Wir danken Andrew Carroll, Bright Chaibang, Purbasha Sarkar (Energy Biosciences Institute, Berkeley), Bahram Parvin (Lawrence Berkeley National Laboratory) und Vincent L. Chiang (North Carolina State University) für eine fruchtbare Zusammenarbeit und hilfreiche Diskussionen. Diese Arbeit wurde von der Energy Biosciences Institute unterstützt. Die Arbeit in der Molecular Foundry wurde durch das Office of Science, Office of Basic Energy Sciences des US Department of Energy unter Vertrag Nr. DE-AC02-05CH1123 unterstützt.

1. Probenvorbereitung <ol><li> Montieren Sie die Anlage hydratisiert Probe, z. B. Pappel Stammholz oder Arabidopsis thaliana Stamm, in dem Mikrotom. </li><li> Cut Dünnschliffen (typischerweise 20 &amp;mgr; m dick) aus dem nativen Gewebe. </li><li> Übertragen Sie die Teilanlage auf einen Glasobjektträger. </li><li> Weichen Sie das Anlagenteil in D 2 O und mit einem Deckglas, das auf den Objektträger versiegelt wird, um die Verdampfung des D 2 O. verhindern Die Anlage Abschnitt ist nun bereit für die Bildgebung oder es kann für eine spätere Verwendung gespeichert werden. </li></ol> 2. Probenmessung <ol><li> Bewerben Immersionsöl zum Mikroskopobjektiv und / oder das Deckglas. </li><li> Ort und sichern die Objektträger auf dem piezoelektrischen Scan des Mikroskops, mit dem Deckglas vor dem Mikroskopobjektiv. </li><li> Sehen Sie sich die Probe durch das Deckglas mit einer hohen numerischen Apertur Eintauchen Mikroskop-Objektiv (100x, NA = 1,40) und suchen Sie die Probe Gebiet von Interesse. </li><li> Nach dem Ausschalten aller anderen Labor-und Mikroskop-Lichtquellen, ortsaufgelöste mikrospektroskopischen Messungen werden durch die Fokussierung bandpassgefilterten monochromatisches grünes Licht (λ = 532 nm) aus einem cw-Laser auf die Probe mit einer typischen Leistungsaufnahme von 10 bis 30 mW durchgeführt ( siehe Abbildung 1 für eine schematische Darstellung der Einrichtung). Autofluoreszenz kann in einigen Proben kommen, die brauchbare Messungen zu verbieten, in welchem ​​Fall der Anregung mit langwelliger Laserlicht kann es ratsam sein. </li><li> Die rückgestreuten Stokes-Raman verschobenen Licht durch das Mikroskop-Objektiv gesammelt, geht durch einen dichroitischen Spiegel, eine Lochkamera, die als räumliche Filter in der konfokalen Aufbau dient, und ein Langpassfilter und wird in den Schlitz eines Gitters konzentriert Spektrometer, wo das Licht spektral zerlegt und detektiert wird durch eine gekühlte CCD-Kamera, so dass ein Raman-Spektrum. Ein Raman-Spektrum von Pappelholz ist in Abbildung 2 gezeigt, mit charakteristischen Lignin Bands im Spektralbereich zwischen 1.600 und 1.700 cm -1. </li><li> Für die chemische Bildgebung und Visualisierung der räumlichen Verteilung Lignin ist ein zweidimensionales spektrale Karte von Rastern der Probe durch den Laserfokus mit dem piezoelektrischen Scan Bühne und die Aufnahme eines Raman-Spektrum für jede Probe Position erworben. Dreidimensionale spektrale Karten kann durch Stapeln zweidimensionale Karten, für die der Laserfokus nacheinander entlang der z-Richtung wurde verstärkt erzeugt werden. </li></ol> 3. Data Analysis <ol><li> Für die chemische Bildgebung und Lignin Visualisierung, werden die erhobenen