standards (IS) to the IS columns (H-K). For those monomers that are not derivatized, namely fatty acid methyl esters (FAMEs), and dicarboxylic acid dimethyl esters (DCA DMEs), use 17:0 FAME (IS1) as IS for quantification (purple-shaded cells in the monomer table; Supplemental Files 2/3). For hydroxylated monomers, including primary alcohols, ferulic acid and &#969;-hydroxy acids, use 15:0 15-hydroxy FAME (IS2) as IS of choice for compound quantification (green-shadded cells in the monomer table; Supplemental Files 2/3).</li>
	<li>Add dry leaf weight for each replicate to columns AX-BA (Supplemental File 2) or AQ-AT (Supplemental File 3); alternatively, scan leaves to calculate surface areas and add area values to the monomer table to express monomer loads per unit of surface area.</li>
</ol>

<ol>
	<li>Kolattukudy, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biopolyester+membranes+of+plants:+cutin+and+suberin.">Biopolyester membranes of plants: cutin and suberin.</a> Science. 208 (4447), 990-1000 (1980).</li><li>Kolattukudy, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polyesters+in+higher+plants.">Polyesters in higher plants.</a> Adv. Biochem. Eng. Biot. 71, 1-49 (2001).</li><li>Yeats, T. H., Rose, J. K. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+formation+and+function+of+plant+cuticles.">The formation and function of plant cuticles.</a> Plant Physiol. 163 (1), 5-20 (2013).</li><li>Pollard, M., Beisson, F., Li, Y., Ohlrogge, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Building+lipid+barriers:+biosynthesis+of+cutin+and+suberin.">Building lipid barriers: biosynthesis of cutin and suberin.</a> Trends Plant Sci. 13 (5), 236-246 (2008).</li><li>Beisson, F., Li-Beisson, Y., Pollard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solving+the+puzzles+of+cutin+and+suberin+polymer+biosynthesis.">Solving the puzzles of cutin and suberin polymer biosynthesis.</a> Curr. Opin. Plant Biol. 15 (3), 329-337 (2012).</li><li>Heredia, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biophysical+and+biochemical+characteristics+of+cutin,+a+plant+barrier+biopolymer.">Biophysical and biochemical characteristics of cutin, a plant barrier biopolymer.</a> Biochim. Biophys. Acta. 1620 (1-3), 1-7 (2003).</li><li>Bonaventure, G., Beisson, F., Ohlrogge, J., Pollard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+aliphatic+monomer+composition+of+polyesters+associated+with+Arabidopsis+epidermis:+occurrence+of+octadeca-cis-6,+cis-9-diene-1,+18-dioate+as+the+major+component.">Analysis of the aliphatic monomer composition of polyesters associated with Arabidopsis epidermis: occurrence of octadeca-cis-6, cis-9-diene-1, 18-dioate as the major component.</a> Plant J. 40 (6), 920-930 (2004).</li><li>Franke, 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=Apoplastic+polyesters+in+Arabidopsis+surface+tissues+-+A+typical+suberin+and+a+particular+cutin.">Apoplastic polyesters in Arabidopsis surface tissues - A typical suberin and a particular cutin.</a> Phytochemistry. 66 (22), 2643-2658 (2005).</li><li>Vogg, G., 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=Tomato+fruit+cuticular+waxes+and+their+effects+on+transpiration+barrier+properties:+functional+characterization+of+a+mutant+deficient+in+a+very-long-chain+fatty+acid+-ketoacyl-CoA+synthase.">Tomato fruit cuticular waxes and their effects on transpiration barrier properties: functional characterization of a mutant deficient in a very-long-chain fatty acid -ketoacyl-CoA synthase.</a> J. Exp. Bot. 55 (401), 1401-1410 (2004).</li><li>Molina, I., Bonaventure, G., Ohlrogge, J., Pollard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lipid+polyester+composition+of+Arabidopsis+thaliana+and+Brassica+napus+seeds.">The lipid polyester composition of Arabidopsis thaliana and Brassica napus seeds.</a> Phytochemistry. 67 (23), 2597-2610 (2006).</li><li>Li-Beisson, Y., 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=Acyl-lipid+metabolism.">Acyl-lipid metabolism.</a> The Arabidopsis Book. 11, e0161 (2013).</li><li>Molina, I., Ohlrogge, J. B., Pollard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deposition+and+localization+of+lipid+polyester+in+developing+seeds+of+Brassica+napus+and+Arabidopsis+thaliana.">Deposition and localization of lipid polyester in developing seeds of Brassica napus and Arabidopsis thaliana.</a> Plant J. 53 (3), 437-449 (2008).</li><li>Xiao, F., 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=Arabidopsis+CYP86A2+represses+Pseudomonas+syringae+type+III+genes+and+is+required+for+cuticle+development.">Arabidopsis CYP86A2 represses Pseudomonas syringae type III genes and is required for cuticle development.</a> EMBO J. 23 (14), 2903-2913 (2004).</li><li>Christie, W. W., Han, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Lipid Analysis - Isolation, Separation, Identification and Lipidomic Analysis. , 446 (2010).</li><li>Walton, T. J., Kolattukudy, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+the+structures+of+cutin+monomers+by+a+novel+depolymerization+procedure+and+combined+gas+chromatography+and+mass+spectrometry.">Determination of the structures of cutin monomers by a novel depolymerization procedure and combined gas chromatography and mass spectrometry.</a> Biochemistry. 11 (10), 1885-1896 (1972).</li><li>Matzke, K., Riederer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparative+study+into+the+chemical+constitution+of+cutins+and+suberins+from+Picea+abies+(L.)+Karst.,+Quercus+robur+L.,+and+Fagus+sylvatica+L.">A comparative study into the chemical constitution of cutins and suberins from Picea abies (L.) Karst., Quercus robur L., and Fagus sylvatica L.</a> Planta. 185 (2), 233-245 (1991).</li><li>Riederer, M., Sch&#246;nherr, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+gas+chromatographic+analysis+of+methyl+esters+of+hydroxy+fatty+acids+derived+from+plant+cutin.">Quantitative gas chromatographic analysis of methyl esters of hydroxy fatty acids derived from plant cutin.</a> J. Chromatogr. 360, 151-161 (1986).</li><li>Moire, L., Schmutz, A., Buchala, A., Yan, B., Stark, R., Ryser, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glycerol+Is+a+Suberin+Monomer.+New+Experimental+Evidence+for+an+Old+Hypothesis.">Glycerol Is a Suberin Monomer. New Experimental Evidence for an Old Hypothesis.</a> Plant Physiol. 119 (3), 1137-1146 (1999).</li><li>Gra&#231;a, J., Schreiber, L., Rodrigues, J., Pereira, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glycerol+and+glyceryl+esters+of+omega-hydroxyacids+in+cutins.">Glycerol and glyceryl esters of omega-hydroxyacids in cutins.</a> Phytochemistry. 61 (2), 205-215 (2002).</li><li>Eglinton, G., Hunneman, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gas+chromatographic-mass+spectrometric+studies+of+long+chain+hydroxy+acids-I:+The+constituent+cutin+acids+of+apple+cuticle.">Gas chromatographic-mass spectrometric studies of long chain hydroxy acids-I: The constituent cutin acids of apple cuticle.</a> Phytochemistry. 7 (2), 313-322 (1968).</li><li>Espelie, K. E., K&#246;ller, W., Kolattukudy, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=9,16-dihydroxy-10-oxo-hexadecanoic+acid,+a+novel+component+in+citrus+cutin.">9,16-dihydroxy-10-oxo-hexadecanoic acid, a novel component in citrus cutin.</a> Chem. Phys. Lipids. 32 (1), 13-26 (1983).</li><li>Holloway, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracuticular+lipids+of+spinach+leaves.">Intracuticular lipids of spinach leaves.</a> Phytochemistry. 13 (10), 2201-2207 (1974).</li><li>Holloway, P. J., Deas, A. H. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epoxyoctadecanoic+acids+in+plant+cutins+and+suberins.">Epoxyoctadecanoic acids in plant cutins and suberins.</a> Phytochemistry. 12 (7), 1721-1735 (1973).</li><li>Holloway, P. J., Cutler, D. F., Alvin, K. L., Price, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The chemical constitution of plant cutins. , 45-85 (1982).</li><li>Croteau, R., Fagerson, I. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+constituent+cutin+acids+of+cranberry+cuticle.">The constituent cutin acids of cranberry cuticle.</a> Phytochemistry. 11 (1), 353-363 (1972).</li></ol>

