We thank Dr. M. Rosenberg for her advice and help with anatomical sectioning. We also thank D. Eisler for his technical assistance. This research was supported by grants from FP7-PEOPLE-IRG-2008, Binational Agriculture research and Development (BARD; IS-4246-09), and Israel Science Foundation (ISF; 592/13).

1. Crescimento de Plantas <ol><li> Superfície esterilizar as sementes. <ol><li> Esterilizar sementes (até 30 mg) por via húmida ou seca. Como um exemplo, para o método de esterilização a seco, adicionar 1 ml de HCl glacial a 37% em 50 ml de lixívia, num copo de vidro em um exsicador fechado durante pelo menos 4 horas e até durante a noite. Execute esta etapa em uma capa química. </li></ol></li><li> Espalhe as sementes estéreis em placas com agar vegetal 0,8% em 0,5x Murashige e Skoog (MS). Enrole placas com Parafilm ou fita porosa e cobrir com folha de alumínio. </li><li> Armazenar as placas a 4 ° C durante 1 - 4 dias para promover a germinação uniforme. </li><li> Transferir as placas para uma câmara de crescimento adequado para o cultivo de plântulas de Arabidopsis. Colocar as placas verticalmente para manter o crescimento da raiz na parte superior do meio de agar e para evitar a penetração. </li><li> mudas de transferência para fixador 7 dias após a germinação (DAG). </li></ol> 2. Fixação <ol><li> Preparar 50 ml de Richardson Solution através da combinação de volumes iguais de 1% bórax (agente de buffer), 1% de azul de metileno e 1% Azure (corantes histológicos) em água destilada duas vezes. Filtrar através de uma seringa Impulsionada 0,22 PVDF Filtro Unidade antes do uso. </li><li> Prepare 50 ml de fixador: 1,25% de glutaraldeído em 0,05 M de sódio cacodilato (agente tampão). Nota: Estes materiais são tóxicos e devem ser manuseados em uma capa química. </li><li> Adicionar 100 uL de solução de Richardson (ver 2.1) para a solução de fixação (ver 2.2) para se obter a solução final fixação. </li><li> Mark cada poço de uma placa de 6 poços com o nome da amostra correspondente. </li><li> Coloque 2 - 3 ml de solução final de fixação (ver 2.3) nas cavidades, de modo que as mudas, uma vez transferidos, serão totalmente cobertos em líquido. </li><li> transferir cuidadosamente, usando uma pinça, até 10 mudas para cada poço. Selar as placas usando Parafilm. Armazenar a 4 ° C, no escuro, durante pelo menos uma noite, e até uma semana. </li></ol> 3. desidratação <ol> &lt;li&gt; Desidratar mudas. Transferi-los entre poços de novas placas de 6 poços, contendo concentrações crescentes de etanol, como listados a seguir: 10% de etanol durante 15 min; Etanol a 30% durante 15 min; Etanol a 50% durante 15 min; Etanol a 70% durante 15 min; 85% de etanol durante 15 min; Etanol a 95% durante 15 min; 95% de etanol a 4 ° C, durante a noite, no mínimo. Lidar com mudas, segurando-os cuidadosamente por seus cotilédones, de preferência com uma pinça de plástico. </li></ol> 4. Infiltração <ol><li> Prepare meio de infiltração. Misture o conteúdo de um saco do ativador kit historesina com 50 ml kit líquido resina básica em uma pequena garrafa de vidro. Mexa com ímã para 20 min. forma de infiltração pode ser mantida a 4 ° C durante algumas semanas. </li><li> Mark cada poço de uma nova placa de 6 poços com o nome da amostra correspondente e encher com 2 - 3 ml de meio de infiltração. </li><li> Transferir as mudas de solução de etanol a 95% ao meio de infiltração com a pinça. Certifique-se de que as mudas são fully coberto pelo meio. </li><li> Armazenar a 4 ° C durante pelo menos 4 dias, para assegurar o máximo de penetração nos tecidos da planta. </li></ol> 5. Preparação do bloco <ol><li> Coloque etiquetas pequenas, lápis-marcados com nome da amostra, dentro de cada poço de incorporação de moldes. </li><li> Prepare a incorporação meio pela adição de 1 ml de endurecedor para 15 ml de meio de infiltração (ver 4.1). Não tente para preparar