Daten analysiert mit Hilfe von MATLAB (MathWorks, Version 7,7). Die Daten werden in einem dreidimensionalen hyperspectral Würfel, welche der beiden räumlichen Dimensionen und eine dritte Dimension für die spektrale Signale zusammen angeordnet ist. </li><li> Für das Lignin Analyse wird der Spektralbereich von 1550 und 1700 cm -1 berücksichtigt werden (siehe Abbildung 2). Die räumliche Verteilung von Lignin wird visualisiert durch die Integration der Intensität von 1.550 bis 1.700 cm -1 der Baseline-korrigierten Spektren (siehe Abbildung 3). Als Alternative zur Baseline-Korrektur kann der zweiten Ableitung Spektren berechnet und der zweiten Ableitung Gipfel zur Analyse verwendet. </li><li> . Lignin Lokalisierung und Chemie, insbesondere im Hinblick auf coniferaldehyde und Coniferylalkohol-Einheiten können weitere durch die Auswertung der Fläche unter ausgestattet Gauß-Peaks der drei Bänder zwischen 1.600 und 1.700 cm -1 gefunden (siehe Einschub in Abbildung 2 und Refs 15 analysiert werden - 17). </li><li> Intensity Normalisierung zwischen verschiedenen spektralen Karten erfolgt über ein Verweis auf die Peakhöhe des extrinsischen OD-Bande rund 2.500 cm -1 im Durchschnitt Lumen-Spektren, die durch k-means Clustering Klassifizierung erhalten werden. Dies ist entscheidend und ermöglicht es, Lignin Signalintensitäten zwischen verschiedenen Messungen, Gewebe, Proben und Arten zu vergleichen. </li></ol> 4. Repräsentative Ergebnisse Ein Vertreter Raman-Spektrum von Pappel (Populus angustifolia) Stammholz ist in Abbildung 2 dargestellt. Charakteristisch Lignin Bands sind im Spektralbereich zwischen 1.600 und 1.700 cm -1 gefunden. Als Beispiel ist die räumliche Verteilung von Lignin in einer Pappel Querschnitt in Abbildung 3 dargestellt. Verglichen mit dem sichtbaren Bild, werden morphologisch Zellwand Regionen deutlich unterscheidbar durch verschiedene Lignin Signalintensität. Hohe Lignin Signalintensität in der Zelle Ecken (CC) zu beobachten und, etwas weniger, in der Verbindung Mittellamellen (CML). Niedrigere, aber nicht unerhebliche, Mengen von Lignin sind innerhalb der S2 Wandschicht der Fasern beobachtet. Variabilität von Lignin Signalintensität zu einem gewissen Grad in CC, CML und S2 gefunden, vor allem von Faser zu Faser. Die laterale räumliche Auflösung in unseren Messungen ist ~ 300 nm. Die Qualität der Daten eignet sich gut zur Verholzung zwischen vergleichenBeispiele und weitere sezieren Ligninchemie 15. <img src="/files/ftp_upload/2064/2064fig1.jpg" alt="Abbildung 1" /> Abbildung 1: Eine schematische Darstellung der instrumentalen Setup BP:. Bandpass-Filter, DM: dichroitischen Spiegel, PH: Lochkamera, LP: Langpassfilter. <img src="/files/ftp_upload/2064/2064fig2.jpg" alt="Abbildung 2" /> Abbildung 2: Ein Vertreter Raman-Spektrum von Pappel (Populus angustifolia) Stammholz in D 2 O aufgenommen. Der hervorgehobene Spektralbereich (siehe auch Kasten) markiert den Spektralbereich mit drei Gipfeln zuordenbare Lignin. <img src="/files/ftp_upload/2064/2064fig3.jpg" alt="Abbildung 3" /> Abbildung 3: Raman Lignin Bild (unten) einer Pappel Querschnitt (oben: sichtbares Bild), durch die Integration der Raman-Signal-Intensität von 1.550 bis 1.700 cm -1 erhalten.