No conflicts of interest declared.

Unlike other biopolymers such as DNA and proteins, plant lipid polyesters are not made from a template. Instead, their compositions depend on the specificity of the enzymes present in the tissues that make these extracellular polymers. As such, chemical analyses of the constituent components are critical to understand lipid polyester composition.
Chemical methods to cleave ester bonds include saponification, hydrogenolysis, acid-catalyzed transmethylation, and base-catalyzed transmethylation2. Each of them has advantages and disadvantages. Saponification produces free fatty hydroxy acids that can undergo secondary reactions. Hydrogenolysis with lithium aluminum hydride (LiAlH4)16 has been used for cutin analysis7. Hydrogenolysis reduces functionalized carbons to alcohols and the original structures need to be inferred by deuteriolysis with lithium aluminum deuteride (LiAlD4). The disadvantage of this approach is the requirement of high resolution GC/MS to compare the degree of deuteriation of the fatty polyols obtained to make assignments of their structures. Acid-catalyzed transesterification with methanolic boron trifluoride (BF3) has been frequently used in cutin and suberin depolymerizations8,17,18, but the reagent has a limited shelf life and may introduce artefacts due to side reactions15. Methanolic sulphuric acid also yields methyl esters of the monomers but with larger proportions of 2-hydroxy fatty acids, which presumably are not true lipid polyester components, compared to other methods10.
The NaOMe-catalyzed transesterification method described in this protocol produces fatty acid methyl esters that are derivatized by silylation of hydroxyl groups, providing characteristic mass spectra for identification, or by acetylation to provide more stable derivatives of hydroxyl groups for quantification. One drawback of this technique is that hydrolysis competes with transesterification when water is present in the reaction. Water reacts with NaOMe (the catalyst) and produces NaOH, which in turn hydrolyses fatty acid methyl esters to yield free acids (Figure 2D). This is an undesirable side reaction because two peaks will be present for each fatty acid: a methyl ester and a TMSi ester derivative, thus complicating the analysis. Using anhydrous reagents and adding methyl acetate as a co-solvent to compete with saponification are thus crucial steps to prevent hydrolysis (Figure 2D).
Cutin and suberin contain between 1 and 26% glycerol4. However, this monomer will not be detected by the experimental conditions described in this protocol. Glycerol is highly hydrophilic and, unlike the fatty acid methyl ester monomers, will be eliminated during the aqueous solvent-washing steps. This limitation also applies to other cutin depolymerization methods, but glycerol can be determined in the aqueous layer obtained after transesterification using an enzymatic method. Alternatively, it can be quantified using milder conditions (e.g., 0.05 M NaOMe) without further water extraction to detect all monomers, including glycerol19,20.Although useful for the purpose of glycerol quantification, mild conditions usually give incomplete depolymerization of cutin and suberin.
If a GC coupled to a flame ionization detector (FID) is available, all replicates can be analyzed in this instrument for quantitative purposes, after peaks of a representative sample have been identified by GC/MS. Alternatively, monomers in the GC/FID traces can be identified if their retention indices are known. The flame ionization detector has especially high sensitivity and a broad range of proportionality, which is critical for quantification of major and minor sample components in single runs. In addition, it is robust and easy to maintain and operate15.
The described protocol allows for the reliable and reproducible isolation, identification, and quantification of plant lipid polyester monomers, allowing the chemical characterization of mutants that differ in the composition of one or more lipid polyester monomers. The procedure is scalable, it can be easily adapted to process both small and bulk quantities of various plant materials, including roots, seeds, leaves, stems and flowers. Mass spectral data of lipid polyester monomers from many species have been publishede.g.,21-26 and constitute valuable resources to identify unknown monomers when adapting this protocol to other tissues and/or species. This method is applicable to investigations of the biosynthesis, regulation, and distribution of lipid polyesters in higher plants.

