Tools for Cellulose Analysis in Plant Cell Walls

Abstract

Cellulose is intimately associated with multiple facets of human civilization: central to clothing, shelter, heat, medicine, and food, there are few moments in the average human life that are not spent in direct contact with cellulose or a by-product of its composition. It should come as no surprise then that a considerable amount of time and energy have been spent, and a couple of Nobel Prizes awarded, in research involved with the analysis of cellulose structure and metabolism from various sources. As the understanding of this important biomacromolecule has developed, numerous analytical techniques have been put to use to decipher cellulose biosynthesis, structure, and function within the plant cell wall. The content of this Update is designed to introduce the reader to current and developing tools for cellulose characterization. To begin, “a brief history” on the analysis of cellulose describes how many of the modern analytical techniques used to determine cellulose structure came into use. This leads into the various imaging techniques that interrogate cellulose biosynthesis, especially those that have arisen since the identification of the CELLULOSE SYNTHASE A (CESA) genes. We then turn our attention to recent in vitro biochemical studies of CESA, and in this context we discuss the relationship between CESA and detergent-resistant fractions of the plasma membrane (PM), which have the opportunity to shine new light on the PM-cell wall continuum. Cellulose was named by the French Academy over 171 years ago (Brongniart et al., 1839), subsequent to its characterization in various plant tissues by the famous French plant scientist Anselme Payen (Payen, 1838). His use of different treatments based on sodium hydroxide, potassium hydroxide, or nitric acid to extract and partially digest cellulose from oak and beech wood revealed an element composition comparable to that of starch (Payen, 1838). Classical organic chemistry then allowed for the determination of the β-(1→4) linkage that separates Glc residues in the cellobiose unit (for review, see Hon, 1994). The remarkable nature of cellulose as a polymer of repeating Glc (cellobiose) units (Staudinger, 1926; Haworth, 1932) contributed greatly to the 1937 and 1953 Nobel Prizes in Chemistry. Today, it is understood that cellulose fibrils from many natural sources result from individual glucan chains of cellulose aggregating via hydrogen bonds and Van der Waals forces to form a long thread-like paracrystalline structure termed the microfibril. The route that led to a sophisticated model of cellulose structure began with x-ray diffraction (XRD) studies. The first XRD patterns of cellulose fibers were generated from wood, hemp, and bamboo samples, and although detailed structural data were not initially obtained, it was determined that the crystallites were of a rod-like shape (Nishikawa and Ono, 1913). However, cellulose in a majority of higher plants forms crystalline domains that are not large enough to produce high-resolution crystallographic structure determination (Lai Kee Him et al., 2002; Müller et al., 2002). Therefore, many of the early molecular models developed for the monoclinic unit cell (Meyer and Misch, 1937) and triclinic unit cell (Honjo and Watanabe, 1958) of cellulose were based on algal or tunicate (animal) model systems (Fischer and Mann, 1960). In addition, modern XRD data have been collected at even higher resolution than before using a synchrotron light source and can be paired with the separate analytical technique of neutron diffraction, which in combination with specific deuteration has greatly increased the power to locate hydrogen atoms involved in the intermolecular or intramolecular hydrogen bonding of the cellulose microfibril (Nishiyama et al., 2002, 2003). Simultaneous to the development of XRD methods, Rowen et al. (1947) analyzed cellulose from cotton (Gossypium hirsutum) by infrared (IR) absorption spectroscopy, and later, Marrinan and Mann (1956) recognized that algae (Valonia) and bacterial cellulose yielded IR spectra that were different from those of tunicates, cotton, or ramie (Boehmeria nivea). Eventually, in the early 1980s, the spectroscopic technique of solid-state 13C cross-polarization magic-angle spinning NMR spectroscopy was able to resolve this issue by showing that native cellulose I diffraction data from many natural sources were a composite of diffraction from the two crystalline allomorphs Iα (triclinic unit cell) and Iβ (monoclinic unit cell; Atalla and VanderHart, 1984). 