um volume menor do que 15 mL, para evitar problemas de polimerização. </li><li> Encha metade de cada cavidade no molde com 200 ul de meio de fixação e cobrir com uma peça de película de sobrecarga cortadas com a forma do molde, mas a um tamanho de um milímetro maior em cada lado. Incubar à temperatura ambiente durante pelo menos 2 horas, para permitir que a polimerização. Não exceda 5 horas. </li><li> Preparar um lote adicional de incorporação de forma (ver 5.2) e manter em gelo, para evitar a polimerização durante a pontas de raiz são dispostos no molde. </li><li> Corte cada ponta da raiz sob um escopo de dissecação usando bisturi e posicionar i com muito cuidadot: verticalmente para seções transversais ou horizontalmente para o corte longitudinal, em cerca de 2-3 mm da periferia do molde. Encha o molde completamente com a solução e cobertura de bloqueio com um filme em cima. Note-se que a solução de bloqueamento vai encolher ligeiramente, uma vez que é polimerizada. </li><li> Mantenha a temperatura ambiente durante 2 horas, no mínimo. </li><li> Para secar os blocos, coloque em uma caixa com gel de sílica seca e deixar à temperatura ambiente durante a noite, no mínimo. </li></ol> 6. Seccionamento <ol><li> Prepare facas de vidro de bastões de vidro com uma máquina de faca. Use as facas de vidro para o seccionamento de tecidos. Corte o bastão de vidro em um quadrado e depois cortar o quadrado na diagonal para produzir duas facas, usando uma ferramenta de pontuação diamante. </li><li> O bloqueio em 2 - 3 uM fatias de largura, utilizando uma máquina de corte. Coloque as fatias sobre uma lâmina de vidro, em cima de gotas de água destilada e coloque o slide em um bloco de aquecimento definido para fogo baixo (50 - 60 ° C), até que a água evapora. </li></ol><ol><li> Diluir solução Richardson (ver 2.1) a 0,2% da solução original e utilizar 1 ml de ele para cobrir as fatias. Coloque no bloco de calor durante 5 segundos e depois lave com água destilada. </li><li> Seque o slide em uma placa quente. Observar sob um escopo de dissecação e marcar a parte de trás da lâmina com um marcador, para indicar a posição das fatias de interesse, para facilitar a sua identificação mais tarde sob o microscópio. </li><li> Adicione 100 ml de meio de montagem e cobrir com uma lamela. </li></ol> 8. Aquisição de Imagem <ol><li> Colocar a lâmina coberta contendo as secções de raiz com um microscópio óptico equipado com sistema Polscope adequado para se obter informação retardância e um filtro polarizador / interferência óptica (Figura 1 dC). </li><li> Escolha de ampliação que permite a visualização clara de toda a amostra (por exemplo, 40X neste estudo) e concentrar a microscope em um campo vazio que contém seção do bloco, mas nenhuma raiz seção. Tente encontrar uma região limpa, sem manchas. Utilizar esta região para definir o fundo. </li><li> Defina o sistema de imagem e parâmetros de software: Definir intervalo para 17 nm. Aplicar &quot;Exposição Automática&quot; e obter uma imagem de fundo do campo vazio (ver 8.2). Uma imagem de fundo é usado por experimento. </li></ol> 9. Análise Polscope de Celulose microfibrila Orientação <ol><li> Análise de secções longitudinais. <ol><li> Coloque uma lâmina contendo secções longitudinais sob o microscópio e obter uma imagem de retardamento que. Concentre-se o microscópio em uma seção de raiz de interesse. Fornecer um novo nome de arquivo. </li><li> Capturar uma imagem abriu (imagem com informações retardo) através da janela &quot;Viva abriu&quot;. Certifique-se de capturar uma secção que inclui a parede celular, que pode ser identificado por a celulose corada correndo ao longo da periferia da célula, tal como mostrado na Figura 2C.