Vascular plants rely on extracellular layers that function as waterproof barriers between plant tissues and the external environment. These lipophilic cell wall-associated structures restrict pathogenic infection and regulate the passive transport of gases, water, and dissolved substances in and out of plant tissues1. Such barriers are the plant cuticle, a synapomorphic structure unique to plants2, and different suberin-containing diffusion barriers. The cuticle is an oleophilic layer synthesized by epidermal cells and bound to them via a pectinaceous layer on the extracellular side of the cell wall3-5. It encases the primary aerial organs of higher plants, functioning as a vital interface between plant tissues and the environment.
Cutin, the structural matrix of the cuticle, and suberin are two insoluble glycerolipid polyesters associated with solvent-extractable waxes2,4. These polymeric lipids are composed of saturated and unsaturated fatty acid derivatives and are both structurally and functionally similar. However, they are distinguishable by characteristic differences in chemical composition and deposition sites.
Suberin is an aliphatic polyester located inside the cell walls of certain external and internal tissues forming a secondary wall. Suberized tissues include periderms of roots, tubers and tree bark, root endodermis, seed coat layers, and healed wounds2. Unlike cutin, the suberin polyester typically contains alcohols, saturated and mono-unsaturated dicarboxylic acids, and a large proportion of very-long-chain monomers (C&#8805;20).
Cutin is the most abundant lipid polyester in vascular plants6, and is composed of glycerol and C16-C18 interesterified fatty acid derivatives, such as hydroxy and hydroxy-epoxy substituted fatty acids4. While the composition of cutin polymers varies across tracheophyte species, the most predominant primary monomers are 10, 16-dihydroxy 16:0, 18-hydroxy-9,10-epoxy&#160;18:0, and 9,10,18-trihydroxy 18:0 fatty acids. Interestingly, Arabidopsis leaf and stem cutin is mainly composed of 18:2 dicarboxylic acid 7,8.
Plant cuticles also present a considerable variability in thickness, ranging from a few nanometers to several micrometers9. Since cuticle isolation is a laborious and time-consuming step, particularly for very thin leaf cuticles such as those of Arabidopsis thaliana8, methods that bypass cuticle isolation have been developed and validated7,8. Here, we describe a detailed protocol to study the monomer composition of cutin in Arabidopsis thaliana leaves by sodium methoxide (NaOMe)-catalyzed depolymerization and subsequent gas chromatography/mass spectrometry (GC/MS) analysis. This protocol offers a robust method for assaying the composition of plant lipid polyesters in whole delipidated tissues, and has been adapted from previously reported protocols7,10,11. Whole tissue samples are first homogenized and exhaustively delipidated, removing solvent-extractable lipids including cuticular and epicuticular waxes, membrane lipids, and triacylglycerols. Cell wall-enriched residues are then depolymerized into their constituent methyl ester monomers by sodium methoxide-catalyzed methanolysis. Fatty acid methyl esters are extracted upon acidification, and derivatized to obtain their corresponding trimethylsilyl or acetyl derivatives. Derivatized residues are highly volatile, and can be eluted from a gas chromatography column at a reasonable temperature without altering their structural conformation during GC/MS analysis.