<ol>
	<li>Herrera, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bonkers+about+biofuels.">Bonkers about biofuels.</a> Nat Biotechnol. 24, 755-760 (2006).</li><li>Himmel, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biomass+recalcitrance:+Engineering+plants+and+enzymes+for+biofuels+production.">Biomass recalcitrance: Engineering plants and enzymes for biofuels production.</a> Science. 315, 804-807 (2007).</li><li>Pauly, M., Keegstra, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-wall+carbohydrates+and+their+modification+as+a+resource+for+biofuels.">Cell-wall carbohydrates and their modification as a resource for biofuels.</a> Plant J. 54, 559-568 (2008).</li><li>Pauly, M., Keegstra, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiology+and+metabolism+'Tear+down+this+wall.">Physiology and metabolism 'Tear down this wall.</a> Curr Opin Plant Biol. 11, 233-235 (2008).</li><li>Ragauskas, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+path+forward+for+biofuels+and+biomaterials.">The path forward for biofuels and biomaterials.</a> Science. 311, 484-489 (2006).</li><li>Somerville, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biofuels.">Biofuels.</a> Curr Biol. 17, R115-R119 (2007).</li><li>Ralph, J., Brunow, G., Boerjan, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Lignins in Encyclopedia of Life Sciences. , (2007).</li><li>Chiang, V. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+rags+to+riches.">From rags to riches.</a> Nat Biotechnol. 20, 557-558 (2002).</li><li>Atalla, R. H., Agarwal, U. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Raman+microprobe+evidence+for+lignin+orientation+in+the+cell+walls+of+native+woody+tissue.">Raman microprobe evidence for lignin orientation in the cell walls of native woody tissue.</a> Science. 227, 636-638 (1985).</li><li>Atalla, R. H., Agarwal, U. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recording+Raman+spectra+from+plant+cell+walls.">Recording Raman spectra from plant cell walls.</a> J Raman Spectrosc. 17, 229-231 (1986).</li><li>Fukushima, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+syringyl+to+guaiacyl+ratio+in+lignin+biosynthesis.">Regulation of syringyl to guaiacyl ratio in lignin biosynthesis.</a> J Plant Res. 114, 499-508 (2001).</li><li>Agarwal, U. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Raman+imaging+to+investigate+ultrastructure+and+composition+of+plant+cell+walls:+distribution+of+lignin+and+cellulose+in+black+spruce+wood+(Picea+mariana).">Raman imaging to investigate ultrastructure and composition of plant cell walls: distribution of lignin and cellulose in black spruce wood (Picea mariana).</a> Planta. 224, 1141-1153 (2006).</li><li>Gierlinger, N., Schwanninger, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+imaging+of+poplar+wood+cell+walls+by+confocal+Raman+microscopy.">Chemical imaging of poplar wood cell walls by confocal Raman microscopy.</a> Plant Physiol. 140, 1246-1254 (2006).</li><li>Gierlinger, N., Schwanninger, 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+potential+of+Raman+microscopy+and+Raman+imaging+in+plant+research.">The potential of Raman microscopy and Raman imaging in plant research.</a> Spectrosc Int J. 21, 69-89 (2007).</li><li>Schmidt, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+in+situ+imaging+of+lignification+in+the+cell+wall+of+low+lignin+transgenic+Populus+trichocarpa.">Label-free in situ imaging of lignification in the cell wall of low lignin transgenic Populus trichocarpa.</a> Planta. 230, 589-597 (2009).</li><li>Agarwal, U. P., Argyropoulos, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+Overview+of+Raman+Spectroscopy+as+Applied+to+Lignocellulosic+Materials.">An Overview of Raman Spectroscopy as Applied to Lignocellulosic Materials.</a> Advances in Lignocellulosics Characterization. , 201-225 (1999).</li><li>Agarwal, U. P., Ralph, S. A. <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+ethylenic+residues+in+wood+and+TMP+of+spruce+by+FT-Raman+spectroscopy.">Determination of ethylenic residues in wood and TMP of spruce by FT-Raman spectroscopy.</a> Holzforschung. 62, 667-675 (2008).</li><li>Schmidt, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Raman+imaging+of+cell+wall+polymers+in+Arabidopsis+thaliana.">Raman imaging of cell wall polymers in Arabidopsis thaliana.</a> Biochem Biophys Res Comm. 395, 521-523 (2010).</li></ol>