</li></ol></li><li> análise de imagem. <ol><li> Dê um duplo clique sobre a imagem relevante na pilha de imagens. Selecione &quot;color Orientação Pseudo&quot; na guia &quot;Configurações de vídeo&quot;. </li><li> Para estimar o ângulo de microfibrilas de celulose, combinar com a cor correspondente da parede da célula para que, na roda de cores. NOTA: A roda de cores na Figura 2C reflecte o eixo lento da luz encontra-se ao longo da microfibrilas que indica a direcção ao longo do eixo das microfibrilas. O ângulo de microfibrilas de celulose é o ângulo indicado pela cor e o eixo longitudinal da célula. Por exemplo, a célula de alongamento na inserção mostra a cor ciano, inclinado por cerca de 90 ° em relação ao eixo longitudinal das células. </li><li> Para quantificar o ângulo médio microfibrilas na parede celular, use a ferramenta &quot;retardance e Azimute Stamp&quot; a partir da barra de ferramentas do software. Ao selecionar a ferramenta, aponte e clique sobre a região de interesse na imagem obtida. NOTA: Este will resultado no visor dos ângulos (azimute) em graus (Figura 2C). </li><li> Em um acesso aberto na luz software análise alternativa polarizada, almoço &quot;Pol-Analyzer&quot; plug-in a partir do menu plugin. Na nova janela aberta, selecione a guia &quot;Birefringence&quot; e aberto e, em seguida, salve o arquivo de fundo e o arquivo de imagem. </li><li> Na janela de imagem, selecione &quot;Specs&quot;. e ajustar o &quot;tamanho do pixel ROI &#39;(use 5 pixels para este estudo). No &quot;Specs&quot;. janela, selecione &quot;Mostrar valores na tabela de resultados&quot;. </li><li> Use a ferramenta Polígono para marcar a região de interesse na janela de imagem. Seleccionar &quot;Linhas de orientação&quot; para obter a orientação da microfibrilas de celulose. Nota: Isto irá resultar na exibição dos ângulos (azimute) em graus. NOTA: O ângulo médio calculado por um dos dois métodos deve ser em seguida normalizada para o posicionamento da secção de raiz ao longo da corrediça. Para corrigir desvio dao posicionamento horizontal de toda a secção de raiz ao longo da corrediça, quantificar o ângulo médio de micro fibrilas da parede celular que flanqueia a célula ao longo do seu eixo longitudinal (isto é, perpendicular à secção e ao longo do eixo de crescimento da raiz, a Figura 2C). Este ângulo é de 180 ° quando a seção de raiz é posicionada com precisão ao longo do slide. Para calcular o ângulo de microfibrilas, subtrair o ângulo microfibrila média medida da parede celular de flanco de 180 °. Adicionar a diferença resultou ao ângulo de microfibrilas médio medido da parede da célula que se encontra plana sobre a lâmina. </li></ol></li></ol> 10. Análise de Polscope Celulose cristalina Acumulação <ol><li> Use as seções transversais (transversais). Note que a acumulação de celulose cristalina só pode ser comparada entre as células quando as microfibrilas de celulose, são alinhados em um ângulo semelhante (tal como determinado em 9). </li><li> Dê um duplo clique sobre a imagem relevante na pilha de imagens. Selecione &quot;Pseudo cor &quot;no guia&quot; Configurações de vídeo &quot;. </li><li> Para medir retardance. Zoom-in na imagem. Selecione &quot;Região&quot; e marque todas as regiões de interesse. Mover para a guia &quot;medidas&quot; e &quot;Copiar para área de transferência&quot; todos os valores. <ol><li> Transferir os dados para o Excel e extrair informações necessárias (região de retardance dizer). Normalizar os dados. NOTA: Para explicar as diferenças de espessura entre fatias, expressar os valores relativos a uma borda da parede celular específico no slide. Para normalizar, dividir o retardância à média correspondente à parede da célula de interesse pela retardância média de uma parede celular diferente na mesma imagem. Por exemplo, a parede exterior da célula epidérmica foi normalizada para o interior da parede celular cortical no nosso estudo. </li></ol></li><li> Repita as medições durante pelo menos 3 secções por raiz, com um mínimo de 3 raízes por amostra. </li></ol>