<table><tbody><tr><td>Chemicals</td><td></td><td></td><td></td></tr><tr><td>2-propanol</td><td>Fisher Scientific</td><td>BPA451-4</td><td>Solvent for delipidation</td></tr><tr><td>Anhydrous sodium sulfate</td><td>Fisher Scientific</td><td>S421500</td><td></td></tr><tr><td>Acetic anhydride</td><td>Sigma Aldrich</td><td>320102</td><td>Derivatization agent</td></tr><tr><td>BSTFA (N,O-bis(trimethylsilyl)-trifluoroacetamide)</td><td>Sigma-Aldrich</td><td>15222</td><td>Derivatization agent</td></tr><tr><td>Butylated hydroxytoluene (BHT)</td><td>Sigma-Aldrich</td><td>101162</td><td>Antioxidant</td></tr><tr><td>Calcium chloride, anhydrous</td><td>Fisher Scientific</td><td>C614-3</td><td>Desiccation agent</td></tr><tr><td>Calcium suflate, anhydrous (DRIERITE- 8 MESH with indicator)</td><td>Acros Organics</td><td>219090020</td><td>Desiccation agent</td></tr><tr><td>Chloroform (Trichloromethane)</td><td>Fisher Scientific</td><td>C6074</td><td>Organic solvent</td></tr><tr><td>Glacial acetic acid</td><td>Fisher Scientific</td><td>BP2401212</td><td>Acidification agent</td></tr><tr><td>Helium carrier gas, compressed</td><td>Air Liquide</td><td>ALPHAGAZ1-UN1046</td><td>Carrier gas, GC/MS</td></tr><tr><td>Heptane</td><td>Fisher Scientific</td><td>H3501</td><td>Organic solvent</td></tr><tr><td>Hexanes</td><td>Fisher Scientific</td><td>H3024</td><td>Organic solvent</td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>A4124</td><td>Organic solvent, transmethylation reactive</td></tr><tr><td>Methyl acetate</td><td>Sigma-Aldrich</td><td>296996</td><td>Organic solvent</td></tr><tr><td>Methyl heptadecanoate</td><td>Sigma-Aldrich</td><td>H4515</td><td>Internal standard (1mg/mL stock)</td></tr><tr><td>Methylene dichloride (Dichloromethane)</td><td>Fisher Scientific</td><td>D374</td><td>Organic solvent</td></tr><tr><td>Nitrogen, compressed</td><td>Air Liquide</td><td>ALPHAGAZ1-UN1044</td><td>Carrier gas, GC-FID</td></tr><tr><td>Pentadecanolactone</td><td>Fluka</td><td>76530</td><td>Internal standard (1 mg/ml stock)</td></tr><tr><td>Pyridine</td><td>Sigma-Aldrich</td><td>270970</td><td>Co-solvent for derivatization</td></tr><tr><td>Sodium chloride</td><td>Fisher Scientific</td><td>BP358212</td><td>Saline solution</td></tr><tr><td>Sodium methoxide (25wt.%)</td><td>Sigma-Aldrich</td><td>156256</td><td>Nucleophile</td></tr><tr><td>Toluene</td><td>FIsher Scientific</td><td>T2904</td><td>Organic solvent</td></tr><tr><td>Plant Growth Supplies</td><td></td><td></td><td></td></tr><tr><td>Pro-Mix PGX</td><td>Premier Tech Horticulture Ltd</td><td></td><td>Pro-Mix PGX is recommended to grow Arabidopsis plants (Eddy, R. and Hahn, D.T., 2012,http://docs.lib.purdue.edu/pmag/2)&lt;br/&gt; Purdue Methods for Arabidopsis Growth.&amp;nbsp;</td></tr><tr><td>PermaNest Humidity Dome&amp;nbsp;</td><td>Grower&#039;s Solution, LLC, Cookeville, TN&amp;nbsp;</td><td>GD2211-24</td><td></td></tr><tr><td>Perma-Nest Plant Trays (22x11in)</td><td>Grower&#039;s Solution, LLC, Cookeville, TN&amp;nbsp;</td><td>N/A</td><td></td></tr><tr><td>Square greenhouse pots, 3.5 inch&amp;nbsp;</td><td>Grower&#039;s Solution, LLC, Cookeville, TN&amp;nbsp;</td><td>P86</td><td></td></tr><tr><td>General Purpose Plant Fertilizer, Plant-Prod 20-20-20</td><td>Premier Tech Home and Garden In., Brantford, ON</td><td>N/A</td><td></td></tr><tr><td>Glassware</td><td></td><td></td><td></td></tr><tr><td>13 x 100 mm glass test tube with Teflon-faced screw cap</td><td>Kimble Chase Life Science and Research Products LLC</td><td>45066A-13100</td><td></td></tr><tr><td>16 x 125 mm glass test tube with Teflon-faced screw cap</td><td>Kimble Chase Life Science and Research Products LLC</td><td>45066A-16125</td><td></td></tr><tr><td>20 x 125 mm glass test tubes with Teflon-faced screw cap</td><td>Kimble Chase Life Science and Research Products LLC</td><td>45066A-20125</td><td></td></tr><tr><td>GC vial caps</td><td>National Scientific</td><td>C400051A</td><td></td></tr><tr><td>GC vial microinserts</td><td>National Scientific</td><td>C4011631</td><td></td></tr><tr><td>GC vials</td><td>National Scientific</td><td>C40001</td><td></td></tr><tr><td>Disposable pasteur pipets</td><td>Fisher Scientific</td><td>1367820B</td><td></td></tr><tr><td>Flasks</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Equipment</td><td></td><td></td><td></td></tr><tr><td>Allegra X15R centrifuge</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Analytic balance</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Belly dancer</td><td></td><td></td><td>A shaker can be used for this purpose if Belly Dancer not available</td></tr><tr><td>DB-5 Capillary GC column</td><td>J&amp;amp;W Scientific, CA, USA;&amp;nbsp;</td><td></td><td>30 m x 0.25 mm x 0.25 &amp;mu;m film thickness</td></tr><tr><td>Desiccator</td><td></td><td></td><td></td></tr><tr><td>Isotemp 202 water bath</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>ISQ LT single quadupole mass spectrometer</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Heat block</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Nitrogen evaporator</td><td></td><td></td><td></td></tr><tr><td>Polytron homogenizer</td><td>Birkmann</td><td></td><td></td></tr><tr><td>Trace 1300 gas chromatograph</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Two-stage regulator</td><td>Air Liquide</td><td>Q1-318B-580</td><td></td></tr><tr><td>Vacuum desiccator</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Vortex mixer</td><td>Fisher Scientific</td><td></td><td></td></tr></tbody></table>

isolation and compositional analysis of plant cuticle lipid polyester monomers

Terrestrial plants produce extracellular aliphatic biopolyesters that modify cell walls of specific tissues. Epidermal cells synthesize cutin, a polyester of glycerol and modified fatty acids that constitutes the framework of the cuticle that covers aerial plant surfaces. Suberin is a related lipid polyester that is deposited on the cell walls of certain tissues, including the root endodermis and the periderm of tubers, tree bark and roots. These lipid polymers are highly variable in composition among plant species, and often differ among tissues within a single species. Here, we describe a detailed protocol to study the monomer composition of cutin in Arabidopsis thaliana leaves by sodium methoxide (NaOMe)-catalyzed depolymerisation, derivatization, and subsequent gas chromatography-mass spectrometry (GC/MS) analysis. This method can be used to investigate the monomers of insoluble polyesters isolated from whole delipidated plant tissues bearing either cutin or suberin. The method can by applied not only to characterize the composition of lipid polymers in species not previously analyzed, but also as an analytical tool in forward and reverse genetic approaches to assess candidate gene function.