Lignocellulose-Materialien sind hierarchische und heterogene sowohl im Hinblick auf Struktur und Zusammensetzung. Für eine eingehende Charakterisierung analytische Tools, die chemische Empfindlichkeit, räumliche Auflösung, und das haben, geben Einblick in diese Materialien in der nativen Kontext sind wünschenswert. Das beschriebene Verfahren bietet die Visualisierung von Lignin und Vergleich der Verholzung von Lignocellulose-Biomasse-Anlage mit einer räumlichen Auflösung, die Sub-um-ohne Färbung oder die Kennzeic...

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<td>pointed brush</td>
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label-free in situ imaging of lignification in plant cell walls

Meeting wachsenden Energiebedarf sicher und effizient ist eine dringende globale Herausforderung. Deshalb hat sich die Forschung in die Produktion von Biokraftstoffen, die kosteneffiziente und nachhaltige Lösungen zu finden sucht sich einen aktuellen und kritischen Aufgabe. Lignocellulose-Biomasse ist bereit, die primäre Quelle von Biomasse für die Umstellung auf flüssige Biokraftstoffe 1-6 geworden. Jedoch stellt die Widerspenstigkeit dieser pflanzlichen Zellwand Materialien zu kostengünstigen und effizienten Abbau ein großes Hindernis für ihre Verwendung bei der Herstellung von Biokraftstoffen und Chemikalien 4. Insbesondere wird Lignin, eine komplexe und unregelmäßige poly-Phenylpropanoid Heteropolymer, problematisch, die Ernte Dekonstruktion von Lignocellulose-Biomasse. Zum Beispiel in die Umwandlung von Biomasse für Biokraftstoffe, hemmt es Verzuckerung in Prozesse bei der Herstellung einfacher Zucker für die Gärung 7 ausgerichtet. Die effektive Nutzung pflanzlicher Biomasse für industrielle Zwecke ist in der Tat weitgehend auf das Ausmaß, in dem die pflanzliche Zellwand ist verholzt. Die Entfernung von Lignin ist ein kostspieliges und limitierende Faktor 8 und Lignin ist daher ein wesentlicher Pflanzenzüchtung und Gentechnik Ziel geworden, um Zellwand der Konvertierung zu verbessern. Analytische Werkzeuge, die die genaue schnelle Charakterisierung der Verholzung der Zellwände von Pflanzen erlauben zunehmend an Bedeutung für die Beurteilung einer großen Anzahl von Populationen. Extractive Verfahren zur Isolierung von nativen Komponenten wie Lignin sind zwangsläufig zerstörerisch, bringt über wesentliche chemische und strukturelle Modifikationen 9-11. Analytische Chemie in-situ-Verfahren sind somit wertvolle Werkzeuge für das kompositorische und strukturelle Charakterisierung von Lignocellulose-Materialien. Raman-Mikroskopie ist eine Technik, die auf unelastisch oder Raman-Streuung von monochromatischem Licht setzt, wie aus einem Laser, wenn die Verschiebung in Energie des Lasers Photonen Molekülschwingungen verbunden ist und stellt eine intrinsische label-free molekularen &quot;Fingerabdruck&quot; der Probe . Raman-Mikroskopie kann nicht-destruktiv und vergleichsweise kostengünstige Messungen mit minimaler Probenvorbereitung leisten, mit Einblicken in chemische Zusammensetzung und die molekulare Struktur in der Nähe von nativen Zustand. Chemical Imaging mittels konfokaler Raman-Mikroskopie wurde bereits für die Visualisierung der räumlichen Verteilung von Cellulose und Lignin im Holz Zellwände 12-14 verwendet. Basierend auf diesen früheren Ergebnissen haben wir kürzlich diese Methode, um Verholzung in Wildtyp-und Lignin-defizienten transgenen Populus trichocarpa (schwarz Pappel) Stammholz 15 Vergleichen angenommen. Die Analyse der Lignin Raman-Banden 16,17 im Spektralbereich zwischen 1.600 und 1.700 cm -1, Lignin Signalintensität und Lokalisation wurden in situ abgebildet. Unser Ansatz visualisiert Unterschiede in Ligningehalt, Lokalisierung und chemische Zusammensetzung. Vor kurzem haben wir gezeigt, Raman Imaging von Zellwandpolymere in Arabidopsis thaliana mit lateraler Auflösung, die Sub-um ist 18 Jahre. Hier ist diese Methode vorgestellt affording Visualisierung von Lignin in pflanzlichen Zellwänden und den Vergleich der Verholzung in verschiedenen Geweben, Mustern oder Arten ohne Färbung oder die Kennzeichnung der Gewebe.