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The authors have no conflicts of interest.

Aqui, apresentamos um método para determinação do acúmulo de celulose cristalina em diferentes tecidos que compõem as raízes de Arabidopsis, mantendo informações anatômicas. Como tal, ela fornece uma etapa adicional para a compreensão de processos de crescimento em plantas com uma resolução celular. Este método pode também ser aplicado para o estudo de espécies de plantas adicionais e órgãos. Um número de pontos devem ser considerados quando se aplica o método. Em...

A parede de célula de planta é uma estrutura dinâmica. células em crescimento são cercadas por uma parede celular primária, cuja organização permite que as células a se expandir. As células que param de crescer depósito de uma parede secundário mais rígido que reforça o suporte mecânico da planta. Ambas as paredes celulares são compostas de microfibrilas de celulose embebidas numa matriz de polissacáridos de estruturas diferentes (por exemplo, hemicelulose e pectina) que variam entre o estágio de desenvolvimento diferentes e tecidos 1,2. A celulose é sintetizada na forma de cadeias de (1,4) -β-D-glucano que são fortemente alinhados para formar a micro fibrilas de uma estrutura cristalina. celulose amorfa refere-se às regiões onde as cadeias de glucano são menos ordenada. A relação entre o cristalino e os domínios amorfos é um parâmetro pensado para afectar as propriedades mecânicas da parede da célula, fornecendo resistência mecânica e característica viscoelástica, respectivamente 3. Vários métodos têm sidodesenvolvidas para detectar e quantificar as duas formas de arranjo de celulose, entre eles de difracção de raios-X e polarização cruzada / ângulo mágico girando RMN de estado sólido 4. Difracção de raios-X pode ser utilizada para determinar a proporção de cristalino contra domínios celulose amorfa na amostra 5. Um método alternativo usa fraccionamento de parede celular conteúdo em material insolúvel em ácido e solúvel em ácido, para distinguir entre a cristalina e a celulose amorfa, ou outros polímeros, respectivamente. Nesta abordagem, a incorporação de glicose marcada ([14 C] glucose) é utilizada para quantificar a 6,7 de celulose. Estes métodos necessitam de grandes volumes de material de planta para análises de órgãos inteiros, na melhor das hipóteses, e, portanto, são inadequadamente sensível à variação específica do tecido na estrutura da parede celular. A visualização das microfibrilas de celulose com uma resolução celular pode ser obtida em estudos de imagem ao vivo combinados com corantes fluorescentes 8,9, que podem identificar mudanças naa orientação das microfibrilas de celulose. No entanto, estes corantes não são utilizados para quantificação, eles não são específicos de celulose cristalina e pode interferir com a estrutura normal da parede celular 8. Polscope é uma técnica de imagiologia que se baseia na capacidade de celulose cristalina para dividir os feixes de luz e retardar a parte da luz 10. retardo de luz é mais forte para as microfibrilas que se situam perpendicularmente à direcção da propagação da luz. Para microfibrilas com orientação semelhante, quanto maior o grau de cristalinidade, quanto maior for o retardamento de luz 11. Assim, polscope é utilizado para estudar ambos os níveis relativos e orientação das microfibrilas de celulose. Raízes exibem crescimento linear, durante o qual as células originárias no nicho de células-tronco, na ponta da raiz, são submetidos a uma série de divisões celulares, antes de se expandir rapidamente 12. As células compreendendo a raiz expandir numa unidireccional (anisotrópica)forma, como ditado por pequenos hormônios molécula de sinalização que impactam as propriedades da parede celular 13. Respostas diferenciadas aos hormônios, no tempo e no espaço, fornecer um meio de garantir o crescimento de órgãos equilibrada 14. Assim, a análise de alta resolução da estrutura da parede celular podem fornecer informações importantes necessários para melhor compreender a relação entre as respostas específicas do tipo de células para crescimento de órgão inteiro. Aqui, nós relatamos a implementação de polscope para estudar a acumulação específica de tecido de celulose cristalina em raízes de Arabidopsis, como observado em seções anatômicas alta qualidade. Este método recentemente descoberto de acumulação específico do tipo de célula de celulose cristalina, em resposta à perturbação espacial da actividade hormonal 15.