13C-NMR spectroscopy would not only confirm that the crystalline structure of the cellulose microfibril in most plants was a composite of Iα and Iβ crystalline forms (Viëtor et al., 2002) but, in combination with Fourier transform IR spectroscopy spectra, suggested that the more crystalline inner chains of the microfibril core are composed primarily of cellulose Iβ, while both forms of cellulose compose the chains in the surrounding paracrystalline sheath (SturcovÁ et al., 2004). Collectively, these early crystallographic and spectroscopic studies laid the foundation for elucidation of the native cellulose structure in plant cell walls. Simultaneously, microscopic analyses showed that cellulose microfibrils were dimensionally different in different cell types of the plant (Roelofsen and Houwink, 1953; Balashov and Preston, 1955). Malcolm Brown and coworkers revealed, first in the green alga Oocystis (Brown and Montezinos, 1976) and then in higher plants (Mueller and Brown, 1980), that the ends of nascent cellulose microfibrils were often associated with globular structures designated as terminal complexes embedded in the PM. The freeze fracture of the PM was imaged on the P-fracture face by transmission electron microscopy (TEM), showing a structure with a 6-fold symmetry (rosette), and remains to this day a fundamental piece of evidence suggesting that this type of structure is involved in cellulose synthesis in plants (Mueller and Brown, 1980). Furthermore, in the study by Kimura and coworkers (1999), TEM was used in combination with an immunoaffinity probe to show that the catalytic subunit of cellulose synthase is associated with the rosette complex in vascular plants. Despite the disadvantages of extensive and often destructive sample preparation and the inability to study live specimens, TEM is unsurpassed in its resolution and has been the method of choice for ultrastructural analysis of the cell wall. Newer techniques in TEM sample preparation are being developed to reduce sample destruction along with the use of promising new affinity tools to be used in conjunction with TEM, such as carbohydrate-binding modules, that will most likely provide enhanced molecular resolution imaging of the cellulose microfibril network (for review, see Sarkar et al., 2009). Integrated with the above-mentioned analytical tools of IR, NMR, XRD, and TEM, the relatively new imaging technique of atomic force microscopy (AFM) has the capacity to provide atom-level resolution of the cellulosic matrix in the cell wall of fresh tissue. Therefore, beyond the compositional structure of cellulose, AFM can offer a spatial view of cellulose microfibril orientation in the polylaminate cell wall. AFM is based on scanning probe microscopy (Binnig and Rohrer, 1986) and uses a physical tip to scan the surface of a specimen to determine its topography, physical properties, and chemical structure (Drake et al., 1989). AFM operates by driving a cantilever with a sharp tip mounted at its end to allow faster scanning across the specimen surface. AFM images are the result of convolutions of the tip and the “true” structure of the specimen at an atomic resolution. Plant cell walls were one of the first biological samples that were examined by AFM (Kirby et al., 1996; van der Wel et al., 1996). The motivation of using AFM for characterizing cell walls is obvious: plant cell walls are relatively stiff and flat, and the molecular features of the microfibril network occur at the nanometer scale. Ideally, AFM could be used to answer some of the key questions regarding the nanostructures within plant cell wall cellulose (Kirby et al., 1996; van der Wel et al., 1996; Engel et al., 1999; Morris et al., 1999; Thimm et al., 2000; Davies and Harris, 2003) beyond that defined by the previously mentioned analytical techniques. To take full advantage of AFM and reduce interference by artifacts, an approach with minimized sample preparation is ideal. Greatest success has used hand-cut sections of fresh or naturally aged dry tissue while operating the AFM device in tapping mode and imaging the inner surface of the native cell wall (Ding and Himmel, 2006; Himmel et al., 2007). Using this strategy, it was possible to image primary and secondary cell walls from maize (Zea mays). AFM imaging of dry primary cell walls documented microfibril dimensions precisely measured at 3 to 5 nm (Ding and Himmel, 2006), consistent with the 36-glucan chain model of cellulose elementary fibril (CEF) biosynthesis. In addition, cellulose macrofibrils, consisting of a bundle of CEFs that split at the end to form smaller bundles and eventually single CEFs, were also observed by AFM (Ding and Himmel, 2006). Each microfibril observed in mature primary cell walls contained only a single CEF with noncellulosic polymers associated with its surface (Ding and Himmel, 2006; Himmel et al., 2007). AFM images of maize cell walls from fresh cells further confirmed these observations (Fig. 1, A and B). AFM images of 3-d-old developing maize coleoptiles (Fig. 1A) showed macrofibrils of 50 to 100 nm in diameter with rather clean surface features, which contained multiple CEFs. By contrast, 4-week old maize stem piths containing mature parenchyma (Fig. 1B) displayed two distinguishable layers. In the upper layer, the fibrils appeared to be small macrofibrils with diameters of 7 to 10 nm and single CEFs with diameters of 3 to 5 nm. In the posterior layer, all microfibrils of 3 to 5 nm diameter were arranged in a parallel orientation (Fig. 1B). Furthermore, in the mature cell walls (Fig. 1B), particle-like features decorated the macrofibrils that may have been formed by hemicelluloses, proteins, or other cell wall polymers (Ding and Himmel, 2006) and were not observed in the macrofibrils of elongating cells (Fig. 1A). A plausible explanation for these differences among cell types is a delay between cellulose biosynthesis and the incorporation of other cell wall polymers into the cell wall. Ultimately, AFM imaging representing the native structure of cellulose provides an enormous opportunity to better understand the molecular architecture in dermal