Watch this Scientific Journal Video about Label-free in situ Imaging of Lignification in Plant Cell Walls at JoVE.com

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

Eine Methode der konfokalen Raman Mikroskopie basiert vorgestellt, bietet label-free-Visualisierung von Lignin in pflanzlichen Zellwänden und den Vergleich der Verholzung in verschiedenen Geweben, Mustern oder Arten.

Label-freie In situ Imaging der Verholzung in pflanzlichen Zellwänden

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

label-free-in-situ-imaging-of-lignification-in-plant-cell-walls

Research

JoVE Journal

Biology

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

12.3K Aufrufe.  University of California, Berkeley. Eine Methode der konfokalen Raman Mikroskopie basiert vorgestellt, bietet label-free-Visualisierung von Lignin in pflanzlichen Zellwänden und den Vergleich der Verholzung in verschiedenen Geweben, Mustern oder Arten.

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

In situ Imaging of the Mouse Thymus Using 2-Photon Microscopy

Two-photon Microscopy (TPM) enables us to image deep into the thymus and document the events that are important for thymocyte development. To follow the migration of individuals in a crowd of thymocytes , we generate neonatal chimeras where less than one percent of the thymocytes are derived from a donor that is transgenic for a ubiquitously express fluorescent protein. To generate these partial hematopoetic chimeras, neonatal recipients are injected with bone marrow between 3-7 days of age. After 4-6 weeks, the mouse is sacrificed and the thymus is carefully dissected and bissected preserving the architecture of the tissue that will be imaged. The thymus is glued onto a coverslip in preparation for ex vivo imaging by TPM. During imaging the thymus is kept in DMEM without phenol red that is perfused with 95% oxygen and 5% carbon dioxide and warmed to 37&deg;C. Using this approach, we can study the events required for the generation of a diverse T cell repertoire.

We present step-by-step instructions for the generation of neonatal chimeras as well as the dissection and preparation of the thymus for ex vivo imaging by 2-Photon Microscopy.

In situ Imaging der Maus Thymus mit 2-Photonen-Mikroskopie

Two-photon Microscopy (TPM) enables us to image deep into the thymus and document the events that are important for thymocyte development.  To follow the ...

Forschung

Biologie

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.

Umfassende Analyse der Zusammensetzung von pflanzlichen Zellwänden (Lignozellulose) Teil 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 ...

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.

Umfassende Analyse der Zusammensetzung von pflanzlichen Zellwänden (Lignozellulose) Teil II: Kohlenhydrate

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Imaging Zellgröße Veränderung der Lebens- Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Glycan Profiling of Plant Cell Wall Polymers using Microarrays