<table><tbody><tr><td>Murashige &amp;amp; Skoog (MS)</td><td>Duchefa</td><td>M0221</td><td></td></tr><tr><td>Borax</td><td>LOBA CHEMIE&amp;nbsp;</td><td>6038</td><td></td></tr><tr><td>Methylene blue</td><td>Sigma</td><td>M-9140</td><td></td></tr><tr><td>Azure</td><td>Sigma</td><td>861065</td><td></td></tr><tr><td>Syringe Driven 0.22 mm PVDF Filter&amp;nbsp;</td><td>MILLEX-GV</td><td></td><td></td></tr><tr><td>Glutaraldehyde</td><td>EMS</td><td>16220</td><td></td></tr><tr><td>Sodium Cacodylate</td><td>Sigma</td><td>C-0250</td><td></td></tr><tr><td>Leica kit historesin&amp;nbsp;</td><td>Leica</td><td>7022 18 500</td><td></td></tr><tr><td>embedding molds</td><td>Agar Scientific</td><td>AGG3530</td><td></td></tr><tr><td>film 100 &amp;micro; P.P.C.&amp;nbsp;</td><td>www.Jolybar.com</td><td></td><td>overhead film</td></tr><tr><td>Knivemaker</td><td>LKB</td><td>7800</td><td></td></tr><tr><td>Ultratome III</td><td>LKB</td><td>8800</td><td>Ultratome</td></tr><tr><td>Shandon immumount&amp;nbsp;</td><td>Thermo</td><td>9990402</td><td>mounting medium</td></tr><tr><td>light microscope</td><td>Nikon</td><td>Eclipse 80i</td><td></td></tr><tr><td>Abrio imaging system</td><td>CRI</td><td>Abrio imaging system</td><td></td></tr><tr><td>Abrio V2.2 software</td><td>CRI</td><td>Abrio V2.2 software</td><td></td></tr><tr><td>Open access polarized light image analysis software &amp;nbsp;</td><td>OPS</td><td>OpenPolScope</td><td>&lt;a href=&quot;http://www.openpolscope.org/&quot;&gt;http://www.openpolscope.org/&lt;/a&gt;</td></tr></tbody></table>

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.

Nós estudamos os efeitos de respostas específicas do tipo de células para brassinosteróides (BRS), usando a raiz de Arabidopsis como um modelo de órgão 15-17. Quando o receptor de BRI1 BR é orientada, no fundo do mutante BRI1, para um subconjunto de células epidérmicas, chamadas de células não-capilares (Figura 2A, B), que inibe a expansão das células unidireccional de células vizinhas, e todo o crescimento da raiz 15.</sup...

Watch this Scientific Journal Video about High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications at JoVE.com

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.

Quantificação de alta resolução Celulose cristalina Acumulação em<em&gt; Arabidopsis</em&gt; Raízes para monitorar celular específico do tecido da parede Modificações

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 ...

high-resolution-quantification-crystalline-cellulose-accumulation

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

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

8.1K visualizações.  Technion-Israel Institute of Technology. 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.