The protocol described in this manuscript is set up to determine lipid polyester monomers (i.e., cutin or suberin), minimizing the contributions of non-cutin lipids10. Figure 1 presents an overview of the assay, which altogether takes between 8 &#8211; 10 days (from initial tissue harvesting to obtaining GC data), depending on how long samples are allowed to dry.
The selected base-catalyzed methanolysis (Figure 2) method to depolymerize polyesters was previously validated for Arabidopsis seeds, which contain both cutin and suberin. Tissues are first homogenized and exhaustively delipidated to remove solvent-extractable lipids. The residue yield after extraction, as percentage of initial fresh weight, is usually 6% for A. thaliana Col-0 leaves. Cell wall-enriched residues are dried in a vacuum desiccator and then depolymerized into their constituent methyl ester monomers by base-catalyzed transmethylation. Two-hour incubation was chosen as the critical time required for proper depolymerization and recovery of lipid polyester components. Longer incubation times resulted in increase in 2-hydroxy acids; these potentially derive from membrane sphingolipids10.
A typical chromatogram if Arabidopsis wild type leaf cutin is shown in Figure 3, for O-TMSi ether derivatives (Figure 3A) and O-acetyl derivatives (Figure 3B). Each peak was identified by comparison to mass spectra from the literature7,8 and a public database12.Our video protocol shows how to prepare TMSi derivatives, but samples can alternatively be acetylated to derivatize hydroxyl groups. Silylated derivatives are good for identification purposes because they give diagnostic mass spectra. However, acetylated derivatives are more stable and a good alternative to silylation once monomers have been identified 10. To help implement this protocol in laboratories that only have GC coupled to flame ionization detector (FID), GC/FID traces corresponding to acetylated derivatives of WT leaf cutin monomers and to a homolog series of fatty acid methyl ester standards are also shown (Supplemental Figure 4).
This method is qualitative and detects quantitative differences between samples, hence its value for mutant analysis. The amounts of individual monomers are determined using the internal standard method of quantification, enabling comparisons of monomer abundance between samples. It should be clarified, however, that the peak size (total ion counts) may not reflect the molar ratios of the monomers in the polyester. We are including editable tables of monomers to calculate monomer amounts in Arabidopsis leaf cutin as fatty acid methyl esters and TMSi derivatives (Supplemental File 1), or acetyl derivatives (Supplemental File 2) of alcohols. These tables may need to be adapted if samples are extracted from different organs or plant species.
As an example, we have analyzed Arabidopsis thaliana Columbia (Col-0) wild type leaves and two previously characterized null-mutant alleles of the CYP86A2/ATT1 gene, att1-1 (m-1) and att1-2 (m-2)13,14. Cytochrome P450 monooxygenases of the CYP86A subfamily encode putative &#969;-oxydases and participate in suberin and cutin monomer biosynthesis. Our results (Figure 4) demonstrate significant reductions in the loads of three major lipid monomers in the mutant leaves compared to WT leaves. Consistent with the enzyme&#8217;s predicted function, 16:0, 18:2, and 18:1 dicarboxylates were specifically affected in att1 mutants.
<img alt="Figure 1" src="/files/ftp_upload/53386/53386fig1.jpg" /> 
Figure 1. Overview of lipid polyester analysis. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/53386fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53386/53386fig2.jpg" /> 
Figure 2. Mechanism of the NaOMe-catalyzed transmethylation reaction. The nucleophilic methoxide anions attack the cabonyl carbon of lipid polyesters to form an unstable tetrahedral intermediate (A), which readily dissociates into fatty acid methyl esters and alkoxide anions (B). These alkoxides are conjugate bases, and react with methanol, regenerating the catalytically active methoxide anions, thereby sustaining additional depolymerization reactions (C). If water is present in the system, it will react with sodium methoxide to form sodium hydroxide, a strong base that irreversibly hydrolyses esters to produce undesirable free fatty acids. Methyl acetate is added as a co-solvent15 to remove small amounts of sodium hydroxide within the system (D). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/53386fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53386/53386fig3.jpg" /> 
Figure 3. Representative total ion chromatogram of wild-type Arabidopsis thaliana leaf cutin monomers. (A) O-trimethylsilyl (TMSi) ether and (B) acetate hydroxyl derivatives. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/53386fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53386/53386fig4.jpg" /> 
Figure 4. Cutin monomer composition of Arabidopsis thaliana WT and two null mutant alleles of the CYP86A2 gene (mutant-1=att1-1; mutant-2=att1-2). Error bars represent standard deviation of the mean (n = 4). Adapted from 13, with permission of &#169; Blackwell Publishing (2007). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/53386fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 1" src="/files/ftp_upload/53386/53386supfig1.jpg" /> 
Supplemental Figure 1. Peak integration results table from the GC/MS software. Peaks corresponding to identified monomers and internal standards are identified by their retention time (column C) and area values are tabulated in column D. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/Supplemental_Figure_1.tif" target="_blank">Please click here to download this file.</a>
<img alt="Figure 2" src="/files/ftp_upload/53386/53386supfig2.jpg" /> 
Supplemental File&#160;2. Table of Arabidopsis cutin monomers (trimethylsilyl ether derivatives of hydroxy fatty acid methyl esters). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/Supplemental_File_2_Cutin_template_TMSi.xls" target="_blank">Please click here to download this file.</a>
<img alt="Figure 3" src="/files/ftp_upload/53386/53386supfig3.jpg" /> 
Supplemental File 3. Table of Arabidopsis cutin monomers (O-acetyl derivatives of hydroxy fatty acid methyl esters). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/Supplemental_File_3_Cutin_template_Acetylated_derivatives.xlsx" target="_blank">Please click here to download this file.</a>
<img alt="Figure 4" src="/files/ftp_upload/53386/53386supfig4.jpg" /> 
Supplemental Figure 4. GC/FID traces of (A-B) fatty acid methyl ester (FAME) retention index standards (peaks are labeled with each saturated FAME chain length); and (C) acetylated A. thaliana WT leaf cutin monomers. Numbers on peak correspond to: 16:0 FAME (1), ferulate (2), 18:3 FAME (3), 18:1/18:2 FAMEs (4), 18:0 FAME (5), sinapate (6), 16:0 DCA (7), 16-OH 16:0 FAME (8), 18:2 DCA (9), 18:1 DCA (10) 18:0 DCA (11), 18-OH 18:2 FAME (12), 18-OH 18:1 FAME (13), 20:0 FAME (14), 10,16-diOH 16:0 FAME (15), 24:0 FAME (16). DCA: dicarboxylic acid dimethyl ester; FAME: fatty acid methyl ester; IS1: internal standard 1, 17:0 FAME; IS2: internal standard 2, 15-OH 15:0 FAME. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53386/Supplemental_Figure_4.tif" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Isolation and Compositional Analysis of Plant Cuticle Lipid Polyester Monomers at JoVE.com