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the raw materials for human societies (e.g. wood, paper, textile and biofuel industries)1,2. However, understanding the biosynthesis and function of these components remains challenging.
Cell wall glycans are chemically and conformationally diverse due to the complexity of their building blocks, the glycosyl residues. These form linkages at multiple positions and differ in ring structure, isomeric or anomeric configuration, and in addition, are substituted with an array of non-sugar residues. Glycan composition varies in different cell and/or tissue types or even sub-domains of a single cell wall3. Furthermore, their composition is also modified during development1, or in response to environmental cues4.
In excess of 2,000 genes have Plant cell walls are complex matrixes of heterogeneous glycans been predicted to be involved in cell wall glycan biosynthesis and modification in Arabidopsis5. However, relatively few of the biosynthetic genes have been functionally characterized 4,5. Reverse genetics approaches are difficult because the genes are often differentially expressed, often at low levels, between cell types6. Also, mutant studies are often hindered by gene redundancy or compensatory mechanisms to ensure appropriate cell wall function is maintained7. Thus novel approaches are needed to rapidly characterise the diverse range of glycan structures and to facilitate functional genomics approaches to understanding cell wall biosynthesis and modification.
Monoclonal antibodies (mAbs)8,9 have emerged as an important tool for determining glycan structure and distribution in plants. These recognise distinct epitopes present within major classes of plant cell wall glycans, including pectins, xyloglucans, xylans, mannans, glucans and arabinogalactans. Recently their use has been extended to large-scale screening experiments to determine the relative abundance of glycans in a broad range of plant and tissue types simultaneously9,10,11.
Here we present a microarray-based glycan screening method called Comprehensive Microarray Polymer Profiling (CoMPP) (Figures 1 &amp; 2)10,11 that enables multiple samples (100 sec) to be screened using a miniaturised microarray platform with reduced reagent and sample volumes. The spot signals on the microarray can be formally quantified to give semi-quantitative data about glycan epitope occurrence. This approach is well suited to tracking glycan changes in complex biological systems12 and providing a global overview of cell wall composition particularly when prior knowledge of this is unavailable.

A technique called Comprehensive Microarray Polymer Profiling (CoMPP) for the characterisation of plant cell wall glycans is described. This method combines the specificity of monoclonal antibodies directed to defined glycan-epitopes with a miniature microarray analytical platform allowing screening of glycan occurrence in a broad range of biological contexts.

Glycan Profiling of Plant Cell Wall Polymers mit Microarrays

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

Live-Imaging von apoptotischen Ausverkauf während<em&gt; Drosophila</em&gt; Embryogenese

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix containing polysaccharides that include cellulose microfibrils that form both crystalline structures and cellulose chains of amorphous organization. The orientation of the cellulose fibers and their concentrations dictate the mechanical properties of the cell. Several methods are used to determine the levels of crystalline cellulose, each bringing both advantages and limitations. Some can distinguish the proportion of crystalline regions within the total cellulose. However, they are limited to whole-organ analyses that are deficient in spatiotemporal information. Others relying on live imaging, are limited by the use of imprecise dyes. Here, we report a sensitive polarized light-based system for specific quantification of relative light retardance, representing crystalline cellulose accumulation in cross sections of Arabidopsis thaliana roots. In this method, the cellular resolution and anatomical data are maintained, enabling direct comparisons between the different tissues composing the growing root. This approach opens a new analytical dimension, shedding light on the link between cell wall composition, cellular behavior and whole-organ growth.

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

Hohe Auflösung Quantifizierung kristalliner Cellulose Akkumulation in<em&gt; Arabidopsis</em&gt; Roots Gewebespezifische Zellwand Änderungen überwachen

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix ...

Hybrid-Cut: An Improved Sectioning Method for Recalcitrant Plant Tissue Samples

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some plant cells have rigid cell walls, tough fibers and crystals (calcium oxalate, silica, etc.), and high water content that often disrupt tissue integrity during plant tissue sectioning.
This study establishes a simple Hybrid-Cut tissue sectioning method. This protocol modifies a paraffin-based sectioning technique and improves the integrity of tissue sections from different plants. Plant tissues were embedded in paraffin before sectioning in a cryostat at -16 &#176;C. Sectioning under low temperature hardened the paraffin blocks, reduced tearing and scratching, and improved tissue integrity significantly. This protocol was successfully applied to calcium oxalate-rich Phalaenopsis orchid tissues as well as recalcitrant tissues such as reproductive organs and leaves of rice, maize, and wheat. In addition, the high quality of tissue sections from Hybrid-Cut could be used in combination with in situ hybridization (ISH) to provide spatial expression patterns of genes of interest. In conclusion, this protocol is particularly useful for recalcitrant plant tissue containing high crystal or silica content. Good quality tissue sections enable morphological and other biological studies.