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

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.

Análise da composição completa de paredes celulares das plantas (biomassa lignocelulósica) Parte II: Os hidratos de carbono

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 ...

Biologia

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

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

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

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

Label-free In situ Imaging de lignificação na parede celular das plantas

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

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through nanoindentations with an atomic force microscope (AFM) coupled with an optical inverted fluorescence microscope. The method consists of applying controlled forces to the sample while measuring its deformation, allowing quantifying parameters such as the apparent Young's modulus of cell walls at subcellular resolutions. It requires a careful mechanical immobilization of the sample and correct selection of indenters and indentation depths. Although it can be used only in external tissues, this method allows characterizing mechanical changes in plant cell walls during development and enables the correlation of these microscopic changes with the growth of an entire organ.

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

The atomic force microscopy indentation protocol offers the possibility to dissect the role of the physical properties of the cell wall of a particular cell of a tissue or organ during normal or constrained growth (i.e., under water deficit).

Microscopia de Força Atômica para Estudo das Propriedades Físicas de Células Epidérmicas de Raízes Vivas de Arabidopsis

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through ...

Biologia do Desenvolvimento

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Longo prazo, de alta resolução de imagem Lapse Confocal Tempo de<em&gt; Cotilédone Arabidopsis</em&gt; Epiderme durante a germinação

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Using Flatbed Scanners to Collect High-resolution Time-lapsed Images of the Arabidopsis Root Gravitropic Response

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories can face is the development and attainment of these methods. Observation of phenotypes in a process of interest is a typical objective of research labs studying gene function and this is often achieved through image capture. A particular process that is amenable to observation using imaging approaches is the corrective growth of a seedling root that has been displaced from alignment with the gravity vector. Imaging platforms used to measure the root gravitropic response can be expensive, relatively low in throughput, and/or labor intensive. These issues have been addressed by developing a high-throughput image capture method using inexpensive, yet high-resolution, flatbed scanners. Using this method, images can be captured every few minutes at 4,800 dpi. The current setup enables collection of 216 individual responses per day. The image data collected is of ample quality for image analysis applications.

This protocol describes a process for rapid collection of images of Arabidopsis seedlings responding to a gravity stimulus using commercially-available flatbed scanners. The method allows for inexpensive, high-volume capture of high-resolution images amenable for downstream analysis algorithms.

Usando scanners de mesa para coletar imagens de alta resolução tempo decorrido da Arabidopsis Root gravitrópico Response

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories ...

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) micro/nano-indentations, for a JPK&#160;AFM. Specifically, in this protocol we measure the apparent Young&#8217;s modulus of cell walls at subcellular resolutions across regions of up to 100 &#181;m&#160;x 100 &#181;m in floral meristems, hypocotyls, and roots. This requires careful preparation of the sample, the correct selection of micro-indenters and indentation depths. To account for cell wall properties only, measurements are performed in highly concentrated solutions of mannitol in order to plasmolyze the cells and thus remove the contribution of cell turgor pressure.
In contrast to other extant techniques, by using different indenters and indentation depths, this method allows simultaneous multiscale measurements, i.e. at subcellular resolutions and across hundreds of cells comprising a tissue. This means that it is now possible&#160;to spatially-temporally characterize the changes that take place in the mechanical properties of cell walls during development, enabling these changes to be correlated with growth and differentiation. This represents a key step to understand how coordinated microscopic cellular changes bring about macroscopic morphogenetic events.
However, several limitations remain: the method can only be used on fairly small samples (around 100 &#181;m in diameter) and only on external tissues; the method is sensitive to tissue topography; it measures only certain aspects of the tissue&#8217;s complex mechanical properties. The technique is being developed rapidly and it is likely that most of these limitations will be resolved in the near future.

We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

Mapeamento baseado em AFM das propriedades elásticas de paredes celulares: em tecido, os celulares, e as Resoluções Subcelulares

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) ...