Isolation and Compositional Analysis of Plant Cuticle Lipid Polyester Monomers

Lipid polyesters constitute the structural components of two cell wall modifications, the plant cuticle and suberin-containing diffusion barriers. In this video, we describe a method to depolymerize cutin from whole delipidated leaves. The method can be applied to investigating mutants compromised in either cutin or suberin biosynthesis.

Terrestrial plants produce extracellular aliphatic biopolyesters that modify cell walls of specific tissues. Epidermal cells synthesize cutin, a polyester ...

isolation-compositional-analysis-plant-cuticle-lipid-polyester

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

16.7K Views.  Algoma University. Lipid polyesters constitute the structural components of two cell wall modifications, the plant cuticle and suberin-containing diffusion barriers. In this video, we describe a method to depolymerize cutin from whole delipidated leaves. The method can be applied to investigating mutants compromised in either cutin or suberin biosynthesis.

Video: Isolation and Compositional Analysis of Plant Cuticle Lipid Polyester Monomers

Isolation and Chemical Characterization of Lipid A from Gram-negative Bacteria

Lipopolysaccharide (LPS) is the major cell surface molecule of gram-negative bacteria, deposited on the outer leaflet of the outer membrane bilayer. LPS can be subdivided into three domains: the distal O-polysaccharide, a core oligosaccharide, and the lipid A domain consisting of a lipid A molecular species and 3-deoxy-D-manno-oct-2-ulosonic acid residues (Kdo). The lipid A domain is the only component essential for bacterial cell survival. Following its synthesis, lipid A is chemically modified in response to environmental stresses such as pH or temperature, to promote resistance to antibiotic compounds, and to evade recognition by mediators of the host innate immune response. The following protocol details the small- and large-scale isolation of lipid A from gram-negative bacteria. Isolated material is then chemically characterized by thin layer chromatography (TLC) or mass-spectrometry (MS). In addition to matrix-assisted laser desorption/ionization-time of flight (MALDI-TOF) MS, we also describe tandem MS protocols for analyzing lipid A molecular species using electrospray ionization (ESI) coupled to collision induced dissociation (CID) and newly employed ultraviolet photodissociation (UVPD) methods. Our MS protocols allow for unequivocal determination of chemical structure, paramount to characterization of lipid A molecules that contain unique or novel chemical modifications. We also describe the radioisotopic labeling, and subsequent isolation, of lipid A from bacterial cells for analysis by TLC. Relative to MS-based protocols, TLC provides a more economical and rapid characterization method, but cannot be used to unambiguously assign lipid A chemical structures without the use of standards of known chemical structure. Over the last two decades isolation and characterization of lipid A has led to numerous exciting discoveries that have improved our understanding of the physiology of gram-negative bacteria, mechanisms of antibiotic resistance, the human innate immune response, and have provided many new targets in the development of antibacterial compounds.

Isolation and characterization of the lipid A domain of lipopolysaccharide (LPS) from gram-negative bacteria provides insight into cell surface based mechanisms of antibiotic resistance, bacterial survival and fitness, and how chemically diverse lipid A molecular species differentially modulate host innate immune responses.

Lipopolysaccharide (LPS) is the major cell surface molecule of gram-negative bacteria, deposited on the outer leaflet of the outer membrane bilayer. LPS ...

Chemistry

Isolation and Preparation of Bacterial Cell Walls for Compositional Analysis by Ultra Performance Liquid Chromatography

The bacterial cell wall is critical for the determination of cell shape during growth and division, and maintains the mechanical integrity of cells in the face of turgor pressures several atmospheres in magnitude. Across the diverse shapes and sizes of the bacterial kingdom, the cell wall is composed of peptidoglycan, a macromolecular network of sugar strands crosslinked by short peptides. Peptidoglycan&#8217;s central importance to bacterial physiology underlies its use as an antibiotic target and has motivated genetic, structural, and cell biological studies of how it is robustly assembled during growth and division. Nonetheless, extensive investigations are still required to fully characterize the key enzymatic activities in peptidoglycan synthesis and the chemical composition of bacterial cell walls. High Performance Liquid Chromatography (HPLC) is a powerful analytical method for quantifying differences in the chemical composition of the walls of bacteria grown under a variety of environmental and genetic conditions, but its throughput is often limited. Here, we present a straightforward procedure for the isolation and preparation of bacterial cell walls for biological analyses of peptidoglycan via HPLC and Ultra Performance Liquid Chromatography (UPLC), an extension of HPLC that utilizes pumps to deliver ultra-high pressures of up to 15,000 psi, compared with 6,000 psi for HPLC. In combination with the preparation of bacterial cell walls presented here, the low-volume sample injectors, detectors with high sampling rates, smaller sample volumes, and shorter run times of UPLC will enable high resolution and throughput for novel discoveries of peptidoglycan composition and fundamental bacterial cell biology in most biological laboratories with access to an ultracentrifuge and UPLC.

The bacterial cell wall is composed of peptidoglycan, a macromolecular network of sugar strands crosslinked by peptides. Ultra Performance Liquid Chromatography provides high resolution and throughput for novel discoveries of peptidoglycan composition. We present a procedure for the isolation of cell walls (sacculi) and their subsequent preparation for analysis via UPLC.

The bacterial cell wall is critical for the determination of cell shape during growth and division, and maintains the mechanical integrity of cells in the ...

Use of Arabidopsis eceriferum Mutants to Explore Plant Cuticle Biosynthesis

The plant cuticle is a waxy outer covering on plants that has a primary role in water conservation, but is also an important barrier against the entry of pathogenic microorganisms.  The cuticle is made up of a tough crosslinked polymer called "cutin" and a protective wax layer that seals the plant surface.  The waxy layer of the cuticle is obvious on many plants, appearing as a shiny film on the ivy leaf or as a dusty outer covering on the surface of a grape or a cabbage leaf thanks to light scattering crystals present in the wax.  Because the cuticle is an essential adaptation of plants to a terrestrial environment, understanding the genes involved in plant cuticle formation has applications in both agriculture and forestry. Today, we'll show the analysis of plant cuticle mutants identified by forward and reverse genetics approaches.


The plant cuticle is a waxy outer covering on plants that has a primary role in water conservation but is also an important barrier against the entry of pathogenic microorganisms. In this video, we demonstrate the analysis of plant cuticle mutants identified by forward and reverse genetics approaches.

The plant cuticle is a waxy outer covering on plants that has a primary role in water conservation, but is also an important barrier against the entry of ...

Chemical Isolation, Quantification, and Separation of Skin Lipids from Reptiles

Reptiles signal to conspecifics using lipids in their skin, primarily to enable mate tracking and assessment. The isolation of these lipids has utility in research focused on evolutionary patterns and mechanisms of chemical communication, in addition to understanding the waterproofing role of lipids in the evolution of terrestrial life. In an applied approach, such skin-based cues have potential use for wildlife managers dealing with invasive species. The main steps for quantifying reptile skin lipids in the protocol presented here include extraction, total lipid determination, and fractionation via column chromatography, the latter process resulting in purified eluates of compounds which can then either be analyzed to assign compound identifications (e.g., gas chromatography-mass spectrometry [GC-MS]) and/or used directly in more refined bioassays. Skin lipids can be extracted from living skin, shed skin, or dead whole animals, using nonpolar organic solvents (e.g., hexane, benzene, toluene). Extraction solubilizes the lipids and, then, the solvent can be evaporated to yield a measurable lipid-only extract. Fractionation involves the separation of the total lipid extract into specific eluates via traditional column chromatography. The total lipid extract is first bound to a substrate-based column (e.g., alumina) and, then, individual eluates (&#34;fractions&#34;) of solvent at specific volumes are passed sequentially through the column to elute sets of compounds from the lipid mixture based on common polarity. The fractions progress in polarity at a standardized sequence by increasing the relative amount of polar solvent (e.g., diethyl ether) in nonpolar solvent. In this manuscript, we describe several methods for extracting skin lipids of reptiles and, then, provide a standard protocol for isolating different sets of compounds based on polarity, using traditional column chromatography. Whole lipid extracts or specific fractions can, then, be used in bioassays to determine any biological activity elicited by the compounds therein.

In reptiles, skin lipids from conspecifics are crucial for sexual signaling, with potential use in invasive species management. Here, we describe protocols for extracting skin lipids from shed skin or whole animals, determining and analyzing the total lipid mass, and separating the lipids using fractionation via column chromatography.

Reptiles signal to conspecifics using lipids in their skin, primarily to enable mate tracking and assessment. The isolation of these lipids has utility in ...

Environment

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 ...

Comprehensive Compositional Analysis of Plant Cell Walls (Lignocellulosic biomass) Part II: Carbohydrates

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 transportation2 and other products such as biocomposite materials6. 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, and the polyphenol lignin5 and thus sometimes termed lignocellulosics. However, plant cell walls have evolved to be recalcitrant to degradation as walls contribute extensively to the strength and structural integrity of the entire plant. Despite its necessary rigidity, the cell wall is a highly dynamic entity that is metabolically active and plays crucial roles in numerous cell activities such as plant growth and differentiation5. 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 depolymerisation 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 (Figure 1). The method starts of with preparing destarched cell wall material. The resulting lignocellulosics are then split up to determine its monosaccharide composition of the hemicelluloses and other matrix polysaccharides1, and its content of crystalline cellulose7. The protocol for analyzing the lignin components in lignocellulosic biomass is discussed in Part I3.

Comprehensive Compositional Analysis of Plant Cell Walls Lignocellulosic biomass Part II: Carbohydrates

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 wall derived carbohydrates.