cell layers, particularly when combined with the confocal live cell imaging described below. Emerging tools for cellulose analysis in plant cell walls. The background is set with the model plant Arabidopsis; a simple schematic is overlaid on the stem to indicate various facets of cell wall cellulose synthesis. A, AFM height image of fibril structures in the elongating primary cell wall of maize contained primarily macrofibrils that split at the end (indicated by the arrow). B, The mature primary cell wall contained both small macrofibrils in the upper-most layer that can also split at the end (indicated by the arrow) and parallel-arranged single elementary fibrils. Particle-like features were clearly seen and were typically associated with the macrofibrils (indicated by the circle). Bar in A = 100 nm. C, Confocal image of upper hypocotyl cells of transgenic plants expressing YFP::CESA6, red fluorescent protein (RFP)::TUA5 (tubulin), and the overlay of the two compatible fluorophors in a merged imaged reveals the coincidence of CESA with microtubules. Bar = 5 μm. D to F, Electron micrographs of cellulose microfibrils extracted from primary walls of blackberry cells (D and E) and of cellulose synthesized in vitro (F) using detergent extracts as a source of enzyme. The cellulose from cell walls was observed before (D) and after (E) treatment with the Updegraff reagent (Lai Kee Him et al., 2002). A cellulose microfibril synthesized in vitro associated to a globular particle, which most likely corresponds to the enzyme complex (not treated with the Updegraff reagent; F). G, Electron micrograph of cellulose microfibrils synthesized in vitro by DRMs isolated from hybrid aspen cells grown as suspension cultures (negative staining with 2% uranyl acetate). The inset shows the magnification of putative DRM structures that carry the active cellulose synthase machinery and from which cellulose microfibrils are being formed. Bars in E to G = 0.1 μm. As an analytical tool, confocal microscopy has rapidly evolved from being a challenging technique with limited accessibility to a high-throughput tool providing quantitatively precise localization data for 30% of the proteome in the model plant Arabidopsis (Arabidopsis thaliana; Chalfie et al., 1994; Cutler et al., 2000; Tian et al., 2004; Heazlewood et al., 2007). The capacity to visualize a protein relies on excitation of an autofluorescent protein (AFP), such as the GFP derived from the jellyfish Aqueora victoriae, fused to a plant protein of interest. Despite the most famous AFP being the GFP characterized 16 years ago (Chalfie et al., 1994), there are now numerous forms of AFP, including blue, cyan, and yellow (YFP) fluorescent proteins (Goodin et al., 2007). Designing an AFP expression fusion can still be a daunting task, given the numerous expression vectors available, the choice of where to fuse the AFP, what promoter to use, and whether to use stable or transient expression of the chimeric protein fusion (for review, see Goodin et al., 2007). Live cell imaging of CESA has led to a remarkable increase in our understanding of the and in cells primary cell wall synthesis et al., 2006; et al., et al., et al., et al., et al., and in vascular tissue secondary cell wall synthesis and of the primary wall complex with or in Arabidopsis hypocotyl cells revealed at the of the PM. The that these along at of nm et al., 2006) that The majority of CESA, in the in by with a and in a of smaller et al., 2006; et al., et al., 2009). In of cellulose synthase was initially used to the between CESA complexes and the et al., 2006), first described years ago by and It was long that the CESA complexes and cellulose the for this was not confocal microscopy showed that CESA were with that as molecular rather than as to CESA (Fig. et al., 2006). the was to in the of CESA complexes to the PM et al., et al., 2009). of individual that complexes are to that are with et al., 2009). small have been to with the et al., et al., 2009). these observations that may CESA to the PM by with in the has also revealed that CESA is with secondary cell wall synthesis and these also that are in of CESA, which is for cell wall and The is also for CESA primary cell wall of the the of CESA at the PM et al., 2009). have been from of for it was observed that after at the PM the began to after et al., 2006). a would be that the CESA complex is and cellulose after at the PM. also are observed to from the suggesting a further of that CESA However, it has not been possible to a CESA from the of its to for this is that the of the cell being imaged it to that a has not of the rather than from the PM. many questions for the and will in confocal resolution. have been in of cellulose biosynthesis by live cell the of was in that the CESA and were of et al., 2006; et al., As mentioned using showed the for in CESA and et al., 2009). the localization of CESA after treatment with of cellulose biosynthesis such as et al., and 1980), et al., and et al., 2006) has allowed for the study of the of for these and of CESA from the of CESA is or CESA is to at the PM et al., 2006; et al., et al., 2009). By contrast, not complexes from at the CESA and CESA et al., has been to to a protein in hybrid aspen further an of the on the of CESA et al., of cellulose biosynthesis have been and using live cell a being et al., which both