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

Arabidopsis thaliana is a model organism commonly used to understand and manipulate various cellular processes in plants, and it has been used extensively in the study of secondary cell wall formation. Secondary cell wall deposition occurs after the primary cell wall is laid down, a process carried out exclusively by specialized cells such as those forming vessel and fiber tissues. Most secondary cell walls are composed of cellulose (40&#8211;50%), hemicellulose (25&#8211;30%), and lignin (20&#8211;30%). Several mutations affecting secondary cell wall biosynthesis have been isolated, and the corresponding mutants may or may not exhibit obvious biochemical composition changes or visual phenotypes since these mutations could be masked by compensatory responses. Staining procedures have historically been used to show differences on a cellular basis. These methods are exclusively visual means of analysis; nevertheless their role in rapid and critical analysis is of great importance. Congo red and calcofluor white are stains used to detect polysaccharides, whereas M&#228;ule and phloroglucinol are commonly used to determine differences in lignin, and toluidine blue O is used to differentially stain polysaccharides and lignin. The seemingly simple techniques of sectioning, staining, and imaging can be a challenge for beginners. Starting with sample preparation using the A. thaliana model, this study details the protocols of a variety of staining methodologies that can be easily implemented for observation of cell and tissue organization in secondary cell walls of plants.

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

Plant cell wall composition varies between tissue types and can include lignin, cellulose, hemicelluloses, and pectin. Various staining techniques have been developed to visualize differences at the cell-type level. This paper is a compilation of commonly used cell wall staining techniques.

Coloração histoquímica de<em&gt; Arabidopsis thaliana</em&gt; Celular Secundária parede Elements

Arabidopsis thaliana is a model organism commonly used to understand and manipulate various cellular processes in plants, and it has been used extensively ...

An Efficient Method for Quantitative, Single-cell Analysis of Chromatin Modification and Nuclear Architecture in Whole-mount Ovules in Arabidopsis

In flowering plants, the somatic-to-reproductive cell fate transition is marked by the specification of spore mother cells (SMCs) in floral organs of the adult plant. The female SMC (megaspore mother cell, MMC) differentiates in the ovule primordium and undergoes meiosis. The selected haploid megaspore then undergoes mitosis to form the multicellular female gametophyte, which will give rise to the gametes, the egg cell and central cell, together with accessory cells. The limited accessibility of the MMC, meiocyte and female gametophyte inside the ovule is technically challenging for cytological and cytogenetic analyses at single cell level. Particularly, direct or indirect immunodetection of cellular or nuclear epitopes is impaired by poor penetration of the reagents inside the plant cell and single-cell imaging is demised by the lack of optical clarity in whole-mount tissues.
Thus, we developed an efficient method to analyze the nuclear organization and chromatin modification at high resolution of single cell in whole-mount embedded Arabidopsis ovules. It is based on dissection and embedding of fixed ovules in a thin layer of acrylamide gel on a microscopic slide. The embedded ovules are subjected to chemical and enzymatic treatments aiming at improving tissue clarity and permeability to the immunostaining reagents. Those treatments preserve cellular and chromatin organization, DNA and protein epitopes. The samples can be used for different downstream cytological analyses, including chromatin immunostaining, fluorescence in situ hybridization (FISH), and DNA staining for heterochromatin analysis. Confocal laser scanning microscopy (CLSM) imaging, with high resolution, followed by 3D reconstruction allows for quantitative measurements at single-cell resolution.

An Efficient Method for Quantitative, Single-cell Analysis of Chromatin Modification and Nuclear Architecture in Whole-mount Ovules in Arabidopsis

We provide here an efficient and reliable protocol for immunostaining, Fluorescence in&#160;situ Hybridization, DNA staining followed by quantitative, high-resolution imaging in whole-mount Arabidopsis thaliana ovules. This method was successfully used to analyze chromatin modifications and nuclear architecture.