Arabidopsis thaliana Polar Glycerolipid Profiling by Thin Layer Chromatography (TLC) Coupled with Gas-Liquid Chromatography (GLC)

Biological membranes separate cells from the environment. From a single cell to multicellular plants and animals, glycerolipids, such as phosphatidylcholine or phosphatidylethanolamine, form bilayer membranes which act as both boundaries and interfaces for chemical exchange between cells and their surroundings. Unlike animals, plant cells have a special organelle for photosynthesis, the chloroplast. The intricate membrane system of the chloroplast contains unique glycerolipids, namely glycolipids lacking phosphorus: monogalactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG), sulfoquinovosyldiacylglycerol (SQDG)4. The roles of these lipids are beyond simply structural. These glycolipids and other glycerolipids were found in the crystal structures of photosystem I and II indicating the involvement of glycerolipids in photosynthesis8,11. During phosphate starvation, DGDG is transferred to extraplastidic membranes to compensate the loss of phospholipids9,12.
Much of our knowledge of the biosynthesis and function of these lipids has been derived from a combination of genetic and biochemical studies with Arabidopsis thaliana14. During these studies, a simple procedure for the analysis of polar lipids has been essential for the screening and analysis of lipid mutants and will be outlined in detail. A leaf lipid extract is first separated by thin layer chromatography (TLC) and glycerolipids are stained reversibly with iodine vapor. The individual lipids are scraped from the TLC plate and converted to fatty acyl methylesters (FAMEs), which are analyzed by gas-liquid chromatography coupled with flame ionization detection (FID-GLC) (Figure 1). This method has been proven to be a reliable tool for mutant screening. For example, the tgd1,2,3,4 endoplasmic reticulum-to-plastid lipid trafficking mutants were discovered based on the accumulation of an abnormal galactoglycerolipid: trigalactosyldiacylglycerol (TGDG) and a decrease in the relative amount of 18:3 (carbons : double bonds) fatty acyl groups in membrane lipids 3,13,18,20. This method is also applicable for determining enzymatic activities of proteins using lipids as substrate6.

Arabidopsis thaliana Polar Glycerolipid Profiling by Thin Layer Chromatography TLC Coupled with Gas-Liquid Chromatography GLC

Composition of polar lipid extracts and the fatty acid composition of individual glycerolipids are determined in a simple and robust lipid profiling experiment. For this purpose, glycerolipids are isolated by thin layer chromatography and subjected to transmethylation of their acyl groups. Fatty acyl methylesters are quantified by gas-liquid chromatography.

Biological membranes separate cells from the environment. From a single cell to multicellular plants and animals, glycerolipids, such as ...

Isolation and Biophysical Study of Fruit Cuticles

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and abiotic stresses and also regulates water flow from the external environment.1 A biopolyester (cutin) and long-chain fatty acids (waxes) form the principal structural framework of the cuticle; the functional integrity of the cuticular layer depends on the outer 'epicuticular' layer as well as the blend consisting of the cutin biopolymer and 'intracuticular' waxes.2 Herein, we describe a comprehensive protocol to extract waxes exhaustively from commercial tomato (Solanum lycopersicum) fruit cuticles or to remove epicuticular and intracuticular waxes sequentially and selectively from the cuticle composite. The method of Jetter and Sch&auml;ffer (2001) was adapted for the stepwise extraction of epicuticular and intracuticular waxes from the fruit cuticle.3,4 To monitor the process of sequential wax removal, solid-state cross-polarization magic-angle-spinning (CPMAS) 13C NMR spectroscopy was used in parallel with atomic force microscopy (AFM), providing molecular-level structural profiles of the bulk materials complemented by information on the microscale topography and roughness of the cuticular surfaces. To evaluate the cross-linking capabilities of dewaxed cuticles from cultivated wild-type and single-gene mutant tomato fruits, MAS 13C NMR was used to compare the relative proportions of oxygenated aliphatic (CHO and CH2O) chemical moieties.
Exhaustive dewaxing by stepwise Soxhlet extraction with a panel of solvents of varying polarity provides an effective means to isolate wax moieties based on the hydrophobic characteristics of their aliphatic and aromatic constituents, while preserving the chemical structure of the cutin biopolyester. The mechanical extraction of epicuticular waxes and selective removal of intracuticular waxes, when monitored by complementary physical methodologies, provides an unprecedented means to investigate the cuticle assembly: this approach reveals the supramolecular organization and structural integration of various types of waxes, the architecture of the cutin-wax matrix, and the chemical composition of each constituent. In addition, solid-state 13C NMR reveals differences in the relative numbers of CHO and CH2O chemical moieties for wild-type and mutant red ripe tomato fruits. The NMR techniques offer exceptional tools to fingerprint the molecular structure of cuticular materials that are insoluble, amorphous, and chemically heterogeneous. As a noninvasive surface-selective imaging technique, AFM furnishes an effective and direct means to probe the structural organization of the cuticular assembly on the nm-&mu;m length scale.

Aerial plant organs are protected by the cuticle, a supramolecular biopolyester-wax assembly. We present protocols to monitor selective removal of epi- and intracuticular waxes from tomato fruit cuticles on molecular and micro scales by solid-state NMR and atomic force microscopy, respectively, and to assess the cross-linking capacity of engineered cuticular biopolyesters.

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and ...

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they are believed to serve a central role in the adaptation and remodeling of the thylakoid membrane under rapidly changing environmental conditions. The capacity to isolate plastoglobules of high purity has greatly facilitated their study through proteomic, lipidomic, and other methodologies. With plastoglobules of high purity and yield, it is possible to investigate their lipid and protein composition, enzymatic activity, and protein topology, among other possible molecular characteristics. This article presents a rapid and effective protocol for the isolation of plastoglobules from chloroplasts of plant leaf tissue and presents methodological variations for the isolation of plastoglobules and related lipid droplet structures from maize leaves, the desiccated leaf tissue of the resurrection plant, Eragrostis nindensis, and the cyanobacterium, Synechocystis sp. PCC 6803. Isolation relies on the low density of these lipid-rich particles, which facilitates their purification by sucrose density flotation. This methodology will prove valuable in the study of plastoglobules from diverse species.

A fast and efficient protocol is presented for the isolation of plastoglobule lipid droplets associated with various photosynthetic organisms. The successful preparation of isolated plastoglobules is a crucial first step that precedes detailed molecular investigations such as proteomic and lipidomic analyses.

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they ...