and CESA, further the between these the protein of all these small an enormous opportunity to new or in cell wall cellulose biosynthesis. Despite all the evidence the biochemical analysis of the CESA has been a in the of plant cell wall biosynthesis. The enzyme complex is and this has limited the of it and its biochemical in of proteins from plant typically to the of the of cellulose synthase 1999; and In addition, further is to the of synthase from the PM. This enzyme uses the as cellulose synthase and in vitro et al., Kee Him et al., 2002; et al., even is a cell wall that forms at the cell to cell or in cells or cell structures and the of glucan by synthase is typically in in vitro and this has further the of the and cellulose synthase In addition, to the incorporation of Glc from into has been associated with cellulose biosynthesis (for review, see et al., 2004; 2007). It has been that and cellulose be by the enzyme the of different the catalytic of and cellulose is with this (for review, see and However, one may still that evidence that the of the CESA the of and is still The answer to this will be when an active cellulose synthase preparation of synthase The first in vitro synthesis of cellulose from plant extracts was using cotton and et al., 1996; and Brown, However, it was only years that the amount of cellulose synthesized in vitro was by the of that allow the of enzyme complexes in an form (Lai Kee Him et al., 2002). The use of and to extract cellulose synthase from of cell suspension cultures of blackberry allowed the synthesis of cellulose in This possible the and characterization of the cellulose synthesized in vitro (Lai Kee Him et al., 2002). the cellulose formed in the in vitro was to more crystalline than the cellulose extracted from the primary walls of the as measured by XRD analysis (Lai Kee Him et al., 2002). This was further by the that cellulose from primary walls was to the Updegraff reagent 1, D and E) while the in microfibrils were not (Lai Kee Him et al., 2002). the samples were not treated with this the individual microfibrils synthesized in vitro were associated with globular structures that most likely to the enzyme complex for (Fig. this the method has been in other plant such as hybrid aspen et al., and cells et al., for which was as the It that the type of detergent used for enzyme to be determined for different plant which different As for the blackberry cellulose synthase (Lai Kee Him et al., higher of were with the cellulose synthase from hybrid aspen when the cells were at with to cellulose and synthesized in vitro et al., 2004). To this the of cellulose to in in vitro synthesis Despite these important it remains that the in vitro to be combined with analyses of the in vitro to determine the of β-(1→4) formed. This can be using specific that are not by which is not the for the biochemical evidence of the synthesis of cellulose is by the of the polymer synthesized in the of to the specific more characterization can be for by analysis after of the electron XRD TEM, and NMR (for review, see 2007). However, for XRD and NMR the of the in vitro is to enough polymer for the characterization. NMR is particularly in of the amount of at 10 a method based on the use of in which the Glc is in 13C has been developed and the analysis of in vitro with as as 100 of et al., 2004). The analysis of the cellulose synthesized in vitro can be further by the that is with cellulose in crystalline cellulose such as that synthesized in vitro by the enzyme from blackberry is not in (Lai Kee Him et al., 2002). However, it is important to in that some crystalline may be synthesized and by in In the tools to cellulose synthase in vitro and the cellulose formed are This a opportunity to detailed biochemical analysis of the cellulose synthase complex when a preparation can be To this has been important has been in this with the of complexes from Arabidopsis using and the specific et al., 2009). However, it was not possible in this to for the was not The and tools mentioned for the characterization of cellulose synthesized in vitro will be most also for the biochemical analysis of individual catalytic of cellulose which may be possible in the with the development of more expression systems for to the of these tools and of specific synthase it was that and cellulose are in detergent-resistant structures biochemical as in cells et al., 2009). The were active in vitro and able to that were as and cellulose et al., 2009). The glucan sample as cellulose, in was not treated with to This allowed for the of the detergent-resistant membrane which are as globular structures or of globular structures (Fig. It was not whether both and cellulose are in the or different of DRMs et al., 2009). The relationship between DRMs and is et al., 2006), the used for DRM may the of such structures et al., it remains that the of DRMs of specific of membrane proteins to various et al., the of DRMs is a tool for understanding the of and cellulose with the that with and that of a higher of and than the et al., 2009). In addition, DRMs a form of isolated that can be further using or other such as that between specific and This combined with the and the detailed analysis of the after treatment with a promising to proteins that with the enzyme We for to it was not possible to all of the that have contributed to and we for