Um método eficiente para Quantitativa, Análise de células Único de cromatina Modificação e Arquitetura Nuclear em Todo-mount óvulos em<em&gt; Arabidopsis</em

In flowering plants, the somatic-to-reproductive cell fate transition is marked by the specification of spore mother cells (SMCs) in floral organs of the ...

Wide-Field, Real-Time Imaging of Local and Systemic Wound Signals in Arabidopsis

Plants respond to mechanical stresses such as wounding and herbivory by inducing defense responses both in the damaged and in the distal undamaged parts. Upon wounding of a leaf, an increase in cytosolic calcium ion concentration (Ca2+ signal) occurs at the wound site. This signal is rapidly transmitted to undamaged leaves, where defense responses are activated. Our recent research revealed that glutamate leaking from the wounded cells of the leaf into the apoplast around them serves as a wound signal. This glutamate activates glutamate receptor-like Ca2+ permeable channels, which then leads to long-distance Ca2+ signal propagation throughout the plant. The spatial and temporal characteristics of these events can be captured with real-time imaging of living plants expressing genetically encoded fluorescent biosensors. Here we introduce a plant-wide, real-time imaging method to monitor the dynamics of both the Ca2+ signals and changes in apoplastic glutamate that occur in response to wounding. This approach uses a wide-field fluorescence microscope and transgenic Arabidopsis plants expressing Green Fluorescent Protein (GFP)-based Ca2+ and glutamate biosensors. In addition, we present methodology to easily elicit wound-induced, glutamate-triggered rapid and long-distance Ca2+ signal propagation. This protocol can also be applied to studies on other plant stresses to help investigate how plant systemic signaling might be involved in their signaling and response networks.

Wide-Field, Real-Time Imaging of Local and Systemic Wound Signals in Arabidopsis

Extracellular glutamate-triggered systemic calcium signaling is critical for the induction of plant defense responses to mechanical wounding and herbivore attack in plants. This article describes a method to visualize the spatial and temporal dynamics of both these factors using Arabidopsis thaliana plants expressing calcium- and glutamate-sensitive fluorescent biosensors.

Imagens em campo amplo e em tempo real de sinais de feridas locais e sistêmicas em Arabidopsis

Plants respond to mechanical stresses such as wounding and herbivory by inducing defense responses both in the damaged and in the distal undamaged parts. ...

Imagem de alta resolução da biossíntese de clorofila em mudas de Arabidopsis

Non-invasive Assay for Chlorophyll Biosynthesis Kinetics Determination during Early Stages of Arabidopsis De-etiolation

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly dynamic process of chlorophyll biosynthesis is triggered during the shift from the dark to the light in flowering plants. At the moment when etiolated seedlings are exposed to the first traces of sunlight, rapid (in order of seconds) conversion of protochlorophyllide into chlorophyllide is mediated by unique light-accepting protein complexes, leading via subsequent metabolic steps to the production of fully functional chlorophyll. Standard techniques for chlorophyll content analysis include pigment extraction from detached plant tissues, which does not apply to studying such fast processes. To investigate chlorophyll kinetics in vivo with high accuracy and spatiotemporal resolution in the first hours after light-induced de-etiolation, an instrument and protocol were developed. Here, we present a detailed procedure designed for statistically robust quantification of chlorophyll in the early stages of Arabidopsis de-etiolation.

Autor em destaque: medição não invasiva de alta resolução da síntese de clorofila durante o desestiolamento

Here, we describe an advanced tool designed for chlorophyll biosynthesis monitoring during the early stages of Arabidopsis seedling de-etiolation. The novel methodology provides non-invasive real-time chlorophyll fluorescence imaging at high spatial and temporal resolution.

Ensaio Não Invasivo para Determinação da Cinética de Biossíntese de Clorofila durante os Estágios Iniciais da Desetiolação de Arabidopsis

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly ...

Quantificação de alta resolução Celulose cristalina Acumulação em

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