This protocol describes a simple Hybrid-Cut tissue sectioning method that is useful for recalcitrant plant tissues. Good quality tissue sections enable anatomical studies and other biological studies including in situ hybridization (ISH).

Hybrid-Cut: Ein verbesserter Schnittverfahren für widerspenstige Pflanzengewebeproben

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some ...

Identification of Novel Regulators of Plant Transpiration by Large-Scale Thermal Imaging Screening in Helianthus Annuus

Plant adaptation to biotic and abiotic stresses is governed by a variety of factors, among which the regulation of stomatal aperture in response to water deficit or pathogens plays a crucial role. Identifying small molecules that regulate stomatal movement can therefore contribute to understanding the physiological basis by which plants adapt to their environment. Large-scale screening approaches that have been used to identify regulators of stomatal movement have potential limitations: some rely heavily on the abscisic acid (ABA) hormone signaling pathway, therefore excluding ABA-independent mechanisms, while others rely on the observation of indirect, long-term physiological effects such as plant growth and development. The screening method presented here allows the large-scale treatment of plants with a library of chemicals coupled with a direct quantification of their transpiration by thermal imaging. Since evaporation of water through transpiration results in leaf surface cooling, thermal imaging provides a non-invasive approach to investigate changes in stomatal conductance over time. In this protocol, Helianthus annuus seedlings are grown hydroponically and then treated by root feeding, in which the primary root is cut and dipped into the chemical being tested. Thermal imaging followed by statistical analysis of cotyledonary temperature changes over time allows for the identification of bioactive molecules modulating stomatal aperture. Our proof-of-concept experiments demonstrate that a chemical can be carried from the cut root to the cotyledon of the sunflower seedling within 10 minutes. In addition, when plants are treated with ABA as a positive control, an increase in leaf surface temperature can be detected within minutes. Our method thus allows the efficient and rapid identification of novel molecules regulating stomatal aperture.

Identification of Novel Regulators of Plant Transpiration by Large-Scale Thermal Imaging Screening in Helianthus Annuus

We provide a method for identifying modulators of foliar transpiration by large-scale screening of a compound library.

Identifizierung neuartiger Regulatoren der Pflanzentranspiration durch großangelegtes Thermisches Bildgebungsscreening in Helianthus Annuus

Plant adaptation to biotic and abiotic stresses is governed by a variety of factors, among which the regulation of stomatal aperture in response to water ...

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits obesity and many associated diseases including cardiovascular disorders, type II diabetes, and cancer. To advance the current understanding of lipid metabolic regulation, quantitative methods to precisely measure in vivo lipid storage levels in time and space have become increasingly important and useful. Traditional approaches to analyze lipid storage are semi-quantitative for microscopic assessment or lacking spatio-temporal information for biochemical measurement. Stimulated Raman scattering (SRS) microscopy is a label-free chemical imaging technology that enables rapid and quantitative detection of lipids in live cells with a subcellular resolution. As the contrast is exploited from intrinsic molecular vibrations, SRS microscopy also permits four-dimensional tracking of lipids in live animals. In the last decade, SRS microscopy has been widely used for small molecule imaging in biomedical research and overcome the major limitations of conventional fluorescent staining and lipid extraction methods. In the laboratory, we have combined SRS microscopy with the genetic and biochemical tools available to the powerful model organism, Caenorhabditis elegans, to investigate the distribution and heterogeneity of lipid droplets across different cells and tissues and ultimately to discover novel conserved signaling pathways that modulate lipid metabolism. Here, we present the working principles and the detailed setup of the SRS microscope and provide methods for its use in quantifying lipid storage at distinct developmental timepoints of wild-type and insulin signaling deficient mutant C. elegans.

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows selective, label-free imaging of specific chemical moieties and it has been effectively employed to image lipid molecules in vivo. Here, we provide a brief introduction to the principle of SRS microscopy and describe methods for its use in imaging lipid storage in Caenorhabditis elegans.

Markierungsfreie Bildgebung der Lipidspeicherdynamik in Caenorhabditis elegans mit stimulierter Raman-Streumikroskopie

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits ...