Plant Physiology

Minorsky, Peter V.

doi: 10.1104/pp.113.900470pmid: N/A

Root Adaptation to Salinity Requires a Type of Annexin Salinity (NaCl) stress impairs plant growth and inflicts severe crop losses. In roots, increasing extracellular NaCl causes Ca2+ influx and an increase in cytosolic free Ca2+ ([Ca2+]cyt). Manipulation of membrane voltage by varying external concentrations of K+ and Ca2+ has indicated that both hyperpolarization- and depolarization-activated plasma membrane Ca2+-permeable channels can operate in generating the NaCl-induced increase in [Ca2+]cyt. The genetic identities of the Ca2+-permeable channels involved in generating the [Ca2+]cyt signal are unknown. Annexin 1 (AtANN1) is a potential candidate. Plant annexins have been shown to form Ca2+-permeable channels in planar lipid bilayers. These soluble proteins are capable of membrane binding and insertion. The most abundant annexin in Arabidopsis (Arabidopsis thaliana; AtANN1) can exist as a plasma membrane protein and is responsible for the root epidermal plasma membrane Ca2+- and K+-permeable conductance that is activated by extracellular hydroxyl radicals, the most reactive of the reactive oxygen species. Laohavisit et al. (pp. 253–262) have examined the involvement of AtANN1 in the generation of root and root epidermal NaCl-induced [Ca2+]cyt elevation. Luminescent detection of [Ca2+]cyt showed that AtANN1 responds to high extracellular NaCl by mediating reactive oxygen species-activated Ca2+ influx across the plasma membrane of root epidermal protoplasts. Electrophysiological analysis revealed that root epidermal plasma membrane Ca2+ influx currents activated by NaCl are absent from the Atann1 loss-of function-mutant. Adaptive signaling and salt-responsive production of secondary roots are also impaired in the loss of function mutant, thus identifying AtANN1 as a key component of root cell adaptation to NaCl. Root Plasticity under Nutrient Deficiency The environmental regulation of root system architecture (RSA) allows plants to display a high level of root plasticity. Nutrient availability can exert a profound impact on RSA by altering the number, length, angle, and diameter of roots and root hairs. Plants can respond to heterogeneities in the availability of soil resources by allocating roots where the most favorable conditions are found. For example, when grown under limited phosphorus availability, roots exhibit a shallower architecture that results from the inhibition of primary root elongation and the concomitant increase in lateral root formation. Although such changes often determine the nutrient efficiency or stress tolerance of plants, a comprehensive and comparative analysis of root morphological responses to different nutrient deficiencies has not yet been conducted. Gruber et al. (pp. 161–179) raised Arabidopsis plants on agar plates at four levels of deficiency for 12 nutrients and quantified seven root traits. In combination with measurements of biomass and elemental concentrations, they observed that the nutritional status and type of nutrient determined the extent and type of changes in RSA. This systematic comparison of RSA responses to nutrient deficiencies provides a comprehensive view on the overall changes in root plasticity induced by the deficiency of single nutrients and provides a solid basis for the identification of nutrient-sensitive steps in the root developmental program. The advances presented may eventually find some applicability in the high-throughput screening of RSA mutants. SUPPRESSOR OF MAX2 1 Controls Seed Germination Strigolactones (SLs) and karrikins (KARs) comprise two classes of butenolide signaling molecules that stimulate seed germination. Both classes of chemicals serve as chemical signals that activate postfire germination in the soil seed bank, and both control plant growth through a shared MAX2-dependent pathway. An SL biosynthetic pathway and candidate KAR/SL receptors have been characterized, but signaling downstream of MAX2 is poorly defined. A screen for genetic suppressors of the enhanced seed dormancy phenotype of max2 in Arabidopsis led to the identification of the suppressor of max2 1 (smax1) mutant. The smax1 mutation restores the seed germination and seedling photomorphogenesis phenotypes of max2 but does not affect the lateral root formation, axillary shoot growth, or senescence phenotypes of max2.Stanga et al. (pp. 318–330) conclude that SMAX1 is an important component of KAR/SL signaling during seed germination and seedling growth but is not necessary for all MAX2-dependent responses. They hypothesize that one or more SMXL proteins may also act downstream of MAX2 to control the diverse developmental responses to KAR and SL. Nuclear Trapping Controls the Position-Dependent Localization of CAPRICE Positional information is a major factor establishing differential gene expression during multicellular development. Moreover, some of the regulated genes themselves, in turn, encode additional positional signals that further refine the gene expression patterns, via cell-cell communication events. Given the importance of positional information, there is great interest in defining the molecular nature and action of the molecules that mediate cell-cell communication during development. The Arabidopsis root epidermis provides an excellent model system to study the molecular basis of cell fate determination and differentiation. A lateral inhibition mechanism mediated by an R3 single-repeat MYB protein, CAPRICE (CPC), has been proposed to explain the specification of the two types of root epidermal cells (hair [H] cells and nonhair [N] cells). However, it is not clear how CPC acts preferentially in the H-position cells, rather than the N-position cells, where its gene is expressed. To explore this mystery, Kang et al. (pp. 193–204) examined the effect of misexpressed CPC on cell fate specification and CPC localization in the root epidermis. They show that CPC is able to move readily within the root epidermis when its expression level is high and that CPC can induce the hair cell fate in a cell-autonomous manner. When misexpressed in different tissues in the root, CPC was able to induce the hair cell fate in the root epidermis, indicating long-distance movement of CPC. It appears that CPC is capable of moving from the stele tissue in the center of the root to the outermost epidermal layer, where it can induce the hair cell fate. In addition, the CPC protein accumulates primarily in the nuclei of H-position cells in the meristematic region, and this localization requires the H-cell-expressed ENHANCER OF GLABRA3 (EGL3) bHLH transcription factor. These results suggest that cell-cell movement of CPC occurs readily within the meristematic region of the root and that EGL3 preferentially traps the CPC protein in the H-position cells of the epidermis. Distribution and Speciation of Toxic Selenium in Hydrated Roots Selenium (Se) is an essential micronutrient for humans and other animals. At elevated concentrations, however, it is toxic, and the concentration range from deficiency to lethality is unusually narrow. Plants represent a direct entrance to the wider food chain as the main sources of dietary Se. The uptake and accumulation of Se by plants is an important process in controlling health risks resulting from Se deficiency or toxicity. Selenium toxicity to plants has been observed in arid and semiarid soils derived from seleniferous rocks and shales, although anthropogenic contamination is also of concern. It is, therefore, important that the mechanisms of Se uptake, transformation, and toxicity in plants, are understood to reduce health risks in animal and Se toxicity in plants. By means of synchrotron-based x-ray absorption spectroscopy and x-ray fluorescence microscopy, Wang et al. (pp. 407–418) have quantified the longitudinal and radial distribution of Se in its different forms in hydrated roots of cowpea (Vigna unguiculata) exposed to either selenite or selenate. Selenate was found to be 9-fold more toxic to the roots than selenite, most likely because of increased accumulation of organoselenium (e.g. seleno-Met) in selenate-treated roots. Se concentrations in the bulk root tissue were approximately 18-fold higher in the selenate treatment. Although the proportion of Se converted to organic forms was higher for selenite (100%) than for selenate (26%), the absolute concentration of organoselenium was actually approximately 5-fold higher for selenate-treated roots. The uptake of selenate (probably via sulfate transporters) occurred at a much higher rate than for selenite. In addition, the longitudinal and radial distribution of Se in roots differed markedly; the highest tissue concentrations were in the endodermis and cortex approximately ≥4 mm behind the apex when exposed to selenate but in the meristem (approximately 1 mm from the apex) when exposed to selenite. The examination of the distribution and speciation of Se in hydrated roots provide valuable data in understanding the Se uptake, transport, and toxicity. Chromatin Remodeling in Gibberellin Signaling SWI/SNF-type chromatin remodeling complexes (CRCs) are evolutionary conserved in eukaryotes. They consist of a central Snf2-type ATPase in association with several core subunits that correspond to orthologous proteins in yeast. In mammals, the core noncatalytic subunits of SWI/SNF-type complexes, such as SWI3, directly interact with nuclear hormone receptors and coactivators. All known core subunits of SWI/SNF complexes are conserved in plants. CRCs are involved in regulating transcription, DNA replication and repair, and the cell cycle. Mutations of conserved subunits of plant CRCs severely impair growth and development. Sarnowska et al. (pp. 305–317) show that inactivating SWI3C, the core component of Arabidopsis SWI/SNF CRCs, interferes with normal functioning of several plant hormone pathways and alters transcriptional regulation of key genes of gibberellin (GA) biosynthesis. The resulting reduction of GA4 causes severe inhibition of hypocotyl and root elongation, which can be rescued by exogenous GA treatment. Such developmental abnormalities are characteristic of Arabidopsis mutants impaired in GA biosynthesis and signaling. The swi3c mutation markedly decreases the levels of bioactive GA derivatives by causing pathway-wide alteration in the transcription of genes involved in the biosynthesis and inactivation of GAs. Furthermore, the swi3c mutation also down-regulates the expression of GID1 GA-receptor genes, which may affect the GA perception in leaves. Moreover, SWI3C physically interacts in the nucleus with several DELLA proteins, and with SPY, which appears to act upstream of SWI3C in the GA response pathway. Physical interactions of SWI3C with DELLAs and SPY suggest that the function of SWI3C-containing SWI/SNF CRCs may be required for some of the DELLA-mediated effects, such as the activation of GID1 and GA3ox genes involved in GA perception and biosynthesis, respectively. Author notes www.plantphysiol.org/cgi/doi/10.1104/pp.113.900470 © 2013 American Society of Plant Biologists. All Rights Reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Blatt, Mike; Martin, Cathie

doi: 10.1104/pp.113.900471pmid: N/A

The past few years have seen a small number of celebrated cases of scientific fraud that have found their way into the general media. Many more examples of inappropriate data handling come across the editorial desks of virtually every scientific journal. These have focused editors’ attention on inappropriate data handling and fraudulent image manipulation. The Plant Cell and Plant Physiology are no exceptions. Two decades ago, the practicalities of image handling meant that the boundaries were well-defined between what was acceptable and what was not; the darkroom skills needed posed a significant technical barrier to inappropriate manipulation of image data, particularly manipulation done without the intention to deceive but simply to “clean up” the image. The ethical boundaries are as clear-cut today as they were a quarter century ago, but many of the technical barriers to inappropriate manipulation have all but disappeared with the advent of digital image acquisition, storage, and handling. Adobe Photoshop was introduced in 1990 for Macintosh and in 1992 for PCs; its widespread application, and the broader acceptance of digital formats during this past decade, have simplified greatly the tasks of image preparation. They also mean that much less skill is needed to manipulate images. Indeed, a common problem arising from digital formats is that many scientists inadvertently manipulate their image data, often in ways that result in the loss of important information, to make their data look as good as possible. The Journal of Cell Biology carried out a detailed study over the past decade and, commendably, has shared this information publicly. The study found that 10% of articles accepted for publication included inappropriate manipulations of image data that contravened journal policy, even if they did not alter the conclusions drawn from the data (International Society of Managing and Technical Editors, 2013). A surprisingly large number of the authors appeared unaware that they had handled image data inappropriately and, in many cases, were not conscious of the ethical issues and consequences of their actions. As editors, how do we maintain ethical standards in publishing? And, as scientists, how do we educate our students and support our peers to understand what is (and what is not) acceptable practice when handling image data? It is essential to recognize that digital images are data, in fact arrays of numerical data, and must be treated as such. As scientists, we assume that images will not have been altered in any way that affects the visual impression; the quantitative and qualitative relationships within images (data arrays) must be maintained. If these relationships are altered, then such alterations must be fully documented and explained. There are two defining principles behind these expectations: (1) We expect honesty and transparency in scientific reporting, and (2) We expect the scientist, as author, to understand the consequences of processing image data to ensure that any transformations are quantitatively rigorous and comply with ethical standards. There are a few simple rules to follow in meeting these expectations (Rossner and Yamada, 2004; North, 2006; Cromey, 2010): Raw image data must be saved and archived intact and without alteration as part of good laboratory practice. Processing of digital images should be done on a copy of the image data file, not on the original. Retaining raw image data is important because they serve as the standard against which the final image can be compared, and they ensure a route for recovery should a mistake be made during processing. We recommend that image data are saved in TIF format. JPEG compression affects the resolution of the image, and information is lost in the process of conversion. Simple adjustments, applied uniformly, to the entire image are generally acceptable. Changes to brightness, contrast, and color balance fall into this category because they affect the image in a linear fashion. However, it is not acceptable to adjust brightness or contrast levels to such an extent that image data are truncated or lost (giving a white or black background; see Fig. 1B). Such changes may give a clearer picture of bands which are “of interest” in a gel, but they will mask background, including information that is important for quantification and validation. We will not accept image data that are processed in this way. Cropping and resizing an image is usually acceptable, but both may be construed as inappropriate manipulation, on occasion. If cropping, ask whether your motivation is to improve the composition of the image or to hide something that complicates interpretation. The former reason is acceptable; the latter is not. Digital filtering of an image is not encouraged because it can easily mask important information. Most filters use mathematical functions that are nonlinear. There are circumstances in which digital filtering is a necessary part of the experimental methodology. If so, filter processing must be clearly justified and documented in the figure legend or under “Materials and Methods.” Such documentation should include reference to the software version, specification of the filters, and any special settings that were used. Combining images is acceptable only if it is clear to the reader that the images are from separate sources. It is acceptable to combine the images of two similar gels or two parts of the same gel in one figure, but only if a visible gap is left between the images or the images are separated and each surrounded by a box. It is not acceptable to splice two gel images together so that they appear to be adjacent tracks from a single gel. Selective alteration or processing of one region of an image is not acceptable. Such manipulations include “cloning” or copying objects or sections within or between images, “smudging,” blurring, blending, and other manipulations that are applied locally within an image. Common examples (see Fig. 1A) involve sections of an image that have been cloned or blended, to clean up a dirty preparation or to mask an unwanted blemish. Such manipulations constitute inappropriate handling at best and are unethical. If the data require such processing, repeat the experiment. When comparing digital images, it is important that they be acquired under identical conditions, and any postacquisition image processing must be applied identically. If the background or color balance must be adjusted between images within a group, this must be acknowledged in the figure legend or under “Materials and Methods” (see Fig. 1B). Quantitative analysis of images should always be carried out on uniformly processed image data, and the data should be calibrated to a known standard. Most instruments, including fluorescent microscopes, are prone to fluctuations and drift over time, so it is advisable to include appropriate internal standards as checks against such changes. Image data should be documented both with representative images as well as with quantitative statistical analysis of sufficient numbers of experiments. It should be self-evident that experiments that include image data should be repeated and the data analyzed for significance. We expect conclusions drawn from image data to be justified based on their quantitative assessment, not on anecdotal observations. Figure 1. Open in new tabDownload slide Examples of inappropriate image manipulation. A, The gel has been cleaned up to hide a stronger band above the main band at 80 kD in the rightmost lane. Adjusting the exposure and gamma correction in the magnified view (top right) highlights a pattern of pixel “smearing,” indicated by the red arrow, that differs from the pixel pattern elsewhere in the gel image. B, Green fluorescent protein expression in the protoplasts appears roughly equivalent with little signal detectable in the control (left). Adjusting the exposure and contrast to the maximum across the image set (bottom), however, demonstrates that the images have not been processed identically. The first image is completely black, and the color balance between the second and third clearly differs when comparing the backgrounds. Figure 1. Open in new tabDownload slide Examples of inappropriate image manipulation. A, The gel has been cleaned up to hide a stronger band above the main band at 80 kD in the rightmost lane. Adjusting the exposure and gamma correction in the magnified view (top right) highlights a pattern of pixel “smearing,” indicated by the red arrow, that differs from the pixel pattern elsewhere in the gel image. B, Green fluorescent protein expression in the protoplasts appears roughly equivalent with little signal detectable in the control (left). Adjusting the exposure and contrast to the maximum across the image set (bottom), however, demonstrates that the images have not been processed identically. The first image is completely black, and the color balance between the second and third clearly differs when comparing the backgrounds. As editors, we have a responsibility to the readers and authors of Plant Physiology and The Plant Cell to ensure that what we publish is sound scientifically and meets the highest ethical standards. We can help authors become aware of data mishandling and the ethical consequences of inappropriate manipulations and address the probable 10% of articles falling into the category of data handling that is simply misguided or ethically ignorant. Most inappropriate data handling is relatively easy to spot and is often flagged by reviewers. From an editorial and educational standpoint, it is always best to identify and deal with such instances before an article is accepted. To this end, Plant Physiology and The Plant Cell will now have available the facility to analyze cases of suspect mishandling using the forensic tools used by The Rockefeller Press journals, including The Journal of Cell Biology. We are confident that these tools will give our editors the resources they need to handle problems of inappropriate data handling as and when questions arise. We hope, too, that our approach to these issues will help strengthen the scientific community and the reliability of the data we publish. LITERATURE CITED Cromey DW ( 2010 ) Avoiding twisted pixels: ethical guidelines for the appropriate use and manipulation of scientific digital images . Sci Eng Ethics 16 : 639 – 667 Google Scholar Crossref Search ADS PubMed WorldCat International Society of Managing and Technical Editors (2013) Image manipulation in scientific publishing: interview with Liz Williams, PhD. http://www.ismte.org/Interview_with_Liz_Williams Image_Manipulation_in_Scientific_Publishing_Interview_with_Liz_Williams_PhD. Accessed August 16, 2013 North AJ ( 2006 ) Seeing is believing? A beginners’ guide to practical pitfalls in image acquisition . J Cell Biol 172 : 9 – 18 Google Scholar Crossref Search ADS PubMed WorldCat Rossner M Yamada KM ( 2004 ) What’s in a picture? The temptation of image manipulation . J Cell Biol 166 : 11 – 15 Google Scholar Crossref Search ADS PubMed WorldCat Author notes www.plantphysiol.org/cgi/doi/10.1104/pp.113.900471 © 2013 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2013. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Yeats, Trevor H.; Rose, Jocelyn K.C.

doi: 10.1104/pp.113.222737pmid: 23893170

Abstract The plant cuticle is an extracellular hydrophobic layer that covers the aerial epidermis of all land plants, providing protection against desiccation and external environmental stresses. The past decade has seen considerable progress in assembling models for the biosynthesis of its two major components, the polymer cutin and cuticular waxes. Most recently, two breakthroughs in the long-sought molecular bases of alkane formation and polyester synthesis have allowed construction of nearly complete biosynthetic pathways for both waxes and cutin. Concurrently, a complex regulatory network controlling the synthesis of the cuticle is emerging. It has also become clear that the physiological role of the cuticle extends well beyond its primary function as a transpiration barrier, playing important roles in processes ranging from development to interaction with microbes. Here, we review recent progress in the biochemistry and molecular biology of cuticle synthesis and function and highlight some of the major questions that will drive future research in this field. The first plant colonizers of land, approximately 450 million years ago in the mid-Paleozoic era, faced a daunting set of challenges associated with their new terrestrial environment, including desiccation, temperature extremes, gravity, and increased exposure to UV radiation (Waters, 2003; Leliaert et al., 2011). The transition from an exclusively aquatic to a terrestrial life style, therefore, would have necessitated the evolution of a toolbox of morphological and physiological features, some of which are apparent through studies of the fossil record or by examining extant plant lineages. For example, the development of architecturally complex cell walls for biomechanical support and structural protection, which typify modern land plants, can be traced back to divergence and radiation within the Charophycean green algae, their immediate ancestors (Sørensen et al., 2011). However, the most critical adaptive trait for survival during terrestrialization would have been the ability to retain water in increasingly dehydrating habitats. Consequently, the capacity to synthesize, deposit, and maintain a hydrophobic surface layer, or cuticle, over the surfaces of aerial organs was arguably one of the most important innovations in the history of plant evolution. This idea is borne out by both fossil evidence (Edwards, 1993) and the ubiquity of cuticles among all extant embryophytes, from bryophytes (Budke et al., 2012) to angiosperms. Armed with a protective skin, together with a range of adaptive strategies for acquiring and conserving water, as well as for avoiding or tolerating water stress, embryophytes now thrive in a wide range of desiccating environments (Ogburn and Edwards, 2010; Aroca et al., 2012; Delaux et al., 2012; Jones and Dolan, 2012; Obata and Fernie, 2012; Gaff and Oliver, 2013). Accordingly, cuticles from a broad range of species, and in various ecological and agricultural contexts, have been studied from the perspective of their role as the primary barrier to transpirational water loss. However, it is now clear that cuticles play numerous other roles in plant development, physiology, and interactions with the abiotic environment and other organisms. Indeed, in recent years, there have been many instances of unexpected associations between the cuticle and diverse aspects of plant biology. In parallel, the past decade has seen considerable progress in understanding the biosynthesis of the major cuticle components and the complex regulatory networks that control cuticle synthesis and assembly. This review summarizes recent progress in elucidating the biochemistry and molecular biology of cuticle synthesis and function and highlights some of the connections to other aspects of plant biology, including signaling, pathogen defense, and development. Given the broad scope and space limitation, not every aspect of cuticle biosynthesis is covered in depth, and recent specialized reviews focusing on cuticle biomechanical properties (Domínguez et al., 2011), defensive functions (Reina-Pinto and Yephremov, 2009), and transport barrier properties (Burghardt and Riederer, 2006) may be of further interest. In addition, key ongoing questions in the field are discussed, and potential future approaches to resolving those questions are suggested. CUTICLE STRUCTURE, BIOSYNTHESIS, AND ASSEMBLY Plant cuticles are composite structures, composed of a covalently linked macromolecular scaffold of cutin and a variety of organic solvent-soluble lipids that are collectively termed waxes. Although the cuticle is usually considered independently from the underlying polysaccharide cell wall of the epidermis, the two structures are physically associated and have some overlapping functions. Indeed, the cuticle can be considered a specialized lipidic modification of the cell wall, just as lignification is a common modification of plant secondary cell walls. The microscopic structure of the cuticle is often divided into two domains based on histochemical staining and their presumed chemical composition: a cutin-rich domain with embedded polysaccharides, which is referred to as the “cuticular layer,” and an overlying layer that is less abundant in polysaccharides but enriched in waxes, referred to as the “cuticle proper” (Fig. 1A). The waxes are either deposited within the cutin matrix (intracuticular wax) or accumulate on its surface as epicuticular wax crystals, or films. These epicuticular waxes can confer distinct macroscopic surface properties: epicuticular films are responsible for the glossy appearance common to many leaves and fruits, while epicuticular wax crystals account for the dull, glaucous appearance of broccoli (Brassica oleracea) leaves and Arabidopsis (Arabidopsis thaliana) stems. Cuticle architectural organization can be discerned using a number of microscopic techniques. Scanning electron microscopy can reveal the elaborate and diverse morphologies of epicuticular wax crystals (Fig. 1B; Jeffree, 2006), while transmission electron microscopy shows the distinct patterning of interior layers of the cuticle, although this approach does not allow the visualization of wax structures (Fig. 1C). Cuticles vary considerably in their architecture and, depending on species and ontogeny, differ dramatically in hickness, from the nanometer to the micrometer scale (Jeffree, 2006). In the latter case, light microscopy can be used to elucidate the fine structures of the cuticle and epidermal cell wall (Fig. 1D), while histochemical staining coupled with confocal microscopy can further resolve three-dimensional cuticle architecture (Buda et al., 2009). Figure 1. Open in new tabDownload slide Plant cuticle structure. A, Schematic diagram highlighting the major structural features of the cuticle and underlying epidermal cell layer (not drawn to scale). B, Scanning electron micrograph image of an Arabidopsis leaf epidermis and overlying cuticle, seen in cross section. Bar = 5 μm. (Image courtesy of Dr. Lacey Samuels.) C, Transmission electron micrograph image of an Arabidopsis stem epidermal cell wall and cuticle. Bar = 500 nm. (Image courtesy of Dr. Christiane Nawrath.) D, Light microscopy image showing the cuticle of a mature green-stage tomato fruit stained with Sudan Red and the polysaccharide cell walls stained with Alcian Blue. Bar = 50 μm. Figure 1. Open in new tabDownload slide Plant cuticle structure. A, Schematic diagram highlighting the major structural features of the cuticle and underlying epidermal cell layer (not drawn to scale). B, Scanning electron micrograph image of an Arabidopsis leaf epidermis and overlying cuticle, seen in cross section. Bar = 5 μm. (Image courtesy of Dr. Lacey Samuels.) C, Transmission electron micrograph image of an Arabidopsis stem epidermal cell wall and cuticle. Bar = 500 nm. (Image courtesy of Dr. Christiane Nawrath.) D, Light microscopy image showing the cuticle of a mature green-stage tomato fruit stained with Sudan Red and the polysaccharide cell walls stained with Alcian Blue. Bar = 50 μm. Wax Biosynthesis Wax composition can vary substantially with species, ontogeny, and environmental growth conditions (Jenks and Ashworth, 1999). In most cases, the majority of compounds comprising the cuticular wax are derived from very-long-chain fatty acids (VLCFAs; C20–C34), including alkanes, aldehydes, primary and secondary alcohols, ketones, and esters (Table I). In some species, various lipophilic secondary metabolites, such as pentacyclic triterpenoids, flavonoids, and tocopherols, can also be substantial components (Jetter et al., 2006). There has been impressive progress in revealing the molecular biology underlying VLCFA-derived wax biosynthesis, and to this end, Arabidopsis has provided an excellent experimental model (Bernard and Joubès, 2013). In addition to its well-known advantages as a genetic system, the presence of stem epicuticular wax crystals, which impart a glaucous appearance in the wild type, has enabled an easy screen for wax-deficient mutants. Such mutants, termed eceriferum (cer; Koornneef et al., 1989), typically exhibit a glossy stem phenotype, and it has primarily been through molecular analyses of these and other wax mutants that an increasingly complete pathway for acyl wax biosynthesis has been established. Major acyl-lipid classes found in cuticular waxes Table I. Major acyl-lipid classes found in cuticular waxes Most classes occur as homologous series with broad distributions of chain lengths, compounds with typical average chain lengths are shown. Open in new tab Table I. Major acyl-lipid classes found in cuticular waxes Most classes occur as homologous series with broad distributions of chain lengths, compounds with typical average chain lengths are shown. Open in new tab Wax biosynthesis begins with de novo C16 or C18 fatty acid biosynthesis in the plastid of epidermal cells (Fig. 2). These long-chain fatty acid compounds are converted to CoA thioesters by a long-chain acyl-coenzyme A synthase (LACS) isozyme and are ultimately transferred to the endoplasmic reticulum (ER). The mechanism of intracellular trafficking of fatty acid from the chloroplast to the ER remains unknown, although heterologous expression of Arabidopsis LACS1, LACS2, and LACS3 facilitates fatty acid uptake in yeast, suggesting that this class of enzymes may play dual roles in fatty acid trafficking and activation (Pulsifer et al., 2012). For reference, Table II provides a list of the corresponding genes, as well as others discussed in this review. The C16 acyl-CoA is then a substrate for the fatty acid elongase (FAE) complex. Through successive addition of two carbons per cycle derived from malonyl-CoA, the ultimate products of this complex are VLCFAs. The complex consists of four core subunits: β-ketoacyl-CoA synthase, β-ketoacyl-CoA reductase, β-hydroxyacyl-CoA dehydratase, and enoyl-CoA reductase. In Arabidopsis, 21 genes are predicted to encode β-ketoacyl-CoA synthase, and for wax biosynthesis, the most important gene, based on the mutant phenotype, is CER6 (Fiebig et al., 2000). Genes encoding the remaining subunits of the FAE complex, represented by KCR1, PAS2, and CER10, respectively, are less redundant, and their pleiotropic mutant phenotypes underscore the shared importance of the FAE in generating VLCFA precursors for sphingolipid biosynthesis (Zheng et al., 2005; Bach et al., 2008; Beaudoin et al., 2009). An additional family of proteins, composed of CER2, CER26, and CER26-like, appears to be required for the elongation of fatty acids to lengths greater than 28C (Haslam et al., 2012; Pascal et al., 2013). Curiously, these enzymes have sequence homology to BAHD acyltransferases, but conserved catalytic amino acid residues of this family of enzymes are dispensable for the elongation-promoting activity of CER2 (Haslam et al., 2012). The elongation cycles can be terminated by a thioesterase to form free VLCFAs, or the VLCFA-CoA esters can undergo further modifications. Figure 2. Open in new tabDownload slide Cutin and wax biosynthetic pathways. Genes (blue text) are described in the review. Red text denotes compound classes that are typically observed in cuticular wax mixtures. Figure 2. Open in new tabDownload slide Cutin and wax biosynthetic pathways. Genes (blue text) are described in the review. Red text denotes compound classes that are typically observed in cuticular wax mixtures. Cuticle-associated genes discussed in this review Table II. Cuticle-associated genes discussed in this review Gene Symbol . Gene Name . Species . Locus Identifier . Description . ABCG11 ATP-BINDING CASSETTE G11 Arabidopsis At1G17840 ABC half transporter ABCG13 ATP-BINDING CASSETTE G13 Arabidopsis At1G51460 ABC half transporter ABCG32 ATP-BINDING CASSETTE G32 Arabidopsis At2G26910 ABC full transporter BDG BODYGUARD Arabidopsis At1G64670 α/β-Hydrolase family protein BDG3 BODYGUARD3 Arabidopsis At4G24140 Homolog of BDG CD1 CUTIN DEFICIENT1 Tomato Solyc11G006250 Cutin synthase/hydroxyacylglycerol transesterase CD2 CUTIN DEFICIENT2 Tomato Solyc01G091630 Homeodomain-Leu zipper IV transcription factor CER1 ECERIFERUM1 Arabidopsis At1G02205 Involved in alkane formation CER10 ECERIFERUM10 Arabidopsis At3G55360 Enoyl-CoA reductase CER2 ECERIFERUM2 Arabidopsis At4G24510 Required for C28 to C30 elongation of fatty acids CER26 ECERIFERUM26 Arabidopsis At4G13840 Homolog of CER2, required for elongation of fatty acids greater than C30 CER26-like ECERIFERUM26-like Arabidopsis At3G23840 Homolog of CER2 and CER26 CER3 ECERIFERUM3 Arabidopsis At5G57800 Involved in alkane formation CER4 ECERIFERUM1 Arabidopsis At4G33790 VLCFA-CoA by fatty acyl-CoA CER5/ABCG12 ECERIFERUM5/ATP-BINDING CASSETTE G12 Arabidopsis At1G51500 ABC half transporter CER6 ECERIFERUM6 Arabidopsis At1G68530 β-Ketoacyl-CoA synthase CER7 ECERIFERUM7 Arabidopsis At3G60500 Exosomal exoribonuclease CER9 ECERIFERUM9 Arabidopsis At4G34100 Putative E3 ubiquitin ligase CFL1 CURLY FLAG LEAF1 Rice Os02G31140 WW domain-containing protein CYP77A6 CYP77A6 Arabidopsis At3G10570 CYP77A subfamily of cytochrome P450 CYP86A4 CYP86A4 Arabidopsis At1G01600 CYP86A subfamily of cytochrome P450 DCR DEFECTIVE IN CUTICULAR RIDGES Arabidopsis At5G23940 BAHD acyltransferase FDH FIDDLEHEAD Arabidopsis At2G26250 β-Ketoacyl-CoA synthase GPAT6 GLYCEROL-3-PHOSPHATE SN-2-ACYLTRANSFERASE6 Arabidopsis At2G38110 Bifunctional glycerol-3-phosphate sn-2-acyltransferase/phosphatase HDG1 HOMEODOMAIN GLABROUS1 Arabidopsis At3G61150 Homeodomain-Leu zipper IV transcription factor HTH HOTHEAD Arabidopsis At1G72970 Glc-methanol-choline oxidoreductase family protein IRG1 INHIBITOR OF RUST GERM TUBE DIFFERENTIATION1 M. truncatula Medtr5G014400 C2H2 zinc finger transcription factor KCR1 β-KETOACYL-COENZYME A REDUCTASE1 Arabidopsis At1G67730 β-Ketoacyl-CoA reductase LACS1 LONG-CHAIN ACYL-COENZYME A SYNTHASE1 Arabidopsis At2G47240 Long-chain acyl-CoA synthase LACS2 LONG-CHAIN ACYL-COENZYME A SYNTHASE2 Arabidopsis At1G49430 Long-chain acyl-CoA synthase LACS3 LONG-CHAIN ACYL-COENZYME A SYNTHASE3 Arabidopsis At1G64400 Long-chain acyl-CoA synthase LCR LACERATA Arabidopsis At2G45970 CYP86A subfamily of cytochrome P450 LTL1 LI-TOLERANT LIPASE1 Arabidopsis At3G04290 Homolog of CD1 LTPG1 GPI-ANCHORED LIPID TRANSFER PROTEIN1 Arabidopsis At1G27950 GPI-anchored lipid transfer protein LTPG2 GPI-ANCHORED LIPID TRANSFER PROTEIN2 Arabidopsis At3G43720 GPI-anchored lipid transfer protein MAH1 MIDCHAIN ALKANE HYDROXYLASE1 Arabidopsis At1G57750 CYP96A subfamily of cytochrome P450 MYB106 Myb DOMAIN PROTEIN106 Arabidopsis At3G01140 Myb transcription factor MYB16 Myb DOMAIN PROTEIN16 Arabidopsis At5G15310 Myb transcription factor MYB30 Myb DOMAIN PROTEIN30 Arabidopsis At3G28910 Myb transcription factor MYB41 Myb DOMAIN PROTEIN41 Arabidopsis At4G28110 Myb transcription factor MYB96 Myb DOMAIN PROTEIN96 Arabidopsis At5G62470 Myb transcription factor OCL1 OUTER CELL LAYER1 Maize GRMZM2G026643 Homeodomain-Leu zipper IV transcription factor PAS2 PASTICCINO2 Arabidopsis At5G10480 β-Hydroxyacyl-CoA dehydratase SHN2 SHINE2 Arabidopsis At5G11190 Homolog of WIN1/SHN1 WIN1/ SHN1 WAX INDUCER1/SHINE1 Arabidopsis At1G15360 AP2 domain-containing transcription factor WSD1 WAX SYNTHASE/ACYL-COENZYME A:DIACYLGLYCEROL ACYLTRANSFERASE1 Arabidopsis At5G37300 Wax synthase/acyl-CoA:diacylglycerol acyltransferase family protein WXP1 WAX PRODUCTION1 M. truncatula Medtr5G062700 AP2 domain-containing transcription factor – – Arabidopsis At5G33370 Homolog of CD1 Gene Symbol . Gene Name . Species . Locus Identifier . Description . ABCG11 ATP-BINDING CASSETTE G11 Arabidopsis At1G17840 ABC half transporter ABCG13 ATP-BINDING CASSETTE G13 Arabidopsis At1G51460 ABC half transporter ABCG32 ATP-BINDING CASSETTE G32 Arabidopsis At2G26910 ABC full transporter BDG BODYGUARD Arabidopsis At1G64670 α/β-Hydrolase family protein BDG3 BODYGUARD3 Arabidopsis At4G24140 Homolog of BDG CD1 CUTIN DEFICIENT1 Tomato Solyc11G006250 Cutin synthase/hydroxyacylglycerol transesterase CD2 CUTIN DEFICIENT2 Tomato Solyc01G091630 Homeodomain-Leu zipper IV transcription factor CER1 ECERIFERUM1 Arabidopsis At1G02205 Involved in alkane formation CER10 ECERIFERUM10 Arabidopsis At3G55360 Enoyl-CoA reductase CER2 ECERIFERUM2 Arabidopsis At4G24510 Required for C28 to C30 elongation of fatty acids CER26 ECERIFERUM26 Arabidopsis At4G13840 Homolog of CER2, required for elongation of fatty acids greater than C30 CER26-like ECERIFERUM26-like Arabidopsis At3G23840 Homolog of CER2 and CER26 CER3 ECERIFERUM3 Arabidopsis At5G57800 Involved in alkane formation CER4 ECERIFERUM1 Arabidopsis At4G33790 VLCFA-CoA by fatty acyl-CoA CER5/ABCG12 ECERIFERUM5/ATP-BINDING CASSETTE G12 Arabidopsis At1G51500 ABC half transporter CER6 ECERIFERUM6 Arabidopsis At1G68530 β-Ketoacyl-CoA synthase CER7 ECERIFERUM7 Arabidopsis At3G60500 Exosomal exoribonuclease CER9 ECERIFERUM9 Arabidopsis At4G34100 Putative E3 ubiquitin ligase CFL1 CURLY FLAG LEAF1 Rice Os02G31140 WW domain-containing protein CYP77A6 CYP77A6 Arabidopsis At3G10570 CYP77A subfamily of cytochrome P450 CYP86A4 CYP86A4 Arabidopsis At1G01600 CYP86A subfamily of cytochrome P450 DCR DEFECTIVE IN CUTICULAR RIDGES Arabidopsis At5G23940 BAHD acyltransferase FDH FIDDLEHEAD Arabidopsis At2G26250 β-Ketoacyl-CoA synthase GPAT6 GLYCEROL-3-PHOSPHATE SN-2-ACYLTRANSFERASE6 Arabidopsis At2G38110 Bifunctional glycerol-3-phosphate sn-2-acyltransferase/phosphatase HDG1 HOMEODOMAIN GLABROUS1 Arabidopsis At3G61150 Homeodomain-Leu zipper IV transcription factor HTH HOTHEAD Arabidopsis At1G72970 Glc-methanol-choline oxidoreductase family protein IRG1 INHIBITOR OF RUST GERM TUBE DIFFERENTIATION1 M. truncatula Medtr5G014400 C2H2 zinc finger transcription factor KCR1 β-KETOACYL-COENZYME A REDUCTASE1 Arabidopsis At1G67730 β-Ketoacyl-CoA reductase LACS1 LONG-CHAIN ACYL-COENZYME A SYNTHASE1 Arabidopsis At2G47240 Long-chain acyl-CoA synthase LACS2 LONG-CHAIN ACYL-COENZYME A SYNTHASE2 Arabidopsis At1G49430 Long-chain acyl-CoA synthase LACS3 LONG-CHAIN ACYL-COENZYME A SYNTHASE3 Arabidopsis At1G64400 Long-chain acyl-CoA synthase LCR LACERATA Arabidopsis At2G45970 CYP86A subfamily of cytochrome P450 LTL1 LI-TOLERANT LIPASE1 Arabidopsis At3G04290 Homolog of CD1 LTPG1 GPI-ANCHORED LIPID TRANSFER PROTEIN1 Arabidopsis At1G27950 GPI-anchored lipid transfer protein LTPG2 GPI-ANCHORED LIPID TRANSFER PROTEIN2 Arabidopsis At3G43720 GPI-anchored lipid transfer protein MAH1 MIDCHAIN ALKANE HYDROXYLASE1 Arabidopsis At1G57750 CYP96A subfamily of cytochrome P450 MYB106 Myb DOMAIN PROTEIN106 Arabidopsis At3G01140 Myb transcription factor MYB16 Myb DOMAIN PROTEIN16 Arabidopsis At5G15310 Myb transcription factor MYB30 Myb DOMAIN PROTEIN30 Arabidopsis At3G28910 Myb transcription factor MYB41 Myb DOMAIN PROTEIN41 Arabidopsis At4G28110 Myb transcription factor MYB96 Myb DOMAIN PROTEIN96 Arabidopsis At5G62470 Myb transcription factor OCL1 OUTER CELL LAYER1 Maize GRMZM2G026643 Homeodomain-Leu zipper IV transcription factor PAS2 PASTICCINO2 Arabidopsis At5G10480 β-Hydroxyacyl-CoA dehydratase SHN2 SHINE2 Arabidopsis At5G11190 Homolog of WIN1/SHN1 WIN1/ SHN1 WAX INDUCER1/SHINE1 Arabidopsis At1G15360 AP2 domain-containing transcription factor WSD1 WAX SYNTHASE/ACYL-COENZYME A:DIACYLGLYCEROL ACYLTRANSFERASE1 Arabidopsis At5G37300 Wax synthase/acyl-CoA:diacylglycerol acyltransferase family protein WXP1 WAX PRODUCTION1 M. truncatula Medtr5G062700 AP2 domain-containing transcription factor – – Arabidopsis At5G33370 Homolog of CD1 Open in new tab Table II. Cuticle-associated genes discussed in this review Gene Symbol . Gene Name . Species . Locus Identifier . Description . ABCG11 ATP-BINDING CASSETTE G11 Arabidopsis At1G17840 ABC half transporter ABCG13 ATP-BINDING CASSETTE G13 Arabidopsis At1G51460 ABC half transporter ABCG32 ATP-BINDING CASSETTE G32 Arabidopsis At2G26910 ABC full transporter BDG BODYGUARD Arabidopsis At1G64670 α/β-Hydrolase family protein BDG3 BODYGUARD3 Arabidopsis At4G24140 Homolog of BDG CD1 CUTIN DEFICIENT1 Tomato Solyc11G006250 Cutin synthase/hydroxyacylglycerol transesterase CD2 CUTIN DEFICIENT2 Tomato Solyc01G091630 Homeodomain-Leu zipper IV transcription factor CER1 ECERIFERUM1 Arabidopsis At1G02205 Involved in alkane formation CER10 ECERIFERUM10 Arabidopsis At3G55360 Enoyl-CoA reductase CER2 ECERIFERUM2 Arabidopsis At4G24510 Required for C28 to C30 elongation of fatty acids CER26 ECERIFERUM26 Arabidopsis At4G13840 Homolog of CER2, required for elongation of fatty acids greater than C30 CER26-like ECERIFERUM26-like Arabidopsis At3G23840 Homolog of CER2 and CER26 CER3 ECERIFERUM3 Arabidopsis At5G57800 Involved in alkane formation CER4 ECERIFERUM1 Arabidopsis At4G33790 VLCFA-CoA by fatty acyl-CoA CER5/ABCG12 ECERIFERUM5/ATP-BINDING CASSETTE G12 Arabidopsis At1G51500 ABC half transporter CER6 ECERIFERUM6 Arabidopsis At1G68530 β-Ketoacyl-CoA synthase CER7 ECERIFERUM7 Arabidopsis At3G60500 Exosomal exoribonuclease CER9 ECERIFERUM9 Arabidopsis At4G34100 Putative E3 ubiquitin ligase CFL1 CURLY FLAG LEAF1 Rice Os02G31140 WW domain-containing protein CYP77A6 CYP77A6 Arabidopsis At3G10570 CYP77A subfamily of cytochrome P450 CYP86A4 CYP86A4 Arabidopsis At1G01600 CYP86A subfamily of cytochrome P450 DCR DEFECTIVE IN CUTICULAR RIDGES Arabidopsis At5G23940 BAHD acyltransferase FDH FIDDLEHEAD Arabidopsis At2G26250 β-Ketoacyl-CoA synthase GPAT6 GLYCEROL-3-PHOSPHATE SN-2-ACYLTRANSFERASE6 Arabidopsis At2G38110 Bifunctional glycerol-3-phosphate sn-2-acyltransferase/phosphatase HDG1 HOMEODOMAIN GLABROUS1 Arabidopsis At3G61150 Homeodomain-Leu zipper IV transcription factor HTH HOTHEAD Arabidopsis At1G72970 Glc-methanol-choline oxidoreductase family protein IRG1 INHIBITOR OF RUST GERM TUBE DIFFERENTIATION1 M. truncatula Medtr5G014400 C2H2 zinc finger transcription factor KCR1 β-KETOACYL-COENZYME A REDUCTASE1 Arabidopsis At1G67730 β-Ketoacyl-CoA reductase LACS1 LONG-CHAIN ACYL-COENZYME A SYNTHASE1 Arabidopsis At2G47240 Long-chain acyl-CoA synthase LACS2 LONG-CHAIN ACYL-COENZYME A SYNTHASE2 Arabidopsis At1G49430 Long-chain acyl-CoA synthase LACS3 LONG-CHAIN ACYL-COENZYME A SYNTHASE3 Arabidopsis At1G64400 Long-chain acyl-CoA synthase LCR LACERATA Arabidopsis At2G45970 CYP86A subfamily of cytochrome P450 LTL1 LI-TOLERANT LIPASE1 Arabidopsis At3G04290 Homolog of CD1 LTPG1 GPI-ANCHORED LIPID TRANSFER PROTEIN1 Arabidopsis At1G27950 GPI-anchored lipid transfer protein LTPG2 GPI-ANCHORED LIPID TRANSFER PROTEIN2 Arabidopsis At3G43720 GPI-anchored lipid transfer protein MAH1 MIDCHAIN ALKANE HYDROXYLASE1 Arabidopsis At1G57750 CYP96A subfamily of cytochrome P450 MYB106 Myb DOMAIN PROTEIN106 Arabidopsis At3G01140 Myb transcription factor MYB16 Myb DOMAIN PROTEIN16 Arabidopsis At5G15310 Myb transcription factor MYB30 Myb DOMAIN PROTEIN30 Arabidopsis At3G28910 Myb transcription factor MYB41 Myb DOMAIN PROTEIN41 Arabidopsis At4G28110 Myb transcription factor MYB96 Myb DOMAIN PROTEIN96 Arabidopsis At5G62470 Myb transcription factor OCL1 OUTER CELL LAYER1 Maize GRMZM2G026643 Homeodomain-Leu zipper IV transcription factor PAS2 PASTICCINO2 Arabidopsis At5G10480 β-Hydroxyacyl-CoA dehydratase SHN2 SHINE2 Arabidopsis At5G11190 Homolog of WIN1/SHN1 WIN1/ SHN1 WAX INDUCER1/SHINE1 Arabidopsis At1G15360 AP2 domain-containing transcription factor WSD1 WAX SYNTHASE/ACYL-COENZYME A:DIACYLGLYCEROL ACYLTRANSFERASE1 Arabidopsis At5G37300 Wax synthase/acyl-CoA:diacylglycerol acyltransferase family protein WXP1 WAX PRODUCTION1 M. truncatula Medtr5G062700 AP2 domain-containing transcription factor – – Arabidopsis At5G33370 Homolog of CD1 Gene Symbol . Gene Name . Species . Locus Identifier . Description . ABCG11 ATP-BINDING CASSETTE G11 Arabidopsis At1G17840 ABC half transporter ABCG13 ATP-BINDING CASSETTE G13 Arabidopsis At1G51460 ABC half transporter ABCG32 ATP-BINDING CASSETTE G32 Arabidopsis At2G26910 ABC full transporter BDG BODYGUARD Arabidopsis At1G64670 α/β-Hydrolase family protein BDG3 BODYGUARD3 Arabidopsis At4G24140 Homolog of BDG CD1 CUTIN DEFICIENT1 Tomato Solyc11G006250 Cutin synthase/hydroxyacylglycerol transesterase CD2 CUTIN DEFICIENT2 Tomato Solyc01G091630 Homeodomain-Leu zipper IV transcription factor CER1 ECERIFERUM1 Arabidopsis At1G02205 Involved in alkane formation CER10 ECERIFERUM10 Arabidopsis At3G55360 Enoyl-CoA reductase CER2 ECERIFERUM2 Arabidopsis At4G24510 Required for C28 to C30 elongation of fatty acids CER26 ECERIFERUM26 Arabidopsis At4G13840 Homolog of CER2, required for elongation of fatty acids greater than C30 CER26-like ECERIFERUM26-like Arabidopsis At3G23840 Homolog of CER2 and CER26 CER3 ECERIFERUM3 Arabidopsis At5G57800 Involved in alkane formation CER4 ECERIFERUM1 Arabidopsis At4G33790 VLCFA-CoA by fatty acyl-CoA CER5/ABCG12 ECERIFERUM5/ATP-BINDING CASSETTE G12 Arabidopsis At1G51500 ABC half transporter CER6 ECERIFERUM6 Arabidopsis At1G68530 β-Ketoacyl-CoA synthase CER7 ECERIFERUM7 Arabidopsis At3G60500 Exosomal exoribonuclease CER9 ECERIFERUM9 Arabidopsis At4G34100 Putative E3 ubiquitin ligase CFL1 CURLY FLAG LEAF1 Rice Os02G31140 WW domain-containing protein CYP77A6 CYP77A6 Arabidopsis At3G10570 CYP77A subfamily of cytochrome P450 CYP86A4 CYP86A4 Arabidopsis At1G01600 CYP86A subfamily of cytochrome P450 DCR DEFECTIVE IN CUTICULAR RIDGES Arabidopsis At5G23940 BAHD acyltransferase FDH FIDDLEHEAD Arabidopsis At2G26250 β-Ketoacyl-CoA synthase GPAT6 GLYCEROL-3-PHOSPHATE SN-2-ACYLTRANSFERASE6 Arabidopsis At2G38110 Bifunctional glycerol-3-phosphate sn-2-acyltransferase/phosphatase HDG1 HOMEODOMAIN GLABROUS1 Arabidopsis At3G61150 Homeodomain-Leu zipper IV transcription factor HTH HOTHEAD Arabidopsis At1G72970 Glc-methanol-choline oxidoreductase family protein IRG1 INHIBITOR OF RUST GERM TUBE DIFFERENTIATION1 M. truncatula Medtr5G014400 C2H2 zinc finger transcription factor KCR1 β-KETOACYL-COENZYME A REDUCTASE1 Arabidopsis At1G67730 β-Ketoacyl-CoA reductase LACS1 LONG-CHAIN ACYL-COENZYME A SYNTHASE1 Arabidopsis At2G47240 Long-chain acyl-CoA synthase LACS2 LONG-CHAIN ACYL-COENZYME A SYNTHASE2 Arabidopsis At1G49430 Long-chain acyl-CoA synthase LACS3 LONG-CHAIN ACYL-COENZYME A SYNTHASE3 Arabidopsis At1G64400 Long-chain acyl-CoA synthase LCR LACERATA Arabidopsis At2G45970 CYP86A subfamily of cytochrome P450 LTL1 LI-TOLERANT LIPASE1 Arabidopsis At3G04290 Homolog of CD1 LTPG1 GPI-ANCHORED LIPID TRANSFER PROTEIN1 Arabidopsis At1G27950 GPI-anchored lipid transfer protein LTPG2 GPI-ANCHORED LIPID TRANSFER PROTEIN2 Arabidopsis At3G43720 GPI-anchored lipid transfer protein MAH1 MIDCHAIN ALKANE HYDROXYLASE1 Arabidopsis At1G57750 CYP96A subfamily of cytochrome P450 MYB106 Myb DOMAIN PROTEIN106 Arabidopsis At3G01140 Myb transcription factor MYB16 Myb DOMAIN PROTEIN16 Arabidopsis At5G15310 Myb transcription factor MYB30 Myb DOMAIN PROTEIN30 Arabidopsis At3G28910 Myb transcription factor MYB41 Myb DOMAIN PROTEIN41 Arabidopsis At4G28110 Myb transcription factor MYB96 Myb DOMAIN PROTEIN96 Arabidopsis At5G62470 Myb transcription factor OCL1 OUTER CELL LAYER1 Maize GRMZM2G026643 Homeodomain-Leu zipper IV transcription factor PAS2 PASTICCINO2 Arabidopsis At5G10480 β-Hydroxyacyl-CoA dehydratase SHN2 SHINE2 Arabidopsis At5G11190 Homolog of WIN1/SHN1 WIN1/ SHN1 WAX INDUCER1/SHINE1 Arabidopsis At1G15360 AP2 domain-containing transcription factor WSD1 WAX SYNTHASE/ACYL-COENZYME A:DIACYLGLYCEROL ACYLTRANSFERASE1 Arabidopsis At5G37300 Wax synthase/acyl-CoA:diacylglycerol acyltransferase family protein WXP1 WAX PRODUCTION1 M. truncatula Medtr5G062700 AP2 domain-containing transcription factor – – Arabidopsis At5G33370 Homolog of CD1 Open in new tab Primary alcohols can be produced from VLCFA-CoA by fatty acyl-CoA reductase, an enzyme encoded by CER4 in Arabidopsis (Rowland et al., 2006). Free primary alcohols can occur in the wax mixture, or they can be esterified to a fatty acid in order to form wax esters. In this case, the alcohol is coupled to an acyl group derived from fatty acyl-CoA. The Arabidopsis enzyme responsible for this is WSD1, an enzyme of the wax synthase/diacylglycerol acyltransferase family (Li et al., 2008). A second branch of acyl wax biosynthesis leads to the formation of aldehydes and, ultimately, alkanes. Interestingly, in Arabidopsis, LACS1, which is also required for C16 cutin monomer biosynthesis, appears to have an additional specificity for C30 VLCFA and is required for the normal accumulation of downstream wax compounds (Lü et al., 2009). This suggests that conversion of an intracellular pool of free VLCFA back to VLCFA-CoA is an important route to aldehyde and alkane biosynthesis, rather than VLCFA-CoA directly derived from FAE. A long unresolved question in wax biosynthesis is the enzymatic basis of alkane synthesis. Classical biochemistry, using crude extracts from pea (Pisum sativum), indicated that the reaction likely occurs via the reduction of VLCFA-CoA to an aldehyde intermediate followed by decarbonylation, yielding an alkane that is 1C shorter (Cheesbrough and Kolattukudy, 1984; Schneider-Belhaddad and Kolattukudy, 2000). Although this enzyme was not purified and identified, compelling evidence was recently obtained, through studies of Arabidopsis, that CER1 and CER3 in complex act together to catalyze the formation of alkanes from VLCFA-CoA. It was shown by a split ubiquitin yeast two-hybrid assay and an Arabidopsis split luciferase assay that CER1 interacts with CER3 as well as several isoforms of cytochrome b5. Furthermore, heterologous expression of the combination of CER1, CER3, a cytochrome b5, and LACS1 in yeast resulted in the formation of very-long-chain alkanes (Bernard et al., 2012). This strongly suggests that a complex including CER1 and CER3 with cytochrome b5 as an electron donor catalyzes the reduction and decarbonylation of VLCFA-CoA in order to form cuticular alkanes. Aside from being a major component of the wax mixture, alkanes can undergo further modification to form secondary alcohols and ketones. In Arabidopsis, both of these oxidations are performed by the cytochrome P450 enzyme midchain alkane hydroxylase (MAH1; Greer et al., 2007). Synthesis of Cutin Precursors Cutin is typically composed of interesterified hydroxy fatty acids, with lesser amounts of glycerol, phenylpropanoids, and dicarboxylic acids (Kolattukudy, 2001). Chemical processes that cleave ester bonds, such as saponification, readily release these monomeric constituents, although in some species an additional lipidic polymer, referred to as cutan, remains recalcitrant to such treatments. Cutan is rich in ether and C-C bonds, but its structure is otherwise unknown, and it appears to be restricted to relatively few extant species (Gupta et al., 2006). The hydroxy fatty acids of cutin are typically ω-hydroxy fatty acids, usually with one or two additional midchain hydroxyl groups or an epoxy group (Fig. 3A). Despite extensive surveys of the chemical composition of plant cutins in the 1970s and 1980s (Kolattukudy, 2001), the composition of Arabidopsis cutin was not determined until relatively recently (Bonaventure et al., 2004; Franke et al., 2005). It is important to note that, in this important model species, the cutin of stems and leaves is atypical in that its major component is a dicarboxylic acid (Fig. 3A), implying that the predominant structural motif must be a copolymer with an unknown polyhydroxy compound, presumably glycerol (Pollard et al., 2008). However, despite the atypical composition of its cutin, Arabidopsis has proven to be an important model for deciphering the pathway of cutin biosynthesis, and more recently, it was discovered that the cutin of its floral organs is more typical, in that it is composed primarily of 10,16-dihydroxyhexadecanoic acid (Li-Beisson et al., 2009). Figure 3. Open in new tabDownload slide Typical cutin monomers and polymeric structure. A, Some typical C16 and C18 fatty acid-derived cutin monomers. From top to bottom: 10,16-dihydroxyhexadecanoic acid, 16-hydroxyhexadecanoic acid, 9,10-epoxyoctadecanoic acid, 9,10,18-trihydroxyoctadecanoic acid, and octadeca-cis-6,cis-9-diene-1,18-dioate, the major cutin monomer of Arabidopsis stems and leaves. B, Linear and branched domains made possible by different ester linkages of 10,16-dihydroxyhexadecanoic acid, depicted schematically as indicated. Figure 3. Open in new tabDownload slide Typical cutin monomers and polymeric structure. A, Some typical C16 and C18 fatty acid-derived cutin monomers. From top to bottom: 10,16-dihydroxyhexadecanoic acid, 16-hydroxyhexadecanoic acid, 9,10-epoxyoctadecanoic acid, 9,10,18-trihydroxyoctadecanoic acid, and octadeca-cis-6,cis-9-diene-1,18-dioate, the major cutin monomer of Arabidopsis stems and leaves. B, Linear and branched domains made possible by different ester linkages of 10,16-dihydroxyhexadecanoic acid, depicted schematically as indicated. While there is considerable diversity in the structure of cutin monomers, the pathway for the biosynthesis of 10,16-dihydroxyhexadecanoic acid-based cutin is the most complete, and the major themes of cutin biosynthesis are likely shared for other cutin monomers. Here, we summarize this pathway based on recent molecular genetic and biochemical studies using Arabidopsis and tomato (Solanum lycopersicum). Intracellular Acyltransferases and Hydroxylases The biosynthesis of cutin begins with de novo fatty acid synthesis in the plastid of epidermal cells (Fig. 2). The next three steps occur in the ER and consist of ω-hydroxylation and midchain hydroxylation and the synthesis of an acyl-CoA intermediate. The relative order of these steps is not known, although it has been shown that the ω-hydroxylation precedes the midchain hydroxylation and that the final product of these steps is most likely a dihydroxyhexadecanoic acid-CoA ester (Li-Beisson et al., 2009). The ω-hydroxylase is encoded by members of the CYP86 subfamily of cytochrome P450s (CYP86A4 in Arabidopsis flowers; Li-Beisson et al., 2009), while the midchain hydroxylase is encoded by the CYP77 subfamily (CYP77A6 in Arabidopsis flowers; Li-Beisson et al., 2009). The acyltransferases that synthesize acyl-CoA are encoded by the LACS family, which consists of nine members in Arabidopsis, and both LACS1 and LACS2 appear to be responsible for C16 cutin monomer biosynthesis (Lü et al., 2009). An additional intracellular acyltransferase required for the synthesis of cutin polyester is a glycerol 3-phosphate acyltransferase (GPAT). Recently, it was shown that plants possess a unique subfamily of bifunctional GPATs encoding enzymes with both sn-2-specific glycerol-3-phosphate:acyl-CoA acyltransferase activity as well as phosphatase activity, yielding a 2-monoacylglyceryl ester (Yang et al., 2010). In the case of Arabidopsis floral cutin, this activity is encoded by GPAT6 (Li-Beisson et al., 2009). Although the specific sequence of all intracellular biosynthetic steps will require additional characterization of the substrate specificity of each enzyme, biochemical characterization of Arabidopsis bifunctional GPATs indicates that they have a strong preference for ω-hydroxylated acyl-CoA, suggesting that hydroxylation precedes the transfer to glycerol (Yang et al., 2012). In any case, the ultimate product of the intracellular steps of cutin biosynthesis is likely to be 2-monoacylglyceryl esters of cutin monomers. In the case of 10,16-dihydroxyhexadecanoic acid-based cutin, this is 2-mono(10,16)-dihydroxyhexadecanoyl glycerol (2-MHG). Transport of Cuticle Precursors After the synthesis of wax and cutin precursors, they are exported from the ER, across the plasma membrane, through the polysaccharide cell wall, and to the nascent cuticular membrane. Most of these transport processes are poorly understood, although trafficking of both wax and cutin precursors across the plasma membrane has been shown to depend on ATP-binding cassette (ABC) transporters. In Arabidopsis, CER5/ABCG12 (Pighin et al., 2004) and ABCG11 (Bird et al., 2007) are required for wax export. Both of these encode half transporters, and based on double mutant analysis and bimolecular fluorescence complementation analyses, it has been suggested that an ABCG11/ABCG12 heterodimer is required for wax secretion (McFarlane et al., 2010). ABCG11 is also required for cutin accumulation, and since it is also able to dimerize with itself, it has been proposed that this homodimer is the functional complex responsible for cutin export (McFarlane et al., 2010). Additionally, a third Arabidopsis half transporter, ABCG13, was shown to be required for cutin deposition in flowers (Panikashvili et al., 2011). More recently, full transporters required for cutin deposition were identified in Arabidopsis (ABCG32; Bessire et al., 2011) as well as wild barley (Hordeum spontaneum) and rice (Oryza sativa; Chen et al., 2011). Despite the clear genetic evidence supporting a role for ABC transporters in cuticular lipid export, the substrate specificity of these transporters has not yet been demonstrated in vitro. However, all the ABC transporters that have been implicated in cuticle biosynthesis to date are members of the ABCG subfamily, which has been associated with the transport of lipids and hydrophobic compounds in other systems (Moitra et al., 2011). Moreover, in several cases, intracellular lipidic inclusions were observed in ABC transporter mutants, further supporting their direct involvement in cuticular lipid export (Pighin et al., 2004; Bird et al., 2007; Bessire et al., 2011). Export of some wax compounds also appears to be facilitated by glycosylphosphatidylinositol (GPI)-anchored lipid-transfer proteins (LTPs), LTPG1 and LTPG2, which are bound to the extracellular side of the plasma membrane (Debono et al., 2009; Lee et al., 2009; Kim et al., 2012). These proteins represent a unique class of LTPs, a family of small and typically soluble proteins that bind a variety of lipid substrates in vitro (Yeats and Rose, 2008). A major remaining question is how hydrophobic cuticle precursors are transported across the hydrophilic environment of the polysaccharide cell wall to the cuticle. Apoplastic LTPs have been proposed to play a role, although genetic or biochemical evidence for their involvement in transport is generally lacking (Yeats and Rose, 2008). In the case of the dihydroxyacyl cutin precursor 2-MHG, the glycerol moiety imparts sufficient polarity to allow aqueous solubility at low millimolar concentrations (Yeats et al., 2012b). This suggests that lipid-binding proteins or other factors are not necessary in order to facilitate the transport of this major precursor of cutin biosynthesis. However, the solubility of glyceryl esters of less polar cutin monomers has not been investigated, and they, along with waxes, may require additional factors to increase their solubility in the apoplast. Cutin Polymerization The final step of cutin synthesis is incorporation of the hydroxyacyl monomer into the polymer, but the molecular mechanism of cutin polymerization has been a longstanding enigma. Recent progress in this area was achieved by studying the tomato mutant cutin deficient1 (cd1) and transgenic tomato plants in which CD1 expression was suppressed using an RNA interference strategy (Girard et al., 2012; Yeats et al., 2012b). The cd1 mutant exhibits a severe reduction in the amount of polymerized cutin in the fruit cuticle (Isaacson et al., 2009), although chemical analysis indicated that, unlike wild-type fruit, those of the mutant accumulate nonpolymerized 2-MHG (Yeats et al., 2012b). Cloning of the mutated gene revealed that it encodes a protein of the GDSL-motif lipase/hydrolase (GDSL) family, which localizes to the developing cuticle (Girard et al., 2012; Yeats et al., 2012b). Despite its similarity to lipolytic enzymes, the recombinant protein acts as an acyltransferase in vitro, forming polyester oligomers from 2-MHG (Yeats et al., 2012b). The identification of CD1 as the first known cutin synthase raises several questions about the specificity and generality of the reaction that it catalyzes. Phylogenetic analysis of CD1 and homologous genes indicates that despite belonging to a very large gene family, the subfamily of GDSLs represented by CD1 is relatively small and well conserved, with sequences represented across diverse taxa of land plants (Volokita et al., 2011). In Arabidopsis, its putative orthologs form a five-member gene family, and silencing of the expression of two of these (LTL1 and At5g33370) resulted in plants exhibiting floral organ fusions and lacking nanoridges on the petal surface, phenotypes that are consistent with a cutin deficiency (Shi et al., 2011). An additional putative ortholog of CD1 from Agave americana exhibited similar localization and expression, further supporting a conserved mechanism of CD1-like enzymes acting as cutin synthases (Reina et al., 2007). Despite the presence of a null allele, the cd1 mutant is not completely deficient in cutin, so the identity of additional cutin synthases, or perhaps nonenzymatic mechanisms of cutin synthesis, represents an intriguing line of future research. The polymeric structure of cutin is not well understood. Monomeric composition can provide a “parts list,” but the relative abundance of possible linkages in the polymer is difficult to determine, largely due to the difficulty of solubilizing intact cutin (Serra et al., 2012). Nevertheless, the multiple functionalities present in many cutin monomers suggests that native cutin polymers can range from linear to branched or cross-linked structures (Pollard et al., 2008). For example, in an idealized cutin polymer composed exclusively of 10,16-dihydroxyhexadecanoic acid, the monomers can be joined by esterification of either the terminal or midchain hydroxyl group. Esterification of a single hydroxyl would result in a linear polymer, while esterification of both hydroxyl groups would generate branched structures (Fig. 3B). The identification of the hydroxyl groups that are esterified by CD1 and other cutin synthases should indicate whether the regiospecificity of cutin polymerization is enzymatically controlled and whether specific cutin synthases catalyze the formation of linear or branched domains of the cutin polymer. Moreover, it is not known how branching or cross linking of cutin affects cuticle function, and the identification of additional cutin synthases will allow this to be investigated using genetic approaches. REGULATION OF CUTICLE BIOSYNTHESIS The regulation of cuticle biosynthesis is complex and involves interacting signaling networks associated with environmental stress responses, pathogen responses, and feedback regulation based on the structure and integrity of the cuticle itself. Furthermore, as the cuticle is exclusively synthesized by epidermal cells, the regulation of epidermis identity during development can also be considered to play a regulatory role in cuticle development. This is covered in more depth in an excellent review by Javelle et al. (2011), and we focus here only on direct regulators of cutin and wax biosynthesis (Fig. 4). Even within this restricted context, the analysis of regulatory mutants is complicated by compensatory mechanisms between cutin and wax biosynthesis and other pleiotropic phenotypes. Nevertheless, a complex regulatory network that responds to developmental and environmental cues, mediated by hormones, transcription factors, and posttranscriptional regulation, is beginning to emerge. Figure 4. Open in new tabDownload slide Regulation of cuticle biosynthesis. A summary of the interaction of environmental factors and regulatory genes that are known to influence cutin or wax biosynthesis is shown. Figure 4. Open in new tabDownload slide Regulation of cuticle biosynthesis. A summary of the interaction of environmental factors and regulatory genes that are known to influence cutin or wax biosynthesis is shown. Environment and Hormones A systematic analysis of both cuticle composition and gene expression in Arabidopsis indicates that wax synthesis is induced by water deficit, sodium chloride, and abscisic acid (ABA) treatments (Kosma et al., 2009). In contrast, cutin biosynthesis was reported only to be induced by water deficit and not ABA or sodium chloride, suggesting that, at least in Arabidopsis, the detection of various osmotic stresses is complex and only partially dependent on ABA (Kosma et al., 2009). However, given that ABA is already well established as a regulator of plant responses to water deficit through the regulation of stomatal aperture (Lee and Luan, 2012), ABA regulation of cuticle biosynthesis is an intriguing area for further research aimed at understanding and engineering drought tolerance in crops. In addition, dark and cold treatments have been shown to reduce the expression of several components of the FAE complex (Hooker et al., 2002; Joubès et al., 2008). Several wax biosynthetic genes have been shown to be induced by bacterial pathogens (Raffaele et al., 2008) and during infestation of wheat (Triticum aestivum) by the Hessian fly (Mayetiola destructor; Kosma et al., 2010), but in general, the relevance of the induction of cuticle synthesis to pest or pathogen resistance is poorly understood. Transcription Factors and Cuticle Biosynthesis The first transcription factor gene identified as having a role in regulating cuticle biosynthesis was the AP2 domain-containing WAX INDUCER1/SHINE1 (WIN1/SHN1; Aharoni et al., 2004; Broun et al., 2004). Overexpression of this gene led to glossy leaves with a greater wax load than the wild type and lower transpiration, although this was likely due to a reduced density of stomata rather than the wax phenotype (Aharoni et al., 2004). Later studies indicated that cutin levels are also increased in WIN1/SHN1-overexpressing plants and that the up-regulation of genes encoding cutin biosynthetic enzymes precedes the induction of wax biosynthetic genes (Kannangara et al., 2007). WIN1/SHN1 is part of a three-member gene family in Arabidopsis, and silencing of all three genes led to a reduction in the amount of cutin but not waxes (Shi et al., 2011). These authors also demonstrated that these transcription factors directly activate promoters of several cutin biosynthetic genes, further supporting a primary role in cutin regulation with a downstream effect on wax biosynthesis (Shi et al., 2011). In addition to regulating cutin biosynthesis, the SHN transcription factors also induced the expression of several pectin-modifying enzymes, suggesting a coordination of the synthesis of the cuticle with the polysaccharide cell wall (Shi et al., 2011). This second function of SHN transcription factors in regulating the polysaccharide cell wall is further suggested by experiments in which the overexpression of Arabidopsis SHN2 in rice resulted in a significant increase in the amount of cellulose and a concomitant decrease in lignin (Ambavaram et al., 2011). On the other hand, a general role of WIN1/SHN1-related transcription factors in the regulation of cutin synthesis is indicated by studies of orthologous genes in barley (Hordeum vulgare; Taketa et al., 2008) and tomato (Shi et al., 2013). The balance of evidence thus suggests that SHN transcription factors coordinate not just the synthesis of cutin but also the polysaccharide cell wall of the epidermis. This ultimately highlights the fact that the cuticle is a specialized modification of the cell wall, and like other modifications, such as lignification or suberization, it should be considered within the context of polysaccharide cell wall components. Aside from the SHN family, other AP2 domain transcription factors from different clades may also play a role in cuticle regulation. For example, overexpression of WXP1 from Medicago truncatula in alfalfa (Medicago sativa) induced wax production (Zhang et al., 2005). Recently, two transcription factors, MYB106 and MYB16, were identified as regulators of cuticle biosynthesis that function in a similar manner to WIN1/SHN1 (Oshima et al., 2013). They both appear to act upstream of, and directly activate, WIN1/SHN1 but also some cuticle biosynthetic genes (Oshima et al., 2013). Several other transcription factors of the MYB family have also been implicated in the regulation of wax and cutin biosynthesis in response to environmental stresses. MYB30 is induced during infection by bacterial pathogens, leading to the up-regulation of several genes of the FAE complex, and ectopic overexpression of MYB30 leads to an increased wax load (Raffaele et al., 2008). MYB96 was identified as an ABA-inducible transcription factor that mediates drought tolerance (Seo et al., 2009), in part due to an induction of wax biosynthesis resulting from MYB96 directly activating the promoters of several wax synthesis genes (Seo et al., 2011). While MYB96 positively regulates wax production in response to stress, MYB41 mediates the negative regulation of cutin biosynthesis in response to similar stresses. MYB41 is induced by ABA, drought, and osmotic stress, leading to the down-regulation of cutin biosynthesis genes and the disruption of cuticle structure (Cominelli et al., 2008). Another regulatory factor was identified through characterization of the rice CURLY FLAG LEAF1 (CFL1) gene, which encodes a WW domain-containing protein that negatively regulates cuticle biosynthesis. Studies of the orthologous CFL1 gene in Arabidopsis indicated that it down-regulates cutin biosynthesis by suppressing the function of HDG1, a homeodomain-leucine zipper IV transcription factor (HD-ZIP IV), which has been shown to induce the expression of several cutin biosynthesis genes (Wu et al., 2011). A more general role of HD-ZIP IV proteins in regulating cutin synthesis is further suggested by the homologous tomato gene CD2, which is required for the biosynthesis of cutin in the fruit and other organs (Isaacson et al., 2009; Nadakuduti et al., 2012). In maize (Zea mays), the HD-ZIP IV gene OUTER CELL LAYER1 (OCL1) was shown to be an epidermis-specific positive regulator of wax biosynthesis, although cutin was not quantified in plants overexpressing this gene (Javelle et al., 2010). Interestingly, HD-ZIP IV proteins have also been implicated in regulating other epidermis-specific processes, such as trichome differentiation and the formation of root hairs and stomatal guard cells (Masucci et al., 1996; Nakamura et al., 2006; Takada et al., 2013). Given their additional association with cuticle biosynthesis, it appears that a common feature of members of this protein family is playing key roles in the biology of the plant epidermis and the determination of epidermal cell fate. Beyond Transcription Factors In addition to the network of transcription factors that regulate cuticle biosynthesis, regulatory mechanisms that do not involve direct transcriptional activation or repression by promoter binding have recently been discovered. A recent example resulted from studies of the Arabidopsis cer9 mutant, which exhibits alterations in the amount and composition of leaf and stem waxes. Cloning of the CER9 gene revealed it to encode a protein with sequence similarity to yeast Doa10, an E3 ubiquitin ligase involved in ER-associated degradation of misfolded proteins (Lü et al., 2012). Given the ER localization of wax and cutin biosynthetic processes, the authors proposed a role for CER9 in the homeostasis of key cuticle biosynthetic enzyme levels. Experiments further addressing this hypothesis will be particularly interesting, given the surprising finding that the cer9 mutant actually exhibits enhanced drought tolerance and water use efficiency (Lü et al., 2012). One of the most intriguing mechanisms of cuticle regulation resulted from characterization of the cer7 mutant. CER7 encodes an exosomal exoribonuclease, and the cer7 mutant exhibits reductions in stem wax and transcription of CER3, a major wax biosynthetic enzyme (Hooker et al., 2007). Recently, two suppressors of cer7 that restore the CER3 transcript and stem wax levels were identified, and cloning of the respective genes identified RDR1 and SGS3, two conserved components of the RNA-mediated gene-silencing pathway (Lam et al., 2012). A model was proposed wherein CER7 is involved in the degradation of a small RNA species that negatively regulates the CER3 transcript. Future work involving the identification of such a small RNA species and other components of this pathway will be especially intriguing, since no known plant small RNA species mapped to the CER7-dependent region of the CER3 promoter (Lam et al., 2012). ENIGMATIC FACTORS IN CUTICLE BIOSYNTHESIS In addition to the characterized components of cuticle biosynthesis that can be incorporated into a coherent model, as discussed above, several genes/proteins have been identified that are required for cuticle formation but that lack a clear associated biochemical function that would place them in a specific point in the pathways. One example is HOTHEAD (HTH), a Glc-methanol-choline oxidoreductase family protein that is required for proper cuticle organization (Krolikowski et al., 2003). Chemical analysis indicated that the Arabidopsis hth mutant has wild-type wax levels but abnormal cutin quantity and composition. Specifically, it has decreased levels of dicarboxylic acids and increased amounts of ω-hydroxy acids, leading the authors to suggest that HTH may have a role in the oxidation of ω-hydroxy fatty acids to the dicarboxylic acid cutin monomers that are characteristic of Arabidopsis stem and leaf cuticles (Kurdyukov et al., 2006b). As dicarboxylic acid cutin monomers are unusually abundant in Arabidopsis, it will be interesting to see whether HTH-related proteins are as essential to cuticle formation in other species where this class of monomers is scarce. Another example of an “orphan” cuticle-associated protein resulted from analysis of the Arabidopsis bodyguard (bdg) mutant, which exhibits a microscopically disorganized cuticle with increased permeability but significantly increased levels of wax and cutin (Kurdyukov et al., 2006a). The BDG protein has sequence similarity to the α/β-hydrolase family of proteins, but no enzymatic activity has been reported. The protein is localized in the outer cell wall of the epidermis below the cuticle, which led the authors to propose that BDG may be involved in cutin polymerization, although the increased amounts of polymeric cutin in the mutant would argue against this (Kurdyukov et al., 2006a). Mutation of BDG3, a close homolog of BDG, resulted in the disorganization of floral nanoridges, petal epidermis structures that are composed of cutin (Shi et al., 2011). Moreover, the key cutin regulatory transcription factors SHN1, SHN2, and SHN3 were shown to activate the BDG3 promoter (Shi et al., 2011). Taken together, these results strongly indicate that BDG proteins are closely linked to cutin polymer formation, although their mode of action remains mysterious. Lastly, a defect in the formation of floral nanoridges was also identified in the Arabidopsis mutant defective in cuticular ridges (dcr), which showed a substantial deficiency in floral cutin but a less drastic alteration of leaf and stem cutin (Panikashvili et al., 2009). DCR encodes a protein of the BAHD acyltransferase family that localizes to the cytoplasm, and it has been proposed that it may be involved in acyl transfer of cutin monomers to form precursor intermediates or oligomeric structures (Panikashvili et al., 2009). However, DCR was later biochemically characterized and shown to possess in vitro diacylglycerol acyltransferase activity, leading to the formation of triacylglycerol (Rani et al., 2010). A role for cytoplasmic triacylglycerol intermediates in cutin biosynthesis is not consistent with any known steps in this pathway, yet DCR is clearly required for cutin biosynthesis in Arabidopsis floral organs. Further work will be needed in order to determine the native substrate and product of DCR in order for its role in cutin biosynthesis to be elucidated. FUNCTIONS OF THE CUTICLE The plant cuticle is most typically associated with providing a fixed barrier to excessive transpirational water loss, allowing gas exchange and transpiration to be dynamically controlled by stomata. However, it has evolved a number of secondary functions that are consistent with its place as the outermost layer of primary aerial organs: it forms a physical barrier that is the first line of defense against pests and pathogens; in many species, elaborate epicuticular crystals help to form a self-cleaning surface, preventing dust and other debris from blocking sunlight; in some cases, it can act to screen excessive UV light; finally, as a defining feature of the epidermis, it plays a central role in development by physically establishing organ boundaries. Cuticle Structure and Water Barrier Properties A common perception is that a thick cuticle is associated with a lower water permeability and thus increased tolerance to water stress. However, comparative studies of the water permeability of cuticles from diverse species have indicated that there is no correlation with either the thickness of the cuticle or the amount of wax (Riederer and Schreiber, 2001). Similarly, the amount of cutin is not necessarily an indication of cuticular water permeability (CWP). For example, studies of three tomato mutants (cd1–cd3), each of which has a greater than 95% reduction in fruit cutin levels, revealed only minor increases in the rate of water loss, and even among the mutants there was no clear correlation between cutin amount and susceptibility to desiccation (Isaacson et al., 2009). However, cutin deficiency that leads to organizational defects can be detrimental to the cuticle permeability (Bessire et al., 2011). In contrast to the lack of association with cutin, extensive removal of wax from tomato fruit, accomplished by brief immersion of the fruit in an organic solvent, indicates that waxes contribute approximately 95% of the cuticle-mediated resistance to water diffusion, at least in tomato fruit (Leide et al., 2007). Specific compound classes appear to be associated with water barrier properties of the cuticle; notably, the more nonpolar components, such as alkanes, tend to be associated with decreased CWP, while nonaliphatic wax compounds, such as triterpenoids, are likely a less effective water barrier (Leide et al., 2007; Buschhaus and Jetter, 2012). This is consistent with a model in which cuticular waxes localize within either crystalline or amorphous domains of the cuticle, with aliphatic compounds forming crystallite “rafts” that are impervious to water, forcing water, and other polar metabolites, to diffuse by a circuitous route through the amorphous domains that are formed by more polar and cyclic waxes (Riederer and Schreiber, 1995). The idea that the proportion of alkanes and not the total wax amount has the most significant effect on CWP was illustrated by a recent study with a backcrossed population of Capsicum annum and Capsicum chinense, two pepper species with high and low postharvest water loss rates, respectively. In 20 backcrossed families, CWP was inversely correlated with the amount of alkanes in the wax but not the total amount of wax, and the more rapidly desiccating parent had three times the wax coverage as the parent that exhibited low postharvest water loss (Parsons et al., 2012). In summary, resistance to water loss is primarily attributed to wax and not cutin, but there is not a direct correlation between the amount of either component and CWP. Rather, it appears that CWP is primarily determined by the particular mixture of intracuticular and epicuticular waxes and by their packing and organization within the cuticle architecture. The Lotus Effect A striking feature of many plant leaves is that water tends to bead into drops and roll to the ground, collecting and washing particles and debris from the leaf surface. The efficiency of this self-cleaning mechanism, termed the “lotus effect,” varies between species and during organ ontogeny, but it has been correlated with the abundance of epicuticular wax crystals that repel water and allow a pocket of air to form beneath the droplets (Barthlott and Neinhuis, 1997). It is thought that this self-cleaning surface helps to prevent the buildup of dust that would block sunlight and slow photosynthesis and that this could also play an important role in washing away pathogen spores before they germinate. Despite the apparent advantages of a self-cleaning surface, there is not a clear example of this trait conferring an adaptive advantage. In terms of photosynthesis, there is likely a tradeoff between a self-cleaning surface and the increased dispersion of light by epicuticular wax crystals, as discussed below. Nevertheless, based on the discovery of this effect, surfaces with high degrees of hydrophobicity and microscopic texture have been employed as effective biomimetic technical materials (Bhushan, 2012), and improved self-cleaning surfaces in agricultural crops may be a productive avenue of research. The Cuticle as a Barrier against Pests and Pathogens The plant cuticle presents a physical barrier to pathogens that do not otherwise enter the plant by way of the stomata, wounds, or vectors. However, fungal pathogens have been shown to breach the cuticle using a combination of enzymatic degradation and mechanical rupture. The latter is often accomplished by the formation of a swollen appressorium structure that extends an infectious peg via turgor pressure (Deising et al., 2000). While mechanical rupture may be sufficient for cuticle penetration, particularly of thinner cuticles (Tenberge, 2007), most fungal pathogens also secrete cutinases, a class of small, nonspecific esterases that hydrolyze the cutin polyester and release free cutin monomers (Longhi and Cambillau, 1999). The cutin monomers that are released during polymeric cutin hydrolysis can act as elicitors of plant defense responses and are thus classified as damage-associated molecular patterns. At micromolar concentrations, these compounds induce the production of hydrogen peroxide and other defense responses (Schweizer et al., 1996; Kauss et al., 1999). However, the mechanism of plant perception of free cutin monomers is currently unknown (Boller and Felix, 2009). Cutin appears to be more important than wax for forming a barrier to pathogen entry, although there is not a consistent correlation between cutin amount and pathogen resistance. In tomato fruit, severely decreased cutin levels in three cd mutants was associated with increased susceptibility to infection by Botrytis cinerea surface inoculation and also to opportunistic microbes (Isaacson et al., 2009). However, in Arabidopsis, a number of cutin-deficient mutants and plants that ectopically overexpress fungal cutinases exhibit enhanced resistance to B. cinerea (Bessire et al., 2007, 2011; Chassot et al., 2007; Tang et al., 2007). In this case, increased cuticular permeability appears to enhance the diffusion of inoculum-derived elicitors that induce the production of small, polar antifungal compounds, which in turn inhibit B. cinerea growth (Bessire et al., 2007). Conversely, the Arabidopsis lacs2 mutant and cutinase overexpressers exhibited no alteration in their susceptibility to a range of other fungal pathogens (Bessire et al., 2007), and the lacs2 mutation also increased susceptibility to a normally avirulent strain of Pseudomonas syringae (Tang et al., 2007). Thus, cutin plays an important role as a physical barrier to many pathogens, yet extreme deficiencies in Arabidopsis can result in increased resistance to some pathogens by way of a secondary, but not well understood, mechanism that involves the induction of plant defenses. An additional layer of complexity was suggested by the observation that cutin can induce gene expression in plant pathogens and has been shown to induce appressorium expression in Colletotrichum trifolii (Dickman et al., 2003). This highlights the competing selective pressures to generate and breach cuticle barriers at the frontier of the plant surface (Chassot and Metraux, 2005). Despite the importance of cutin in plant-pathogen interactions, the first surface encountered by foliar pathogens is formed by epicuticular wax crystals and films. In addition to the lotus effect that promotes the washing of spores from the plant surface before germination, there are several indications that the epicuticular wax structures and composition are important in determining fungal pathogen development and, thus, pathogenicity. The C26 aldehyde n-hexacosanyl, a component of cuticular wax in many species of the Poaceae, can induce in vitro appressorium formation by the powdery mildew Blumeria graminis (Tsuba et al., 2002; Ringelmann et al., 2009; Hansjakob et al., 2010). This observation is further corroborated by studies of the maize mutant glossy1, which does not accumulate aldehydes in its wax complement. B. graminis appressorium formation is substantially reduced on the leaf surface of the glossy1 mutant but can be restored to normal levels by the application of n-hexacosanyl (Hansjakob et al., 2011). Another example of the influence of waxes on pathogenicity is provided by the inhibitor of rust tube germination1 (irg1) mutant of M. truncatula, which exhibits decreased amounts of epicuticular wax crystals on the abaxial leaf surface, corresponding to a substantial decrease in wax primary alcohol groups. This surface alteration was shown to reduce spore differentiation of the rust fungal pathogens Phakopsora pachyrhizi and Puccinia emaculata and the anthracnose fungus C. trifolii, resulting in nonhost resistance (Uppalapati et al., 2012). The IRG1 gene was found to encode a C2H2 zinc finger transcription factor that had previously been identified as a regulator of dissected leaf morphology (Chen et al., 2010). Reduced transcript levels of putative MYB96 and CER4 orthologs were also observed in the irg1 mutant, which is consistent with the wax phenotype. The significance of waxes and cutin in pathogen resistance, therefore, is suggested in a general sense, but, as with cuticle permeability, little is known about the relative importance of specific molecular classes or their intermolecular associations and packing within the architecture of the cuticle. Epicuticular waxes may also play an important role in plant-insect interactions; indeed, epicuticular wax crystals can form an unstable surface that prevents insect attachment or locomotion on plant surfaces (Borodich et al., 2010). A striking example of this is seen in the carnivorous pitcher plants (Nepenthes spp.), which catch insects by way of a slippery interior surface that is coated with epicuticular wax crystals (Riedel et al., 2007). For a more detailed review of cuticle chemical ecology, see Müller and Riederer (2005). The Cuticle and Development In addition to providing physical barriers to water and microbes, the cuticle appears to play an important role in defining organ boundaries during development, since plants with cuticles showing increased permeability and structural defects often exhibit numerous ectopic organ fusions. This phenomenon has been observed in a wax-deficient tomato mutant (Smirnova et al., 2013), a range of Arabidopsis mutants with abnormal cuticles (Yephremov et al., 1999; Wellesen et al., 2001; Kurdyukov et al., 2006a; Bird et al., 2007), and transgenic Arabidopsis plants overexpressing a secreted fungal cutinase (Sieber et al., 2000). The fusion zones are often marked by two adjacent polysaccharide cell walls with no visible cuticle separating the two organs, although the fused epidermal layers maintain their identity, as indicated by the differentiation of internal nonfunctional stomata within fusion zones (Sieber et al., 2000). In each of three Arabidopsis mutants exhibiting organ fusions, lacerata, bodyguard, and fiddlehead, ectopic organ fusions and cuticular permeability defects could be partially suppressed by a second mutation in SERRATE (Voisin et al., 2009). SERRATE is a C2H2 zinc finger protein that is required for microRNA biogenesis, and hypomorphic alleles exhibit numerous developmental defects, including serrated leaf margins (Dong et al., 2008). While the mechanism of SERRATE action as a suppressor of cuticle fusions remains unclear, this result suggests the existence of a cuticle integrity pathway that is integrated with epidermal developmental programs. The identification of additional suppressors of cuticle mutant-associated developmental phenotypes should be informative in elucidating the cuticle integrity pathway. Protection against UV Radiation UV light in the UV-B spectrum is a considerable portion of the daylight that reaches the terrestrial surface, and it can threaten plant life by damaging DNA, the photosynthetic apparatus, and membrane lipids (Rozema et al., 1997). As a result, plants have evolved a number of strategies for screening UV-B radiation. These include a variety of soluble flavonoid pigments that are typically localized within the vacuoles of epidermal cells, phenolic compounds present in the polysaccharide cell wall, and lipophilic phenolic molecules that are covalently bound to cutin or associated with waxes (Pfündel et al., 2006). A survey of isolated cuticles from a range of species indicated generally effective screening of the UV-B spectrum but consistently high transmittance in the higher wavelengths that are photosynthetically active (Krauss et al., 1997). In addition to absorbing light, the plant cuticle can reflect light to some degree, presumably depending on the abundance of epicuticular wax crystals. For example, Dudleya brittonnii can reflect up to 83% of UV-B, but this value is substantially reduced when epicuticular waxes are removed (Mulroy, 1979). Smooth, glossy “glabrous” cuticles typically reflect only small amounts of light (less than 10%), but glaucous plant surfaces are moderately reflective and generally show approximately 20% to 30% reflectance in the UV and visible spectra (Pfündel et al., 2006). Waxes reflect both UV and visible light, but not necessarily to the same extent, and the reflectance of UV has been reported to be greater in some cases (Holmes and Keiller, 2002). While light reflection provides an important protective mechanism, especially by limiting damaging UV radiation, there is likely a tradeoff with photosynthetic efficiency under conditions when light intensity is limiting (Pfündel et al., 2006). In this regard, an interesting area of future research might to determine whether relative proportions of UV and visible light reflection can be predictively changed by altering the composition of epicuticular waxes. CONCLUSION AND PERSPECTIVES As described above, several key areas of cuticle biogenesis remain poorly understood. First, the mechanism of intracellular and extracellular transport of wax and cutin precursors remains unknown, although key ABC transporters required for their export across the plasma membrane have been identified (Pighin et al., 2004; Bird et al., 2007; Chen et al., 2011). The first cutin synthase has been identified (Girard et al., 2012; Yeats et al., 2012b), but there are certainly additional cutin synthases, and whether they are closely related to CD1 or belong to distinct protein families remains to be discovered. After cutin is polymerized, is modification of the polymeric structure required to accommodate organ expansion? If so, which enzymes are involved in this process? While our understanding of cuticle biosynthesis at the molecular level remains incomplete, recent progress in deciphering these pathways is bringing us closer than ever to an ability to selectively modify cuticle properties in order to improve agricultural productivity. However, the ability to make such modifications rationally will require an understanding of the complexity of cuticle function at the molecular level, and far less progress has been made in this regard. To this end, further work aimed at understanding the ecophysiological functions of the cuticle in defined mutant backgrounds, as well as in genetically tractable wild species, will provide a framework for understanding the complex interaction of structure, composition, and function of cuticles (Yeats et al., 2012a). While the past decade has seen unprecedented progress in the molecular biology of cuticle biogenesis, many studies have revealed complexities in cuticle function that underscore the fact that the cuticle is much more than just a preformed barrier to water loss. ACKNOWLEDGMENTS We thank Drs. Gregory Buda, Christiane Nawrath, and Lacey Samuels for generously providing microscopy images and Eric Fich, Laetitia Martin, and Dr. Iben Sørensen for helpful comments and discussion. Glossary VLCFA very-long-chain fatty acid ER endoplasmic reticulum FAE fatty acid elongase ABC ATP-binding cassette LTP lipid-transfer protein GPI glycosylphosphatidylinositol ABA abscisic acid CWP cuticular water permeability LITERATURE CITED Aharoni A Dixit S Jetter R Thoenes E van Arkel G Pereira A ( 2004 ) The SHINE clade of AP2 domain transcription factors activates wax biosynthesis, alters cuticle properties, and confers drought tolerance when overexpressed in Arabidopsis . 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Plant Cell 17 : 1467 – 1481 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the U.S. Department of Agriculture Agriculture and Food Research Initiative Competitive Grants Program (grant no. 2011–67013–19399), by the National Institute of Food and Agriculture, and by the National Science Foundation Plant Genome Research Program (grant no. DBI–0606595). 2 Present address: Energy Biosciences Institute, University of California, Berkeley, CA 94720. * Address correspondence to [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Jocelyn K.C. Rose ([email protected]). www.plantphysiol.org/cgi/doi/10.1104/pp.113.222737 © 2013 American Society of Plant Biologists. All Rights Reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Duangpan, Saowapa; Zhang, Wenli; Wu, Yufang; Jansky, Shelley H.; Jiang, Jiming

doi: 10.1104/pp.113.221903pmid: 23898040

Abstract Insertional mutagenesis using transfer DNA or transposable elements, which is an important tool in functional genomics and is well established in several crops, has not been developed in potato (Solanum tuberosum). Here, we report the application of the tobacco (Nicotiana tabacum) Tnt1 retrotransposon as an insertional mutagen in potato. The Tnt1 retrotransposon was introduced into a highly homozygous and self-compatible clone, 523-3, of the diploid wild potato species Solanum chacoense. Transposition of the Tnt1 elements introduced into 523-3 can be efficiently induced by tissue culture. Tnt1 preferentially inserted into genic regions in the potato genome and the insertions were stable during sexual reproduction, making Tnt1 an ideal mutagen in potato. Several distinct phenotypes associated with plant stature and leaf morphology were discovered in mutation screening from a total of 38 families derived from Tnt1-containing lines. We demonstrate that the insertional mutagenesis system based on Tnt1 and the 523-3 clone can be expanded to the genome-wide level to potentially tag every gene in the potato genome. Insertional mutagenesis is one of the most important tools in plant functional genomics. Application of T-DNA, the transfer DNA of the Ti plasmid of Agrobacterium tumefaciens, was the first and the most successful methodology of genome-wide insertional mutagenesis in plants. Many T-DNA lines were developed in two of the most important model plant species, Arabidopsis (Arabidopsis thaliana) and rice (Oryza sativa; Krysan et al., 1999; Jeon et al., 2000; Alonso et al., 2003; Sallaud et al., 2004). These T-DNA stocks have served as the foundation for the identification and characterization of numerous genes in these two species. A major limitation of the T-DNA-based technique is the requirement for a highly efficient transformation system to generate a large number of transgenic lines. Unfortunately, in many plant species, A. tumefaciens-based transformation is either not yet developed or is not efficient enough to produce a sufficient number of T-DNA lines that would allow a genome-wide gene tagging. Several transposable elements (TEs) have been used for insertional mutagenesis in plants, including the Activator and Mutator transposons in maize (Zea mays; Walbot, 1992), the Tam3 transposon in Antirrhinum majus (Luo et al., 1991), the Tos17 retrotransposon in rice (Hirochika, 2010), and the LORE1 retrotransposon in Lotus japonicus (Fukai et al., 2012). Most remarkably, the Tnt1 retrotransposon, originally identified in tobacco (Nicotiana tabacum; Grandbastien et al., 1989), has been successfully used in insertional mutagenesis in several heterologous species, including Arabidopsis (Courtial et al., 2001), Medicago truncatula (d’Erfurth et al., 2003), lettuce (Lactuca sativa; Mazier et al., 2007), and soybean (Glycine max; Cui et al., 2013). Tnt1 was used to generate approximately 12,000 independent lines that represent over 300,000 insertions in M. truncatula (Tadege et al., 2008; Cheng et al., 2011). Many M. truncatula genes have been identified through forward and reverse genetics approaches using the Tnt1 retrotransposon insertion population (Pang et al., 2009; Zhao et al., 2010; Laurie et al., 2011; Tadege et al., 2011; Zhou et al., 2011; Cheng et al., 2012; Bourcy et al., 2013). Potato (Solanum tuberosum) is one of the most important food crops in the world. Cultivated potato is an autotetraploid (2n = 4x = 48) with a highly heterozygous genome. These characteristics make potato a poor model for both forward and reverse genetics research. Very few potato genes have been cloned using the traditional map-based cloning strategy. The lack of fertile homozygous clones makes insertional mutagenesis infeasible in potato. However, the recent sequencing of the potato genome (Xu et al., 2011) has significantly changed the status of potato genetics and genomics research. Highly efficient genotyping systems based on DNA microarrays or DNA sequencing will make genetic mapping and association mapping much more efficient in potato (Hamilton et al., 2011; Felcher et al., 2012). We expect an acceleration of forward genetics-based gene identification in potato. A genome-wide mutagenesis system is urgently needed for the potato research community. Here, we report the development of an insertional mutagenesis system in potato using the Tnt1 retrotransposon. This system is built on a highly homozygous and self-compatible clone (523-3) of the diploid potato species Solanum chacoense (2n = 2x = 24), one of the most widespread wild Solanum species (Miller and Spooner, 1996). S. chacoense is sexually compatible with diploid cultivated potato (Leue and Peloquin, 1980; Hermundstad and Peloquin, 1985). It has been an important germplasm source for the improvement of potato cultivars, especially those for use in the processing market (Love et al., 1998). We demonstrate that this system can be used to potentially tag every potato gene and serve as an important foundation for potato functional genomics research. RESULTS Transposition of Tnt1 Occurred during in Vitro Transformation of Potato A self-compatible clone of S. chacoense was first reported by Hosaka and Hanneman (1998a, 1998b). An S7 clone from the Hanneman program, 523-3, was derived from seven generations of selfing from the original S. chacoense clone. The high level of homozygosity of 523-3 was confirmed by a genome-wide single-nucleotide polymorphism-based genotyping (S.H. Jansky, unpublished data) and the uniform phenotype of the progeny derived from selfing 523-3. The 523-3 clone readily self-pollinates and can generate an average of 100 seeds per fruit. More importantly, this clone can produce tubers in both greenhouse and field conditions. We developed several transgenic 523-3 lines by transforming internode explants using A. tumefaciens strain GV3101 containing an autonomous copy of Tnt1 in plasmid Tnk23 (d’Erfurth et al., 2003; Fig. 1A). A total of 17 transgenic plants (T0) were generated and confirmed by PCR for the presence of both the nptII gene and Tnt1-specific sequences. Genomic DNA was isolated from the T0 plants, double digested with HindIII (three sites in the construct; Fig. 1A) and EcoRI (no site in the construct), and hybridized sequentially by an nptII gene probe (Fig. 1B) and a Tnt1-specific probe (Fig. 1C), respectively. Figure 1. Open in new tabDownload slide Transpositions of Tnt1 elements in transgenic potato lines. A, Structure of the Tnk23 T-DNA plasmid. The two small horizontal arrows point to the Tnt1-specific probe designed from the long terminal repeat (LTR) region. Vertical black arrows point to the three HindIII sites within the Tnt1 element. Each Tnt1 element will result in an 879-bp fragment after HindIII digestion. LB, Left border; RB, right border. B, Southern-blot hybridization of five T0 lines using an nptII-specific probe. Genomic DNAs were double digested with EcoRI and HindIII. Each hybridization band represents a single transgenic Tnt1 site. C, Southern-blot hybridization of the same blot as in B using the Tnt1-specific probe. The expected 879-bp fragment, as indicated by the black arrow, was observed in every line. Each of the remaining bands represents an independent Tnt1 element. Figure 1. Open in new tabDownload slide Transpositions of Tnt1 elements in transgenic potato lines. A, Structure of the Tnk23 T-DNA plasmid. The two small horizontal arrows point to the Tnt1-specific probe designed from the long terminal repeat (LTR) region. Vertical black arrows point to the three HindIII sites within the Tnt1 element. Each Tnt1 element will result in an 879-bp fragment after HindIII digestion. LB, Left border; RB, right border. B, Southern-blot hybridization of five T0 lines using an nptII-specific probe. Genomic DNAs were double digested with EcoRI and HindIII. Each hybridization band represents a single transgenic Tnt1 site. C, Southern-blot hybridization of the same blot as in B using the Tnt1-specific probe. The expected 879-bp fragment, as indicated by the black arrow, was observed in every line. Each of the remaining bands represents an independent Tnt1 element. All T0 lines contained at least one copy of the transgenic Tnt1 element, based on the number of bands hybridized to the nptII gene. The numbers of transgenic Tnt1 elements among the T0 lines ranged from one to six, based on the number of nptII gene-specific bands on the Southern blots (Fig. 1B). The Tnt1-specific probe hybridized to two DNA fragments derived from each Tnt1 element, a common 879-bp fragment and another fragment that represents an individual Tnt1 element (Fig. 1C). The 879-bp fragment was detected in every T0 line. The numbers of total Tnt1 elements among the T0 lines ranged from one to more than 20, based on Tnt1-specific hybridization bands (Fig. 1C). The total numbers of Tnt1 elements in all T0 lines, except line CT1, were more than the numbers of the original transgenic Tnt1 elements. These results suggest that some of the transgenic Tnt1 elements transposed immediately after integrating into the 523-3 genome, resulting in additional Tnt1 elements that are not associated with the nptII gene. Tnt1 Transposition Can Be Induced by in Vitro Regeneration Tissue culture-induced retrotransposition of Tnt1 was previously reported in several plant species (Courtial et al., 2001; d’Erfurth et al., 2003; Mazier et al., 2007; Cui et al., 2013). We wanted to confirm whether tissue culture can also induce Tnt1 transposition in the potato genome. The T0 line CT1, which contains a single copy of Tnt1 (Fig. 1), was selected for in vitro regeneration experiments. Internode explants were cut from in vitro CT1 plants and were transferred to regeneration medium. The explants were kept in an incubator and were transferred to new medium every 2 weeks. Regenerated shoots were then transferred to root-inducing medium. We developed a total of 13 regenerated T1 plants (CT1:1–CT1:13). Southern-blot hybridization of the regenerated plants revealed that each of the 13 T1 plants contains at least one additional copy of Tnt1 compared with CT1 (Fig. 2). The numbers of additional Tnt1 elements ranged from one to more than 20 among the 13 plants. These results showed that tissue culture can effectively induce Tnt1 transpositions in potato. Figure 2. Open in new tabDownload slide Tissue culture-induced transposition of Tnt1 in potato. DNAs from the original transgenic line CT1 and four regenerated lines from CT1 were digested with both EcoRI and HindIII and hybridized with the Tnt1-specific probe. Arrows point to the two bands derived from the original Tnt1 element from CT1. Other bands represent new Tnt1 elements retrotransposed from the original copy. Figure 2. Open in new tabDownload slide Tissue culture-induced transposition of Tnt1 in potato. DNAs from the original transgenic line CT1 and four regenerated lines from CT1 were digested with both EcoRI and HindIII and hybridized with the Tnt1-specific probe. Arrows point to the two bands derived from the original Tnt1 element from CT1. Other bands represent new Tnt1 elements retrotransposed from the original copy. Tnt1 Insertions Are Stable during Sexual Reproduction Reverse genetics using TE as a mutagen involves self-pollination to develop homozygous insertional mutants. Therefore, the stability of inserted TEs during sexual reproduction is essential for maintaining the mutation. To investigate the stability of the Tnt1 elements in the potato genome, we conducted Southern-blot hybridization analysis of 80 selfed progeny derived from each of four T1 lines (CT1:3, CT1:4, CT1:5, and CT1:7). For each T1 line, DNA samples isolated from 20 selfed progenies were pooled, digested with HindIII and EcoRI, and hybridized with the Tnt1-specific probe. We did not observe any additional hybridization bands in these progenies compared with the parental T1 lines (Fig. 3A), suggesting that the Tnt1 elements in these four T1 lines were stable during sexual reproduction. Figure 3. Open in new tabDownload slide Stability of Tnt1 insertions during sexual reproduction. A, Genomic DNAs were pooled from 20 selfed progenies from one of the four lines (CT1:3, CT1:4, CT1:5, and CT1:7), digested with EcoRI and HindIII, and hybridized to a Tnt1-specific probe. An 879-bp fragment can be observed in each lane. The hybridization patterns associated with the selfed progenies are identical to the patterns of the parental lines. B, Southern-blot hybridization analysis of two DNA samples. Lane 1, DNA isolated from CT1:3; lane 2, pooled DNA from 19 progenies of CT1:3 and genomic DNA from CT1:4. Arrows point to the two bands associated with CT1:4. Figure 3. Open in new tabDownload slide Stability of Tnt1 insertions during sexual reproduction. A, Genomic DNAs were pooled from 20 selfed progenies from one of the four lines (CT1:3, CT1:4, CT1:5, and CT1:7), digested with EcoRI and HindIII, and hybridized to a Tnt1-specific probe. An 879-bp fragment can be observed in each lane. The hybridization patterns associated with the selfed progenies are identical to the patterns of the parental lines. B, Southern-blot hybridization analysis of two DNA samples. Lane 1, DNA isolated from CT1:3; lane 2, pooled DNA from 19 progenies of CT1:3 and genomic DNA from CT1:4. Arrows point to the two bands associated with CT1:4. To confirm that the potentially retransposed Tnt1 element(s) is detectable in this pooled hybridization approach, we developed a blot by mixing an equal amount of DNA from each of 19 progenies from CT1:3 with DNA from CT1:4. The Tnt1 elements associated with CT1:4 can be unambiguously detected in the mixed DNA sample (Fig. 3B). Retrotransposition Efficiency Does Not Depend on the Number of Tnt1 Elements We were interested to know if the average number of transpositions during tissue culture can be controlled by using lines with different copy numbers of the Tnt1 element. We conducted regeneration experiments using four lines (CT1, CT1:4, CT1:3, and CT6), which contain one, three, eight, and nine Tnt1 copies, respectively. The Tnt1 copy numbers of 10 regenerated progeny from each of these four lines were determined by Southern-blot hybridization. Interestingly, we did not observe a correlation between initial Tnt1 copy number and the average Tnt1 copy number in the regenerated progenies (Table I). For example, CT1 contains a single Tnt1 element, while the 10 regenerated progenies from CT1 contained an average of 10.4 Tnt1 elements, resulting in an average of 9.4 new insertions. In comparison, CT6 contains nine copies of Tnt1, and the 10 progenies from CT6 contained an average of 13.5 Tnt1 elements, resulting in only an average of 4.5 new insertions. Tnt1 copy numbers in progeny derived from four start lines Table I. Tnt1 copy numbers in progeny derived from four start lines Progeny from CT1 . Total Tnt1 Copies . Progeny from CT6 . Total Tnt1 Copies . Progeny from CT1:3 . Total Tnt1 Copies . Progeny from CT1:4 . Total Tnt1 Copies . CT1:1 >20a CT6:5 11 CT1:3:1 8 CT1:4:2 15 CT1:3 9 CT6:7 16 CT1:3:7 >20 CT1:4:3 >20 CT1:4 3 CT6:9 9 CT1:3:5 14 CT1:4:4 8 CT1:5 2 CT6:11 16 CT1:3:13 12 CT1:4:5 7 CT1:6 2 CT6:15 13 CT1:3:12 10 CT1:4:8 8 CT1:7 10 CT6:17 9 CT1:3:11 8 CT1:4:12 >20 CT1:10 >20 CT6:22 15 CT1:3:10 8 CT1:4:13 12 CT1:12 16 CT6:23 9 CT1:3:6 >20 CT1:4:15 16 CT1:14 >20 CT6:24 18 CT1:3:4 11 CT1:4:16 8 CT1:16 2 CT6:29 19 CT1:3:2 >20 CT1:4:42 9 Averageb 10.4 13.5 13.1 12.3 Parental Tnt1 copies 1 9 8 3 New Tnt1 copies 9.4 4.5 5.1 9.3 Progeny from CT1 . Total Tnt1 Copies . Progeny from CT6 . Total Tnt1 Copies . Progeny from CT1:3 . Total Tnt1 Copies . Progeny from CT1:4 . Total Tnt1 Copies . CT1:1 >20a CT6:5 11 CT1:3:1 8 CT1:4:2 15 CT1:3 9 CT6:7 16 CT1:3:7 >20 CT1:4:3 >20 CT1:4 3 CT6:9 9 CT1:3:5 14 CT1:4:4 8 CT1:5 2 CT6:11 16 CT1:3:13 12 CT1:4:5 7 CT1:6 2 CT6:15 13 CT1:3:12 10 CT1:4:8 8 CT1:7 10 CT6:17 9 CT1:3:11 8 CT1:4:12 >20 CT1:10 >20 CT6:22 15 CT1:3:10 8 CT1:4:13 12 CT1:12 16 CT6:23 9 CT1:3:6 >20 CT1:4:15 16 CT1:14 >20 CT6:24 18 CT1:3:4 11 CT1:4:16 8 CT1:16 2 CT6:29 19 CT1:3:2 >20 CT1:4:42 9 Averageb 10.4 13.5 13.1 12.3 Parental Tnt1 copies 1 9 8 3 New Tnt1 copies 9.4 4.5 5.1 9.3 a If a plant contains more than 20 Tnt1 elements, the number cannot be accurately counted based on Southern-blot hybridization.  bFor average, 20 was used in the calculation if a plant contained more than 20 copies. Open in new tab Table I. Tnt1 copy numbers in progeny derived from four start lines Progeny from CT1 . Total Tnt1 Copies . Progeny from CT6 . Total Tnt1 Copies . Progeny from CT1:3 . Total Tnt1 Copies . Progeny from CT1:4 . Total Tnt1 Copies . CT1:1 >20a CT6:5 11 CT1:3:1 8 CT1:4:2 15 CT1:3 9 CT6:7 16 CT1:3:7 >20 CT1:4:3 >20 CT1:4 3 CT6:9 9 CT1:3:5 14 CT1:4:4 8 CT1:5 2 CT6:11 16 CT1:3:13 12 CT1:4:5 7 CT1:6 2 CT6:15 13 CT1:3:12 10 CT1:4:8 8 CT1:7 10 CT6:17 9 CT1:3:11 8 CT1:4:12 >20 CT1:10 >20 CT6:22 15 CT1:3:10 8 CT1:4:13 12 CT1:12 16 CT6:23 9 CT1:3:6 >20 CT1:4:15 16 CT1:14 >20 CT6:24 18 CT1:3:4 11 CT1:4:16 8 CT1:16 2 CT6:29 19 CT1:3:2 >20 CT1:4:42 9 Averageb 10.4 13.5 13.1 12.3 Parental Tnt1 copies 1 9 8 3 New Tnt1 copies 9.4 4.5 5.1 9.3 Progeny from CT1 . Total Tnt1 Copies . Progeny from CT6 . Total Tnt1 Copies . Progeny from CT1:3 . Total Tnt1 Copies . Progeny from CT1:4 . Total Tnt1 Copies . CT1:1 >20a CT6:5 11 CT1:3:1 8 CT1:4:2 15 CT1:3 9 CT6:7 16 CT1:3:7 >20 CT1:4:3 >20 CT1:4 3 CT6:9 9 CT1:3:5 14 CT1:4:4 8 CT1:5 2 CT6:11 16 CT1:3:13 12 CT1:4:5 7 CT1:6 2 CT6:15 13 CT1:3:12 10 CT1:4:8 8 CT1:7 10 CT6:17 9 CT1:3:11 8 CT1:4:12 >20 CT1:10 >20 CT6:22 15 CT1:3:10 8 CT1:4:13 12 CT1:12 16 CT6:23 9 CT1:3:6 >20 CT1:4:15 16 CT1:14 >20 CT6:24 18 CT1:3:4 11 CT1:4:16 8 CT1:16 2 CT6:29 19 CT1:3:2 >20 CT1:4:42 9 Averageb 10.4 13.5 13.1 12.3 Parental Tnt1 copies 1 9 8 3 New Tnt1 copies 9.4 4.5 5.1 9.3 a If a plant contains more than 20 Tnt1 elements, the number cannot be accurately counted based on Southern-blot hybridization.  bFor average, 20 was used in the calculation if a plant contained more than 20 copies. Open in new tab All progenies from CT1 (one Tnt1 copy) and CT1:4 (three Tnt1 copies) contained more Tnt1 copies than the parental lines, suggesting that retrotransposition occurred in every plant. In contrast, three plants from CT1:3 (eight Tnt1 copies) and three plants from CT6 (nine Tnt1 copies) contained the same numbers of Tnt1 as the parental lines. These results suggest that most of the Tnt1 elements in CT1:3 and CT6 were not activated by tissue culture. Thus, the retrotransposition efficiency of a line is likely dependent on how easily the Tnt1 element(s) can be activated rather than on the number of original Tnt1 elements. Tnt1 Preferentially Inserts into Genic Regions in the Potato Genome We developed a Tnt1-seq method to map the genomic positions of a large number of Tnt1 insertions (Fig. 4). Genomic DNA was isolated from 70 T0 and T1 lines. Tnt1-flanking DNA sequences were isolated by two rounds of DNA walking using PCR. The PCR products were then pooled and used for the preparation of a paired-end Tnt1-seq library (Fig. 4). To check the quality of the library, we cloned and sequenced 20 randomly selected DNA fragments from this library. All clones contained the expected left- and right-border adapters. A total of nine independent inserts were identified (some inserts were identical). This library was sequenced using the Illumina MiSeq platform, resulting in a total of 5,931,429 paired-end sequence reads (150 bp). The sequences from the adapter immediately adjacent to the partial Tnt1 sequence were found to contain sequence errors. Thus, only the sequences adjacent to the left adapter were used in mapping. We mapped 474,424 sequences to the DM1-3 reference genome (PGSC_DM_v3_2.1.11; Xu et al., 2011), corresponding to 1,667 insertion sites, including all nine inserts identified by manual cloning and sequencing. Thus, the 70 different T0 and T1 lines contain an average of 24 Tnt1 insertions. Mapping of the 1,667 insertion sites revealed a near-random distribution on all 12 potato chromosomes (Fig. 5). Figure 4. Open in new tabDownload slide Development of the Tnt1-seq library. DNA walking was performed on pooled genomic DNA from 70 T0 and T1 lines. Each of the Annealing Controlled Primer (ACP) primers (DW-ACP1, DW-ACP2, DW-ACP3, and DW-ACP4) and the TSP1 primer (for Tnt1-specific primer 1) were used to amplify the target region from pooled genomic DNAs in the first PCRs. The second PCR using the DW-ACP-N primer and the TSP2 primer was performed using the first PCR product as a template. The PCR products were then combined and subjected to DNA end blunting, adding adenine (A) to the 3′ end, and paired-end adaptor ligation. The third PCR was conducted to enrich the Tnt1-flanking sequences. Each DNA fragment in the Tnt1-seq library contains an adaptor-specific primer (black), DW-ACP-N primer (red), Tnt1-flanking region (white), Tnt1 sequence from amplification (green), synthesized Tnt1 sequence, which is part of the Tnt1 enrichment-specific primer (purple), and part of the Tnt1 enrichment-specific primer, which was used for Illumina sequencing (sky blue). Figure 4. Open in new tabDownload slide Development of the Tnt1-seq library. DNA walking was performed on pooled genomic DNA from 70 T0 and T1 lines. Each of the Annealing Controlled Primer (ACP) primers (DW-ACP1, DW-ACP2, DW-ACP3, and DW-ACP4) and the TSP1 primer (for Tnt1-specific primer 1) were used to amplify the target region from pooled genomic DNAs in the first PCRs. The second PCR using the DW-ACP-N primer and the TSP2 primer was performed using the first PCR product as a template. The PCR products were then combined and subjected to DNA end blunting, adding adenine (A) to the 3′ end, and paired-end adaptor ligation. The third PCR was conducted to enrich the Tnt1-flanking sequences. Each DNA fragment in the Tnt1-seq library contains an adaptor-specific primer (black), DW-ACP-N primer (red), Tnt1-flanking region (white), Tnt1 sequence from amplification (green), synthesized Tnt1 sequence, which is part of the Tnt1 enrichment-specific primer (purple), and part of the Tnt1 enrichment-specific primer, which was used for Illumina sequencing (sky blue). Figure 5. Open in new tabDownload slide Distribution of Tnt1 insertion sites in the DM1-3 genome (PGSC_DM_v3_2.1.11). Vertical lines above and below the line represent the positions of Tnt1 insertions in the forward and reverse strands of the DM1-3 reference genome. Thick horizontal bars associated with chromosomes 1, 2, 5, and 12 indicate large physical gaps in the pseudomolecules. Figure 5. Open in new tabDownload slide Distribution of Tnt1 insertion sites in the DM1-3 genome (PGSC_DM_v3_2.1.11). Vertical lines above and below the line represent the positions of Tnt1 insertions in the forward and reverse strands of the DM1-3 reference genome. Thick horizontal bars associated with chromosomes 1, 2, 5, and 12 indicate large physical gaps in the pseudomolecules. A total of 533 insertions (33%) were associated with genic regions in the DM1-3 reference genome, including 15% in exons, 13% in introns, 3% in the 3′ untranslated region, and 2% in the 5′ untranslated region. The genic sequences account for approximately 14% of the potato genome; thus, Tnt1 elements preferentially insert in genic regions. In comparison, 78.6% of the Tnt1-flanking sequences matched coding sequence in M. truncatula (Tadege et al., 2008). The difference in the frequencies of Tnt1 insertions into genic regions in the two species may be caused by the differences in the accuracy of annotations of the two genomes and the lengths of the Tnt1-flanking sequences generated for analysis. The DM1-3 genome was based on the cultivated diploid species S. tuberosum group phureja (Xu et al., 2011). The level of sequence divergence between S. tuberosum group phureja and S. chacoense is unknown. The sequence divergence between the two species may prevent accurate mapping of the relatively short Illumina sequence reads derived from S. chacoense to the DM1-3 genome. Preliminary Screening of Mutations Caused by Tnt1 Insertions To investigate the potential of Tnt1 as an insertional mutagen, we developed 38 families by selfing independent T0 and T1 lines. We sowed 30 seeds from each family and then grew 15 randomly selected plants in a greenhouse, along with progenies from wild type 523-3. The germination rate and all visible phenotypes were recorded. Plants from five families showed seven phenotypes that were unambiguously distinguishable from wild-type 523-3 (Table II; Fig. 6). The germination rate of wild-type 523-3 seeds was 90%. In contrast, seeds from the CT4:33 family showed a much lower germination rate of 60%. A Tnt1 element in CT4:33 possibly landed in a gene that is essential to embryo/seed development; thus, progeny homozygous for this insertion may not be viable. Extreme dwarf plants were observed in the CT4:20 family. These plants showed a severely stunted growth and died approximately 1 month after being transplanted into individual pots. Putative mutation phenotypes associated with families selfed from T0 and T1 lines Table II. Putative mutation phenotypes associated with families selfed from T0 and T1 lines Phenotype . Family . Number of Plants Showing the Phenotypea . Low germination rate CT4:33 60% germination rate Extreme dwarf and stunted growth CT4:20 5 Bushy, short internode CT4:50, CT4:33 4, 4 Small leaf CT1:13 3 Small and curling-up leaf CT41 3 Round and curling-up leaf CT4:20 3 Phenotype . Family . Number of Plants Showing the Phenotypea . Low germination rate CT4:33 60% germination rate Extreme dwarf and stunted growth CT4:20 5 Bushy, short internode CT4:50, CT4:33 4, 4 Small leaf CT1:13 3 Small and curling-up leaf CT41 3 Round and curling-up leaf CT4:20 3 a Number of plants from a total of 15 in each family. Open in new tab Table II. Putative mutation phenotypes associated with families selfed from T0 and T1 lines Phenotype . Family . Number of Plants Showing the Phenotypea . Low germination rate CT4:33 60% germination rate Extreme dwarf and stunted growth CT4:20 5 Bushy, short internode CT4:50, CT4:33 4, 4 Small leaf CT1:13 3 Small and curling-up leaf CT41 3 Round and curling-up leaf CT4:20 3 Phenotype . Family . Number of Plants Showing the Phenotypea . Low germination rate CT4:33 60% germination rate Extreme dwarf and stunted growth CT4:20 5 Bushy, short internode CT4:50, CT4:33 4, 4 Small leaf CT1:13 3 Small and curling-up leaf CT41 3 Round and curling-up leaf CT4:20 3 a Number of plants from a total of 15 in each family. Open in new tab Figure 6. Open in new tabDownload slide Visible phenotypes observed in selfed progeny from some T0 and T1 families compared with wild-type 523-3 plants. A, Short internodes of a plant from the CT4:50 family (left) and internodes from a wild-type 523-3 plant (right). B, Bushy phenotype of a plant from the CT4:33 family (left) and a wild-type 523-3 plant (right). C, A plant with small, inward-curling leaves from the CT41 family (left) and a wild-type 523-3 plant (right). D, Rounded, inward-curling leaves of plants from the CT4:20 family (left) and wild-type leaves (right). Figure 6. Open in new tabDownload slide Visible phenotypes observed in selfed progeny from some T0 and T1 families compared with wild-type 523-3 plants. A, Short internodes of a plant from the CT4:50 family (left) and internodes from a wild-type 523-3 plant (right). B, Bushy phenotype of a plant from the CT4:33 family (left) and a wild-type 523-3 plant (right). C, A plant with small, inward-curling leaves from the CT41 family (left) and a wild-type 523-3 plant (right). D, Rounded, inward-curling leaves of plants from the CT4:20 family (left) and wild-type leaves (right). Plants in several families showed distinctive leaf shapes. Plants in the CT4:20 family had large, round leaves that curled inward (Fig. 6D). Some plants from CT4:50 and CT4:33 had short internodes and, as a consequence, were bushy (Fig. 6, A and B). Small and inward-curling leaves were observed in plants from the CT41 family (Fig. 6C). DISCUSSION A mutation caused by a T-DNA or TE insertion into a gene will interrupt only one of the two homologs in diploid species. Selfing of the heterozygous mutation line is required to identify the homozygous recessive mutant. Therefore, insertional mutagenesis is most appropriately applied only in homozygous and self-compatible inbred lines. Most asexually propagated species, such as potato, contain highly heterozygous genomes and are often self-incompatible; thus, they cannot benefit from insertional mutagenesis technology. Ishizaki and Kato (2005) introduced the Tto1 retrotransposon from tobacco into a sterile diploid potato clone. However, transposition of Tto1 was not induced by tissue culture in potato (Ishizaki and Kato, 2005), although Tto1 can be activated by tissue culture in rice and Arabidopsis (Hirochika et al., 1996; Okamoto and Hirochika, 2000). An alternative approach in plant species with a heterozygous genome is to develop activation-tagging lines by transforming a construct containing a strong promoter (Weigel et al., 2000). Insertion of a strong promoter may result in an overexpression of an adjacent gene or a gain-of-function phenotype for the gene. Such a dominant phenotype can be observed in T0 transgenic lines, avoiding the selfing process. The Canadian Potato Genome Project generated 10,000 activation-tagged potato lines (Regan et al., 2006). However, the phenotype-inducing efficiency of the activation-tagging approach is not known in potato. In addition, each tagging line has to be individually transformed. Thus, the value of activation-tagging technology remains to be seen in potato. We have demonstrated that the Tnt1-based insertional mutagenesis system using S. chacoense clone 523-3 can be used to tag individual potato genes. This is because of the following reasons. (1) The 523-3 clone can be easily transformed and regenerated. Thus, many regenerated lines from a selected starter line can be readily developed, and it will be feasible to develop a large number of Tnt1 lines to saturate the entire potato genome (see below). (2) A sufficient amount of seeds from each regenerated line can be readily produced by self-pollination. Each plant can produce dozens of berries, each of which typically contains about 100 seeds. Potato seeds are small and can be stored in small-sized tubes or envelopes in a −20°C freezer for decades. (3) The 523-3 plants are much smaller than typical tetraploid potato cultivars and can be grown in small-sized pots for large-scale screening in greenhouses. (4) The 523-3 clone tuberizes well in both greenhouses and the field, even under long-photoperiod conditions (Kittipadukal et al., 2012). Thus, tuber-related traits can be screened. Tnt1-based genome-wide mutagenesis has already been well demonstrated in M. truncatula (Tadege et al., 2008; Cheng et al., 2011). The Tnt1 element in potato showed similar activity and behavior to that documented in other plant species, including activation by tissue culture, stability during sexual reproduction, and preferential transposition into genic regions (Courtial et al., 2001; d’Erfurth et al., 2003; Mazier et al., 2007; Cui et al., 2013). We examined several factors that may affect the Tnt1 retrotransposition. We observed that the Tnt1 copy numbers in the new regenerants do not correlate with the length of time that the callus was grown on the regeneration medium (data not shown). This is consistent with a similar study in M. truncatula (Tadege et al., 2008). The new Tnt1 copy numbers in regenerants were not correlated with the original Tnt1 copy numbers in the parental lines (Table I). It was demonstrated in Arabidopsis that the transcription of Tnt1 was often silenced in plants containing numerous copies. The silencing of Tnt1 was associated with 24-nucleotide short-interfering RNAs targeting the promoter localized in the long terminal repeat region and with the non-CG site methylation of these sequences (Pérez-Hormaeche et al., 2008). Our data are in agreement with those in Arabidopsis. Potato lines with more copies of Tnt1 generated fewer new Tnt1 copies during regeneration than CT1, a T0 line containing a single copy of Tnt1 (Table I). Thus, most Tnt1 copies in lines with numerous insertions are likely silenced and cannot be activated by tissue culture. Our preliminary mutation screening revealed the potential of the Tnt1-based system to generate potato mutants. Seven distinct phenotypic changes were observed in five of the 38 T0/T1 families. Each phenotype was observed in three to five plants in a particular family (Table II), indicating that each change was likely caused by a recessive mutation of a single gene. More subtle changes were observed in several additional families. In the screening of the M. truncatula mutant population, approximately 30% of the lines showed new recognizable phenotypes (Tadege et al., 2008). However, the screening involved inoculation with arbuscular mycorrhizal fungi and Sinorhizobium meliloti and was performed under low-nitrogen and low-phosphorus conditions. In this study, only a few families were screened, under regular greenhouse growing conditions, but the Tnt1-based system appears to have a similar phenotype-changing efficiency to that in M. truncatula. Our ultimate goal is to develop a 523-3 population that is large enough to tag every potato gene with a Tnt1 element. The average number of Tnt1 insertions will directly affect the number of Tnt1-tagged 523-3 lines needed for mutation saturation. The transposed Tnt1 copy numbers of the regenerants derived from CT1 ranged from one to more than 20, which is similar to the numbers observed in other plant species. The transposition events ranged from four to more than 30 in M. truncatula (d’Erfurth et al., 2003), zero to 26 in Arabidopsis (Courtial et al., 2001), and four to 19 in soybean (Cui et al., 2013). A total of 1,667 insertions was recovered from 70 Tnt1-harboring lines, averaging 24 insertions per line. The possibility of finding an insertion in a given gene can be estimated by P = 1 − (1 − [X/G])n, where P = the possibility of finding an insertion in a given gene, X = the length of the gene in kilobases, G = the genome size in kilobases, and n = the number of insertions in the population (Krysan et al., 1999). Considering that the size of the potato genome is 845 Mb and the average gene size is 2.5 kb, we estimate that approximately 1 million insertions are needed to achieve 95% genome saturation. If each Tnt1 tagging line contains 24 insertions, then 42,000 lines will be required. However, this equation assumes the random insertion of Tnt1. Therefore, the number of mutants needed to saturate the potato gene complement would be significantly reduced due to the preferential insertions of Tnt1 in genic regions. In a comparison, the genome size of M. truncatula is approximately 500 Mb, and approximately 90% of the M. truncatula genes were covered by a population of approximately 12,000 Tnt1 insertion lines (Cheng et al., 2011). MATERIALS AND METHODS Plant Material Solanum chacoense clone 523-3, developed by Robert Hanneman in Madison, Wisconsin (Hosaka and Hanneman, 1998a, 1998b), was used for transformation. For germination, seeds were soaked in 1,500 ppm GA3 for 24 h to break dormancy. The seeds were then sown in square pots (10 cm × 10 cm) using soilless potting mix. After 4 weeks, the seedlings were transplanted to individual pots and maintained in a greenhouse. Growth conditions were a 16-h photoperiod and 18°C and 15°C day and night temperatures, respectively. Potato Transformation and Regeneration Agrobacterium tumefaciens strain LBA4404 was transformed with the Tnk23 vector (Lucas et al., 1995). Internode explants from in vitro 523-3 plantlets were then transformed using the LBA4404 strain following standard A. tumefaciens-mediated potato (Solanum tuberosum) transformation (Bhaskar et al., 2008). The transformed shoots were transferred to rooting medium without antibiotics. In vitro regeneration was performed by cutting the internodes of each Tnt1-containing plant into explants approximately 5 mm long. The explants were then placed on medium used for transformation but without any antibiotics and transferred to new medium every 2 weeks. Regenerated shoots were transferred to rooting medium. Identification of Plants Containing Tnt1 Elements Genomic DNA was extracted from leaf tissue of transgenic plants grown in greenhouses using the Qiagen DNeasy Plant Mini Kit. The oligonucleotides KAN1 (5′-CCAACGCTATGTCCTGATAG-3′) and KAN2 (5′-TTTGTCAAGACCGACCTGTC-3′) were used to verify the presence of the nptII gene, and the oligonucleotides LTR1 (5′-ATGTCCATCTCATTGAAGAAGTA-3′) and LTR2 (5′-GGGAATAAACCCCTTACCAAAA-3′) were used to verify the presence of the Tnt1 element. PCR conditions for both primer pairs were 30 cycles of 94°C for 30 s, 55°C for 30 s, and 72°C for 1 min. For Southern-blot hybridization, approximately 20 μg of genomic DNA was digested with HindIII and EcoRI and then blotted on a membrane. The blots were probed with a 528-bp fragment complementary to nucleotides 141 to 669 of the nptII gene or with a 194-bp fragment complementary to nucleotides 1 to 194 of Tnt1. The blots were hybridized overnight at 65°C using standard protocols (Sambrook et al., 2001). Identification of the Tnt1-Flanking Region DNA walking on pooled genomic DNA from T0 and T1 lines was performed using the DNA walking SpeedUP Kit II (Seegene). Tnt1-specific primers used in the first and second PCR were TSP1 (5′-CCCGAGAGGAGCAACTGATA-3′) and TSP2 (5′-AAGAAATGAGAGTTGAAGCTCTCC-3′), respectively. The PCR products from the second round of DNA walking were used for the preparation of a paired-end Tnt1-seq library following the standard protocol provided by Illumina. Briefly, the protocol includes end blunting, adding an “A” tail, ligation of paired-end adaptors, and enrichment of the flanking region of Tnt1 insertions. The flanking sequences immediately adjacent to Tnt1 were amplified using the Tnt1 enrichment-specific primer 5′-AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACGCTCTTCCGATCTCACATGCCTAATACTTCTTCAATGAG-3′ and the adapter-specific primer 5′-CAAGCAGAAGACGGCATACGAGATCGGTCTCGGCATTCCTGCTGAACCGCTCTTCCGATCT-3′. The amplification program was as follows: 30 s at 98°C; 18 cycles of 10 s at 98°C, 30 s at 65°C, and 30 s at 72°C; and then 5 min at 72°C. DNA fragments between 300 and 550 bp were cut from a 2% agarose gel and purified using MinElute PCR purification columns (Qiagen). The quality of purified DNA (Tnt1-seq library) was checked by cloning into pCR4-TOPO TA vector (Invitrogen). Twenty clones were randomly selected and sequenced by Sanger sequencing. The Tnt1-seq library was then quantified using the Agilent Bioanalyzer 2100 and sequenced on the Illumina MiSeq platform. The sequence reads were mapped to the DM1-3 reference genome using BLAT (Kent, 2002). 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( 2011 ) From model to crop: functional analysis of a STAY-GREEN gene in the model legume Medicago truncatula and effective use of the gene for alfalfa improvement . Plant Physiol 157 : 1483 – 1496 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the National Science Foundation (grant no. IOS–1237969) and Hatch funds to J.J. and by the Strategic Scholarships for Frontier Research Network of Thailand’s Commission on Higher Education to S.D. * Address correspondence to [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Jiming Jiang ([email protected]). [OPEN] Articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.113.221903 © 2013 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2013. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Bajwa, Vikramjit S.; Wang, Xiaofeng; Blackburn, R. Kevin; Goshe, Michael B.; Mitra, Srijeet K.; Williams, Elisabeth L.; Bishop, Gerard J.; Krasnyanski, Sergei; Allen, George; Huber, Steven C.; Clouse, Steven D.

doi: 10.1104/pp.113.221465

Ring, Ludwig; Yeh, Su-Ying; Hücherig, Stephanie; Hoffmann, Thomas; Blanco-Portales, Rosario; Fouche, Mathieu; Villatoro, Carmen; Denoyes, Béatrice; Monfort, Amparo; Caballero, José Luis; Muñoz-Blanco, Juan; Gershenson, Jonathan; Schwab, Wilfried

Muthan, Bagyalakshmi; Roston, Rebecca L.; Froehlich, John E.; Benning, Christoph

doi: 10.1104/pp.113.222513pmid: 23839866

Abstract Diacylglycerol (DAG) is an intermediate in metabolism of both triacylglycerols and membrane lipids. Probing the steady-state pools of DAG and understanding how they contribute to the synthesis of different lipids is important when designing plants with altered lipid metabolism. However, traditional methods of assaying DAG pools are difficult, because its abundance is low and because fractionation of subcellular membranes affects DAG pools. To manipulate and probe DAG pools in an in vivo context, we generated multiple stable transgenic lines of Arabidopsis (Arabidopsis thaliana) that target an Escherichia coli DAG kinase (DAGK) to each leaflet of each chloroplast envelope membrane. E. coli DAGK is small, self inserts into membranes, and has catalytic activity on only one side of each membrane. By comparing whole-tissue lipid profiles between our lines, we show that each line has an individual pattern of DAG, phosphatidic acid, phosphatidylcholine, and triacylglycerol steady-state levels, which supports an individual function of DAG in each membrane leaflet. Furthermore, conversion of DAG in the leaflets facing the chloroplast intermembrane space by DAGK impairs plant growth. As a result of DAGK presence in the outer leaflet of the outer envelope membrane, phosphatidic acid accumulation is not observed, likely because it is either converted into other lipids or removed to other membranes. Finally, we use the outer envelope-targeted DAGK line as a tool to probe the accessibility of DAG generated in response to osmotic stress. Diacylglycerol (DAG) is a central metabolite in plant lipid metabolism. Its glycerol backbone is modified with two acyl chains. If a third acyl chain is added, triacylglycerol (TAG) is formed, whereas if a head group is added, it is converted into polar lipids such as a galactolipid. In green tissues, the majority of DAG is used as an intermediate in galactolipid synthesis, because the extensive thylakoid membrane system consists of approximately 85% galactolipids (Block et al., 1983). Although under normal conditions the galactolipids are exclusively chloroplastic, in Arabidopsis (Arabidopsis thaliana), the DAG used to make galactolipids is derived from assembly pathways in both the chloroplast and the endoplasmic reticulum (ER; Benning, 2009). In both pathways, the bulk of the fatty acids are synthesized in the chloroplast stroma (Browse et al., 1986) in the following order of abundance: 18:1, 16:0, and 18:0 (Wallis and Browse, 2002). In the chloroplast pathway, these fatty acids are directly attached to a glycerol-3-P, generating first lyso-phosphatidic acid (l-PtdOH) and then phosphatidic acid (PtdOH) in the inner leaflet of the chloroplast inner envelope (Fig. 1; Frentzen et al., 1983). The acyltransferases involved are specific to the extent that the sn-2 position of the glycerol backbone predominantly receives a 16:0 fatty acid. PtdOH is then used directly for phosphatidylglycerol (PtdGro) synthesis (Babiychuk et al., 2003) or converted to DAG by a PtdOH phosphatase (Joyard and Douce, 1977). The PtdOH phosphatase activity is known to be associated with the inner envelope, though which leaflet is obscured by the fact that DAG can efficiently flip across membranes (Hamilton et al., 1991) and the actual enzyme has not been unambiguously identified and located (Nakamura et al., 2007). However, the leaflet associations of two other enzymes that use DAG in the inner envelope have been established. MGD1, which uses DAG to synthesize the most abundant galactolipid, monogalactosyldiacylglycerol (MGDG), is on the outer leaflet of the inner envelope membrane (Xu et al., 2005), while SQD2, which uses DAG to generate the less abundant sulfolipid, sulfoquinovosyldiacylglycerol (SQDG), is located on the inner leaflet of the inner envelope membrane (Tietje and Heinz, 1998). Also associated with the inner envelope membrane are a number of fatty acid desaturases, including FAD4, FAD5, FAD6, FAD7, and FAD8 (Joyard et al., 2010). Two of these are specific, generating lipids with signature desaturations: FAD4 desaturates only the 16:0 fatty acid of PtdGro, giving plastidic PtdGro a distinct 16:1 Ɗ3 trans moiety (Browse et al., 1985; Gao et al., 2009), and FAD5 desaturates primarily the 16:0 fatty acid of MGDG, producing 16:1 Ɗ7 cis (Kunst et al., 1989). The remaining desaturases are less specific, with little preference for head group or acyl tail. They further desaturate 16:1 or 18:1 in the cis conformation to 16:2 or 18:2 (FAD6; Browse et al., 1989) and on to 16:3 or 18:3 (FAD7 and FAD8; Wallis and Browse, 2002). The combined actions of these FADs result in the highly desaturated fatty acid profiles seen for most chloroplast lipids. Figure 1. Open in new tabDownload slide Overview of DAG pools in the chloroplast envelope membranes. Processes that are known to have activity feeding into or withdrawing from DAG pools in the chloroplast envelope membranes are shown. Enzymes are indicated, and their substrates and products are connected with black arrows. However, for space reasons, not all reactants are shown. Membrane leaflets are indicated, and enzymes with known membrane topology are displayed correctly, while those without known topology are displayed in the center of the appropriate membrane. The acyl group preferred by each l-PtdOH acyltransferase is given in parentheses. Proposed processes transporting lipids from the ER to the chloroplast are shown with dashed arrows. Enzymes are as follows: 1, ATS1; 2, ATS2; 3, lipid phosphate phosphatase γ; 4, MGD1; 5, SQD2; 6, cytosolic phospholipases; 7, MGD2 or MGD3; 8, SFR2; 9, acyl-CoA:glycerol-3-P acyltransferase; 10, l-PtdOH acyltransferase; 11, PtdOH phosphatase; 12, cytidine diphosphate-choline:DAG cholinephosphotransferase; 13, TGD4; and 14, TGD1, TGD 2, TGD3 lipid transport complex. OE, Chloroplast outer envelope membrane; IE, chloroplast inner envelope membrane; ACP, acyl carrier protein. [See online article for color version of this figure.] Figure 1. Open in new tabDownload slide Overview of DAG pools in the chloroplast envelope membranes. Processes that are known to have activity feeding into or withdrawing from DAG pools in the chloroplast envelope membranes are shown. Enzymes are indicated, and their substrates and products are connected with black arrows. However, for space reasons, not all reactants are shown. Membrane leaflets are indicated, and enzymes with known membrane topology are displayed correctly, while those without known topology are displayed in the center of the appropriate membrane. The acyl group preferred by each l-PtdOH acyltransferase is given in parentheses. Proposed processes transporting lipids from the ER to the chloroplast are shown with dashed arrows. Enzymes are as follows: 1, ATS1; 2, ATS2; 3, lipid phosphate phosphatase γ; 4, MGD1; 5, SQD2; 6, cytosolic phospholipases; 7, MGD2 or MGD3; 8, SFR2; 9, acyl-CoA:glycerol-3-P acyltransferase; 10, l-PtdOH acyltransferase; 11, PtdOH phosphatase; 12, cytidine diphosphate-choline:DAG cholinephosphotransferase; 13, TGD4; and 14, TGD1, TGD 2, TGD3 lipid transport complex. OE, Chloroplast outer envelope membrane; IE, chloroplast inner envelope membrane; ACP, acyl carrier protein. [See online article for color version of this figure.] In unstressed plants, DAG seems to be used primarily in the inner chloroplast envelope. However, several conditions are known to cause extensive DAG use in the chloroplast outer envelope. During phosphate deprivation, MGD2 and MDG3 synthesize MGDG from DAG on the outer leaflet of the outer envelope membrane (Kobayashi et al., 2009). The DAG backbones are probably supplied from the phosphatidylcholine (PtdCho) pool by phospholipase activity, which was shown to be simultaneously up-regulated (Andersson et al., 2004; Nakamura et al., 2005). DAG is also generated during freezing stress by a galactolipid:galactolipid galactosyltransferase named Sensitive to FReezing2 (SFR2). This enzyme transfers the galactosyl head group of MGDG onto another MGDG, giving rise to digalactosyldiacylglycerol (DGDG) and DAG (Moellering et al., 2010). The DAG is subsequently sequestered into a lipid droplet by formation of TAG by an as yet unidentified enzyme. In the ER pathway, fatty acids synthesized in the chloroplast stroma are exported through a still poorly defined mechanism to the ER and activated to acyl-CoAs. Acyltransferases sequentially catalyze formation of l-PtdOH and PtdOH from glycerol-3-P and acyl-CoAs. Again, the acyltransferase working on the sn-2 position of the glycerol backbone is specific, but unlike the chloroplast isoform, it prefers an 18:1 carbon fatty acid (Frentzen et al., 1983). Newly generated PtdOH can be converted to PtdGro or phosphatidyl inositol (PtdIns) (Collin et al., 1999) or hydrolyzed to DAG (Shimojima et al., 2009). DAG can then be further metabolized to TAG and PtdCho. PtdCho acyl groups (18:1/18:1 and 18:1/16:0) are desaturated sequentially by desaturases FAD2 (Okuley et al., 1994) and FAD3 (Browse et al., 1993). These desaturases prefer PtdCho as substrate. The acyl chains modified on PtdCho are transferred to other ER lipids, including DAG, as a result of continual acyl editing of the PtdCho pool (Bates et al., 2012). Furthermore, PtdOH and many of the other extraplastidic phospholipids can be converted to DAG by action of phospholipases (Shimojima et al., 2009). These have as yet partially defined roles in response to stress or recycling of membrane lipids (Testerink and Munnik, 2005). Glycerolipid precursors generated by de novo synthesis, acyl editing, and possibly stress conditions in the ER are transported to the chloroplast by a mechanism that is likely to involve at least two putative lipid transporters: trigalactosyldiacylglycerol4 (TGD4) in the chloroplast outer envelope membrane and the TGD1, TGD2, and TGD3 complex in the inner envelope membrane (Wang and Benning, 2012). The actual lipid species transported remains unclear, but PtdCho, lyso-phosphatidylcholine, PtdOH, and DAG have been discussed in the literature (Andersson and Dörmann, 2009). The DAG moieties are then fully incorporated into all plastidic lipids except PtdGro, presumably using the same pathways that metabolize plastidic DAG, described above. Because of the preference of chloroplast and ER sn-2 acyltransferases for 16 or 18 carbon fatty acids, respectively, the origin of the DAG moieties can be distinguished by positional analysis of the acyl groups on the glycerol backbone (Roughan and Slack, 1982). In Arabidopsis, the chloroplast and ER lipid synthesis pathways contribute nearly equally to mature chloroplast lipids (Browse et al., 1986; Mongrand et al., 1998). Thus, the DAG pools described so far in the chloroplast inner and outer envelope membranes are each of dual origin. A challenge for the analysis of the different DAG pools is that this compound is not a bilayer-forming lipid and thus does not accumulate stably to high levels. Furthermore, during any lengthy fractionation procedure, its levels can be expected to alter, as DAG-modifying enzymes exist in multiple membranes. Moreover, because DAG is quickly metabolized and may have efficient transport systems (Dong et al., 2012), it is difficult to confirm whether metabolizing enzymes are accessing the same or separate DAG pools. To probe different DAG pools of chloroplast membranes in vivo, we have generated a series of stable transgenic Arabidopsis lines in which we target an Escherichia coli DAG kinase (DAGK) to each leaflet of the chloroplast envelope membranes. The basic utility of this approach was previously shown by targeting a DAGK to the chloroplast in tobacco (Nicotiana tabacum) using a single construct fusing the bacterial protein to the Rubisco small subunit N-terminal peptide (Fritz et al., 2007). Here, we present a full phenotypic analysis of these lines, determining which chloroplast membranes show steady-state alterations of DAG and PtdOH levels predicted by ectopic DAGK activity. We further determine the accessibility of DAG pools generated on the outer leaflet of the chloroplast outer envelope membrane during osmotic stress. Having this system established in Arabidopsis will allow characterization of DAG pools in multiple lipid mutant lines. RESULTS Construction and Verification of Targeted DAGK Fusion Proteins To investigate accessibility of DAG pools in each chloroplast membrane, we took advantage of a small, 121-amino acid DAGK from E. coli that spontaneously inserts itself into membranes (Sanders et al., 1996). DAGK converts DAG into PtdOH and was shown to be able to access DAG when inserted into the inside of the inner chloroplast envelope in tobacco (Fritz et al., 2007). DAGK fusion constructs were generated that incorporated a C-terminal hemagglutinin (HA) tag and did or did not include chloroplast-specific N-terminal targeting information, as diagrammed in Figure 2A. Figure 2. Open in new tabDownload slide DAGK fusion proteins remain functional. A, Construction of targeted DAGK fusion proteins. All proteins contain the full E. coli DAGK protein with HA tag. These are fused with N-DGD1, tpTOC75, and tpATS1. Numbers of the first and last residue of the targeting sequence or DAGK are given. B, Bacterial strain FB21625 (dgkA −)-producing proteins indicated below after overnight growth on agar-solidified medium containing isopropyl β-d-1-thiogalactopyranoside and antibiotics and with or without arbutin as indicated at right. (–) indicates spotting of the untransformed strain. PDV1 is a construct without a DAGK fusion in the same pDEST24 vector as the DAGK constructs and is used as a vector control. The vertical white bar separates different sections from the same plate. [See online article for color version of this figure.] Figure 2. Open in new tabDownload slide DAGK fusion proteins remain functional. A, Construction of targeted DAGK fusion proteins. All proteins contain the full E. coli DAGK protein with HA tag. These are fused with N-DGD1, tpTOC75, and tpATS1. Numbers of the first and last residue of the targeting sequence or DAGK are given. B, Bacterial strain FB21625 (dgkA −)-producing proteins indicated below after overnight growth on agar-solidified medium containing isopropyl β-d-1-thiogalactopyranoside and antibiotics and with or without arbutin as indicated at right. (–) indicates spotting of the untransformed strain. PDV1 is a construct without a DAGK fusion in the same pDEST24 vector as the DAGK constructs and is used as a vector control. The vertical white bar separates different sections from the same plate. [See online article for color version of this figure.] To ensure that fusion of targeting peptides and HA tags did not disrupt DAGK function, the constructs were introduced into an E. coli strain lacking DAGK (dgkA −). E. coli lacking DAGK are sensitive to growth on hydroquinone β-d-glucopyranoside (arbutin; Miller et al., 1992; Jerga et al., 2007). Arbutin is an alternative substrate for phosphoglycerol transferase I, which can transfer the head group of PtdGro to arbutin. When added to the media, arbutin essentially allows bulk conversion of PtdGro to DAG (Jackson and Kennedy, 1983). Accordingly, in dgkA − strains, DAG accumulates and prevents growth. However, dgkA − strains producing DAGK proteins with or without targeting peptides grew successfully on arbutin complementing the mutation, while the knockout strain producing a control vector did not (Fig. 2B). Therefore, all DAGK fusion proteins retained activity. E. coli DAGK inserts directionally into membranes and is only active on one face of the membrane (Sanders et al., 1996; Van Horn and Sanders, 2012). Therefore, to test all envelope membranes of the chloroplast, it was targeted to each leaflet of each chloroplast envelope membrane. As a control, untargeted DAGK was used. Because it has no targeting sequence, it is expected to integrate into any membrane accessible to the cytosol, including the chloroplast outer envelope (Fig. 3A). Figure 3. Open in new tabDownload slide DAGK fusion proteins are targeted to all compartments of the chloroplast. A, Expected locations of DAGK proteins based on targeting information and self insertion of DAGK into the nearest membranes. EPM, Extraplastidic membranes; OE, chloroplast outer envelope membrane; IMS, inter membrane space; IE, inner envelope membrane; Th, thylakoid membranes. B, Radiolabeled precursors of proteins as indicated at left were incubated with pea chloroplasts. After import, chloroplasts were incubated with or without Thermolysin or trypsin and were then reisolated and separated into membrane pellet (P) and soluble (S) fractions as indicated at top. Ten percent of the total precursor (TP) before chloroplast incubation is shown as a reference for precursor size. Locations of precursor (pr) and mature (m) proteins are indicated at right; asterisks indicate protease resistant protein fragments. A dashed line separates controls from DAGK fusion proteins. Results are representative of three repeats. [See online article for color version of this figure.] Figure 3. Open in new tabDownload slide DAGK fusion proteins are targeted to all compartments of the chloroplast. A, Expected locations of DAGK proteins based on targeting information and self insertion of DAGK into the nearest membranes. EPM, Extraplastidic membranes; OE, chloroplast outer envelope membrane; IMS, inter membrane space; IE, inner envelope membrane; Th, thylakoid membranes. B, Radiolabeled precursors of proteins as indicated at left were incubated with pea chloroplasts. After import, chloroplasts were incubated with or without Thermolysin or trypsin and were then reisolated and separated into membrane pellet (P) and soluble (S) fractions as indicated at top. Ten percent of the total precursor (TP) before chloroplast incubation is shown as a reference for precursor size. Locations of precursor (pr) and mature (m) proteins are indicated at right; asterisks indicate protease resistant protein fragments. A dashed line separates controls from DAGK fusion proteins. Results are representative of three repeats. [See online article for color version of this figure.] To specifically target DAGK to the outer leaflet of the outer envelope of the chloroplast, the N terminus of digalactosyl diacylglycerol1 (DGD1; N-DGD1) was used, as it was previously shown to be responsible for targeting of DGD1 and interacts directly with the outer envelope membrane (Froehlich et al., 2001). Because of its membrane interaction, DAGK targeted with N-DGD1 is expected to insert only in the chloroplast outer envelope and to face the cytosol (Fig. 3A). To target DAGK simultaneously to the inner leaflet of the outer envelope membrane and the outer leaflet of the inner envelope membrane, it was fused with the transit peptide of the translocon at the outer envelope membrane of chloroplasts, 75 kD (tpTOC75). TOC75 is unique among chloroplast outer envelope membrane proteins in that it has cleavable targeting information, which is removed in two parts (Inoue et al., 2005). The first part is a stroma targeting sequence (Inoue et al., 2001), and the second part, which is necessary to sort TOC75 to the outer envelope membrane, was hypothesized to keep TOC75 from traversing the inner membrane (Baldwin and Inoue, 2006). In a preliminary experiment, tpTOC75 was shown to target soluble GFP to the intermembrane space (R.L. Roston and K. Inoue, unpublished data). When targeted with tpTOC75, DAGK is expected to be able to self insert into inner and outer envelope membranes facing the intermembrane space (Fig. 3A). Targeting of E. coli DAGK to both the inner leaflet of the inner chloroplast membrane and the outer leaflet of the thylakoid membranes was already shown to be possible in tobacco using the stromal targeting transit peptide of the small subunit of Rubisco (SSU; Fritz et al., 2007). A similar approach with the stromal targeting transit peptide of acyltransferase1 (ATS1; tpATS1) was used here (Fig. 3A). In vitro chloroplast protein import and subsequent protease digestion and chloroplast fractionation was used to confirm the expected locations of targeted DAGK proteins. Thermolysin is a protease that digests proteins not protected by the outer envelope membrane (Cline et al., 1984), while Trypsin can digest proteins not protected by the inner envelope membrane of the chloroplast (Jackson et al., 1998). Untargeted DAGK was not recovered with any chloroplast fraction, similar to nonchloroplast control luciferase and unlike our prediction that it would insert into the chloroplast outer envelope (Fig. 3, A and B, compare lanes 1, 2, and 3). By contrast, all of the DAGK fusion proteins were recovered with the chloroplast membrane pellet after fractionation, showing that DAGK inserted itself into chloroplast membranes in each case (Fig. 3B, compare lanes 2 and 3). N-DGD1-DAGK was completely susceptible to Thermolysin treatment (Fig. 3B, compare lanes 2 and 4), similar to control outer envelope protein TOC34. This confirms its location on the outer leaflet of the outer envelope membrane. tpTOC75-DAGK was partially digested by Thermolysin, retaining both protein fragments and some full-length mature protein, in a pattern distinct from TOC34 and control inner envelope protein Accumulation and Replication of Chloroplasts6 (ARC6; Fig. 3B, compare lanes 2 and 4). Existence of tpTOC75-DAGK fragments is consistent with DAGK facing the inner leaflet of the outer envelope membrane, allowing only the small DAGK transmembrane region to be accessed by the protease. Note that no degradation products are seen in the N-DGD1-DAGK Thermolysin digestion, corroborating evidence that tpTOC75-DAGK is in a different position in the outer envelope than that of N-DGD1-DAGK. The mature-length tpTOC75-DAGK (lane 4) is completely protected from Thermolysin and therefore must be inside of the outer envelope membrane. Trypsin digestion of tpTOC75-DAGK does not show any protease-protected fragment (Fig. 3B, compare lanes 7 and 9), indicating that the Thermolysin-resistant regions of tpTOC75-DAGK are not protected by the inner envelope membrane. Therefore, tpTOC75-DAGK must be on the outer face of the inner envelope membrane as well as the inner face of the outer envelope membrane, as predicted. Mature tpATS1-DAGK was recovered with the membrane pellet and protected from both Thermolysin and Trypsin digestion (Fig. 3B), consistent with the protease-protected, stromal region of ARC6. It has potentially inserted into either the inner leaflet of the inner envelope or the outer leaflet of the thylakoid membranes. When DAGK was similarly targeted in tobacco with the transit peptide of SSU (Fritz et al., 2007), it was found to be inserted in both inner envelope and thylakoid membranes; therefore, this is the most likely conclusion here. Together with bacterial complementation, these data show that the DAGK fusion proteins are active and specifically targeted to relevant locations throughout the chloroplast. Phenotypes of Arabidopsis Producing DAGK Proteins The DAGK constructs were transferred into wild-type Arabidopsis and expressed under the control of the Cauliflower mosaic virus 35S promoter. For simplicity, from this point onward, Arabidopsis plants producing DAGK will be referred to by their targeting information only, i.e. tpATS1 for tpATS1-DAGK-producing Arabidopsis, N-DGD1 for N-DGD1-DAGK-producing Arabidopsis, etc. The majority of resulting plants did not show a growth phenotype, including multiple lines of untargeted DAGK, N-DGD1, and tpATS1 (representative examples, Fig. 4A). In all plant lines, the overall level and species of fatty acids were similar to the wild type (Fig. 4, B and C). Similarly, in all lines tested, the DAGK-encoding construct was expressed, because DAGK-encoding mRNA was detectable in all lines, as assayed by reverse transcription (RT)-PCR (Fig. 4D). Figure 4. Open in new tabDownload slide Phenotypic effects of disrupting chloroplastic DAG pools. A, Twelve-week-old wild-type (WT) Arabidopsis or lines producing DAGK or DAGK fused to the respective targeting peptides are shown. A vertical white bar indicates plants grown at a separate time. B, Levels of total fatty acids (FA) are given as micrograms per milligram of dry weight (DW) in the wild type or plants producing DAGK fused to the respective targeting peptides. C, Fatty acid profiles are given as mole percentage of total in the wild type or plants producing DAGK with the respective targeting peptide. Error bars are sds calculated from three or more biological replicates for all graphs. D, RT-PCR results showing levels of DAGK or ACT2 (At3g18780) mRNA in multiple independent lines of untargeted DAGK, N-DGD1, tpTOC75, or tpATS1 or an individual wild-type Arabidopsis. A PCR control (indicated with C) has no template DNA added. Plant lines from which lipid analyses are shown are indicated with an asterisk. Figure 4. Open in new tabDownload slide Phenotypic effects of disrupting chloroplastic DAG pools. A, Twelve-week-old wild-type (WT) Arabidopsis or lines producing DAGK or DAGK fused to the respective targeting peptides are shown. A vertical white bar indicates plants grown at a separate time. B, Levels of total fatty acids (FA) are given as micrograms per milligram of dry weight (DW) in the wild type or plants producing DAGK fused to the respective targeting peptides. C, Fatty acid profiles are given as mole percentage of total in the wild type or plants producing DAGK with the respective targeting peptide. Error bars are sds calculated from three or more biological replicates for all graphs. D, RT-PCR results showing levels of DAGK or ACT2 (At3g18780) mRNA in multiple independent lines of untargeted DAGK, N-DGD1, tpTOC75, or tpATS1 or an individual wild-type Arabidopsis. A PCR control (indicated with C) has no template DNA added. Plant lines from which lipid analyses are shown are indicated with an asterisk. In the previous tobacco study by Fritz et al. (2007), stromally targeted DAGK caused severe growth phenotypes, including loss of chlorophyll and reduced shoot size. Thus, the wild-type appearance of the tpATS1 and N-DGD1 lines was unexpected given that the respective DAGK transcripts were present and shown to encode functional enzyme in the bacterial complementation assay (Fig. 2B). Therefore, DAGK activity in N-DGD1 and tpATS1 chloroplasts was directly measured by incubating solubilized chloroplasts with [γ-32P]ATP and quantifying [32P]PtdOH produced. To establish DAGK assays for isolated chloroplasts, we first validated the assay conditions with commercially available E. coli DAGK. The DAGK was serially diluted and added to wild-type Arabidopsis chloroplasts under assay conditions following those used by Fritz et al. (2007; Fig. 5A). Radiolabel recovered in PtdOH indicated that the assay was robust and in a linear range below 100 ng of DAGK. Using these conditions, chloroplast-associated DAGK activity was then assayed in the wild type, tpATS1, and N-DGD1. Surprisingly, the major product of the assay was not PtdOH, but l-PtdOH, presumably from activity of a phospholipase A. Furthermore, tpATS1 lines appeared to have distinctly more phospholipase A activity than the wild type, as more radioactivity was found in l-PtdOH than in PtdOH in these chloroplasts (Fig. 5B). Increased phospholipase A activity in tpATS1 lines was confirmed by an additional test in which excess commercial E. coli DAGK was added to tpATS1 and wild-type chloroplasts before the assay. Because the exogenous DAGK activity added exceeds the endogenous levels, if phospholipase activity was identical, the relative labeling of PtdOH and l-PtdOH in wild-type and tpATS1 plastid preparations should be similar. However, it was observed that tpATS1chloroplasts had less label in PtdOH (45.1 ± 5.4%) and more in l-PtdOH (125 ± 6.1%) compared with wild-type chloroplasts, consistent with an increased turnover of PtdOH in tpATS1. To reduce the affect of increased phospholipase A activity on the accuracy of DAGK activity measurements, both labeled PtdOH and l-PtdOH were considered products. Assaying DAGK in this manner for both N-DGD1 and tpATS1 lines indicated that both N-DGD1 and tpATS1 have increased radioactivity in PtdOH and l-PtdOH compared with the wild type (Fig. 5C). However, statistical confidence levels were only above 95% for one of the two N-DGD1 lines (other line confidence level, 70.5%), and tpATS1 confidence levels were even lower (68.5% and 55.1%). A time course DAGK assay of the wild type and two independent tpATS1 lines showed a consistent increase of radiolabel in PtdOH and l-PtdOH in tpATS1 compared with the wild type (Fig. 5D). Considering the increase in phospholipase A activity observed in these lines (Fig. 5B), it seems reasonable to suggest that both N-DGD1 and tpATS1 lines have an active DAGK but that tpATS1 plants have compensated for this increase in DAGK activity by concurrently increasing activity of PtdOH-degrading enzymes, including that of at least one phospholipase A. Figure 5. Open in new tabDownload slide Targeted DAGK lines without physical phenotypes show increased DAGK activity. A, Incorporation of label into PtdOH produced by various amounts of E.coli DAGK added directly to wild-type Arabidopsis chloroplasts are presented as a percentage of total radioactivity. Data are fit with a Hill equation. B, Autoradiogram (left) or iodine staining (right) of lipids separated by TLC. On the left, lipids extracted from chloroplasts isolated from the wild type or plants producing tpATS1-DAGK and incubated with [γ-32P]ATP (DAGK assay). Identity of lipid standards generated by partial PtdOH digestion with R. arrhizus lipase are given at right. C, DAGK assays with chloroplasts isolated from the wild type or plants producing DAGK indicated by their respective targeting peptide with label in both PtdOH and l-PtdOH quantified together as percentage of total radioactivity. Results statistically different from the wild type at a 95% confidence level are indicated by an asterisk. D, Time course of l-PtdOH and PtdOH labeling during DAGK assay quantified together as percentage of total radioactivity using chloroplasts isolated from the wild type or two tpATS1-DAGK lines. Figure 5. Open in new tabDownload slide Targeted DAGK lines without physical phenotypes show increased DAGK activity. A, Incorporation of label into PtdOH produced by various amounts of E.coli DAGK added directly to wild-type Arabidopsis chloroplasts are presented as a percentage of total radioactivity. Data are fit with a Hill equation. B, Autoradiogram (left) or iodine staining (right) of lipids separated by TLC. On the left, lipids extracted from chloroplasts isolated from the wild type or plants producing tpATS1-DAGK and incubated with [γ-32P]ATP (DAGK assay). Identity of lipid standards generated by partial PtdOH digestion with R. arrhizus lipase are given at right. C, DAGK assays with chloroplasts isolated from the wild type or plants producing DAGK indicated by their respective targeting peptide with label in both PtdOH and l-PtdOH quantified together as percentage of total radioactivity. Results statistically different from the wild type at a 95% confidence level are indicated by an asterisk. D, Time course of l-PtdOH and PtdOH labeling during DAGK assay quantified together as percentage of total radioactivity using chloroplasts isolated from the wild type or two tpATS1-DAGK lines. Unlike N-DGD1 and tpATS1, lines of tpTOC75 did have growth phenotypes, including reduced growth (Fig. 4A), as well as delayed and reduced seed set. These phenotypes were unlikely to be due to the location of genomic insertion, as multiple independent lines showed similar phenotypes. Because tpTOC75 lines were the only ones showing growth phenotypes, the possibility that expression of the native TOC75 gene was sense suppressed was investigated. RT-PCR specific to TOC75 or to control gene ACTIN2 showed that TOC75 was expressed in all five tpTOC75 lines tested. In corroboration, Arabidopsis RNA interference knockdowns of TOC75 are severely pale (Huang et al., 2011), while tpTOC75 lines were not (Figs. 4A and 6, B and C). Instead, tpTOC75 lines grew more slowly than the wild type on soil (Fig. 4A) and plates (Fig. 6B), developing smaller mature rosettes, as indicated by rosette leaf size (Fig. 6B) and fewer bolts, but they had normal chlorophyll levels. Total chlorophyll level in wild-type plants was 2.19 ± 0.06 mg g–1 fresh weight, while in tpTOC75-1, it was 2.41 ± 0.33 and in tpTOC75-2, 2.10 ± 0.19. Based on our previous experience with chloroplast lipid mutants (Dörmann et al., 1995), the fact that chlorophyll levels remain unchanged indirectly suggests that there are no secondary effects on chloroplast lipid levels per se due to perturbation of photosynthesis. Therefore, we concluded that observed phenotypic effects were directly due to DAGK activity and proceeded with lipid analysis. Figure 6. Open in new tabDownload slide The strong phenotype of tpTOC75-DAGK-producing lines is not due to reduction of TOC75 expression. A, RT-PCR results showing levels of TOC75 (At3g46740) or ACT2 (At3g18780) mRNA in five independent tpTOC75-DAGK lines (1–5) or wild-type (WT) plants. The control (C) has no template DNA added. B, Four-week-old seedlings of the wild type or tpTOC75-DAGK grown on 1% Suc-supplemented Murashige and Skoog medium. C, Full rosette leaves of the 6-week-old wild type or tpTOC75-DAGK grown on soil. Figure 6. Open in new tabDownload slide The strong phenotype of tpTOC75-DAGK-producing lines is not due to reduction of TOC75 expression. A, RT-PCR results showing levels of TOC75 (At3g46740) or ACT2 (At3g18780) mRNA in five independent tpTOC75-DAGK lines (1–5) or wild-type (WT) plants. The control (C) has no template DNA added. B, Four-week-old seedlings of the wild type or tpTOC75-DAGK grown on 1% Suc-supplemented Murashige and Skoog medium. C, Full rosette leaves of the 6-week-old wild type or tpTOC75-DAGK grown on soil. Direct Effects on Levels of DAG and PtdOH Lipids were analyzed from leaves of 6-week-old, soil-grown wild-type Arabidopsis or two independent lines each of untargeted DAGK, N-DGD1, tpTOC75, or tpATS1, respectively. Because of the large number of lines and replicates involved, this work was necessarily completed over a period of time. Each experiment included, at a minimum, three biological replicates from the wild type and two DAGK-producing lines (i.e. the wild type, tpATS1-1, and tpATS1-2) and was repeated once or more with an additional set of separately grown plants to ensure accuracy. For graphical display, each data set was normalized to the wild-type samples analyzed within the same group and then displayed relative to the average of all the wild types in all groups. For statistical analyses of significant changes, each data set was again compared to wild-type samples analyzed in the same group. Because DAGK generates PtdOH from DAG, it was expected that the level of PtdOH would increase while the level of DAG would decrease in DAGK-producing plants. Untargeted DAGK lines had only a small effect on the levels of PtdOH and DAG, with the level of PtdOH appearing slightly reduced (Fig. 7A) but lacking statistical significance (best confidence level, 72%). DAG level changes were not consistent between the two lines tested (Fig. 7B) and are therefore unlikely to be relevant. As untargeted DAGK was shown not to interact with chloroplast membranes (Fig. 3B), it may not have inserted into any membrane or may have preferentially inserted into membranes that are not active in lipid metabolism generating DAG and thus had no effect. Similarly, tpATS1 lines showed no consistent change of DAG levels and an apparent change in PtdOH (Fig. 7, A and B), though again without strong statistical support (confidence level, 73%). Neither the tpATS1 nor the untargeted DAGK lines caused any changes to fatty acid profiles of PtdOH or DAG (Fig. 7, C and D). It should be noted that because fatty acid profiles are highly reproducible, many small changes were observed to be statistically significant (marked by asterisks). However, we have only considered the profile as “changed” if observed statistically significant changes are both consistent (i.e. present in both lines) and are compensated for by alternate statistically significant changes (e.g. 18:3 decreases are offset by 18:1 increases). The lack of observed changes in PtdOH and DAG fatty acid profiles of tpATS1 are likely because DAG in the inner chloroplast membrane is derived from PtdOH by action of a PtdOH phosphatase (Benning, 2009), the activity of which, like the observed increase of phospholipase A activity (Fig. 5B), may be increased in these lines. Thus, introducing a DAGK to the inner envelope likely causes completion of a futile cycle, reversibly converting DAG to PtdOH. The unchanged profiles only reconfirmed that the E. coli DAGK has no preference for specific DAG acyl chains (Walsh et al., 1990) and suggest that the native PtdOH phosphatase does not either. Figure 7. Open in new tabDownload slide PtdOH and DAG levels in DAGK-producing plants. Levels of PtdOH (A) or DAG (B) are given as a molar percentage of total fatty acids in the wild type or plants producing DAGK indicated by their respective targeting peptide. Fatty acid profiles of PtdOH (C) and DAG (D) in the wild type or plants producing DAGK indicated by their respective targeting peptide. Quantification shown is mole percentage of total fatty acids in each lipid. Error bars are sds from three or more biological replicates for all graphs. Levels statistically different from the wild type at a 95% confidence level are indicated by an asterisk. Those at a 99% confidence level are indicated by a double asterisk. Figure 7. Open in new tabDownload slide PtdOH and DAG levels in DAGK-producing plants. Levels of PtdOH (A) or DAG (B) are given as a molar percentage of total fatty acids in the wild type or plants producing DAGK indicated by their respective targeting peptide. Fatty acid profiles of PtdOH (C) and DAG (D) in the wild type or plants producing DAGK indicated by their respective targeting peptide. Quantification shown is mole percentage of total fatty acids in each lipid. Error bars are sds from three or more biological replicates for all graphs. Levels statistically different from the wild type at a 95% confidence level are indicated by an asterisk. Those at a 99% confidence level are indicated by a double asterisk. By contrast, chloroplast outer envelope-localized N-DGD1 lines showed significant decreases in both PtdOH and DAG levels (Fig. 7, A and B). Presumably, DAGK activity did cause a conversion of DAG to PtdOH, because a reduction in the level of DAG was observed. However, the expected increase in PtdOH levels must have been transient, and PtdOH must have been further converted and/or transferred to another membrane. The fatty acid profiles of N-DGD1 DAG were relatively unchanged (Fig. 7D), consistent with the nonspecific activity of E. coli DAGK. However, the fatty acid profile of N-DGD1 PtdOH showed an increase in 18:1 and a decrease in 18:3 moieties (Fig. 7C), suggestive of specificity not attributable to DAGK, and its likely source is the enzyme or transporter resulting in steady-state reduction of PtdOH levels. Alternatively, it is also possible that flux through the PtdOH pool is increased in N-DGD1 plants, and the increase in 18:1 and decrease in 18:3 represents a higher ratio of newly synthesized PtdOH. In tpTOC75 lines, where DAGK is targeted to both the inside of the outer envelope membrane and the outside of the inner envelope membrane (Fig. 3), the expected increase in PtdOH level and decrease in DAG level was finally observed (Fig. 7, A and B). The fatty acid profiles of each were similar to the wild type (Fig. 7, C and D), indicating that although PtdOH accumulated, it did so with acyl groups of normal desaturation levels, possibly because chloroplast desaturases are available in the inner envelope membrane (Joyard et al., 2010). Indirect Effects on Other Lipids PtdOH is a direct precursor for two abundant phospholipids, PtdGro and PtdCho (Benning, 2009), and therefore their levels are of interest. Furthermore, PtdGro levels and fatty acid composition were shown to be affected in tobacco producing tpSSU-DAGK (Fritz et al., 2007). In this study, Arabidopsis producing similarly targeted tpATS1-DAGK did not show any changes to the level of PtdGro, which was indistinguishable from the wild type (Fig. 8A). In fact, no lines showed changes to the level of PtdGro (Fig. 8A). There were, however, statistically significant changes to the fatty acid profile of PtdGro: tpATS1 showed decreases in 16:3 and increases in 18:2 (Fig. 8D), which could be consistent with eukaryotic pathway-derived PtdOH backbones becoming incorporated into plastidic PtdGro. If so, this would confirm the conclusion of the earlier study. Other DAGK lines did not show consistent changes to PtdGro profiles (Fig. 8D). Figure 8. Open in new tabDownload slide Effects on PtdGro, PtdCho, and TAG levels in DAGK-producing plants. Levels of PtdGro (A), PtdCho (B), and TAG (C) are given as a mole percentage of total fatty acids in the wild type or plants producing DAGK indicated by their respective targeting peptides. Fatty acid profiles of PtdGro (D), PtdCho (E), and TAG (F) are given as the mole percentage of total fatty acids in each lipid. Error bars are sds from three or more biological replicates for all graphs. Levels statistically different from the wild type at a 95% confidence level are indicated by an asterisk. Those at a 99% confidence level are indicated by a double asterisk. Figure 8. Open in new tabDownload slide Effects on PtdGro, PtdCho, and TAG levels in DAGK-producing plants. Levels of PtdGro (A), PtdCho (B), and TAG (C) are given as a mole percentage of total fatty acids in the wild type or plants producing DAGK indicated by their respective targeting peptides. Fatty acid profiles of PtdGro (D), PtdCho (E), and TAG (F) are given as the mole percentage of total fatty acids in each lipid. Error bars are sds from three or more biological replicates for all graphs. Levels statistically different from the wild type at a 95% confidence level are indicated by an asterisk. Those at a 99% confidence level are indicated by a double asterisk. In N-DGD1 lines, it was observed that PtdOH levels decreased by approximately 2% of total fatty acids (Fig. 7A) and that DAG levels decreased by approximately 0.1% (Fig. 7B) but that total fatty acids were not decreased (Fig. 4B). Therefore, these reductions must be compensated by increases in other lipid species. Accordingly, observed levels of PtdCho increased by approximately 2% (Fig. 8B) and TAG by approximately 0.15% (Fig. 8C). The fatty acid profile of PtdCho and TAG did not change to resemble that of PtdOH or DAG (compare Fig. 7, C and D to Fig. 8, E and F), suggesting that if a direct substrate-product relationship was in place, normal desaturation of product lipids was maintained. Levels and fatty acids of PtdCho and TAG were similar to the wild type for all other DAGK lines (Fig. 8, B, C, E, and F). Levels and compositions of other major plant lipids, including MGDG, DGDG, SQDG, PtdIns, and phosphatidylethanolamine were also quantified (Supplemental Fig. S1). However, no statistically relevant deviations from the wild type were observed among the DAGK-producing plants. Use of N-DGD1-DAGK to Probe DAG Pools Generated by SFR2 SFR2 is a processive galactosyltransferase also known as galactolipid:galactolipid galactosyltransferase, which is activated by freezing conditions or leaf infiltration with osmotically active compounds (Moellering et al., 2010). When activated, it removes a galactosyl moiety from MGDG and transfers it to another MGDG or higher order oligogalactolipid, increasing the length of the product’s head group and producing DAG as a product. The bulk of DAG formed this way is further metabolized to TAG. However, the availability of SFR2-generated DAG to alternate pathways is unknown. Hence, SFR2-generated DAG accessibility was tested by comparing PtdOH, DAG, and TAG levels before and after SFR2 activation in N-DGD1 or wild-type Arabidopsis. Before SFR2 activation, levels of PtdOH and DAG were reduced, while TAG levels were increased in N-DGD1 compared with the wild type (Figs. 7, A and B, 8C, and 9A). After SFR2 activation by MgCl2 leaf infiltration, PtdOH, DAG, and TAG levels of the wild type and N-DGD1 were similar (Fig. 9B). It should be noted that the small steady-state increase in TAG levels (approximately 0.1%) or decrease in DAG levels (approximately 0.05%) before SFR2 activation may be too small to be discernible after SFR2 activation, when TAG levels are approximately 12% and DAG approximately 1.4% of total fatty acids. Instead, the focus should be on the levels of PtdOH, which remain relatively similar before and after SFR2 activation in the wild type but not in N-DGD1. The relative increase in N-DGD1 PtdOH levels after SFR2 activation is consistent with DAGK accessing the DAG pool generated by SFR2 activity. Interestingly, although fatty acid profiles of PtdOH, DAG, and TAG after SFR2 activation were dramatically different than those before SFR2 activation (compare Fig. 7, C, D, and F to Fig. 9, C, D, and E), no profiles showed significant differences between the wild type and N-DGD1. Specifically, DAG and TAG profiles after activation have increased levels of 16:3 fatty acids (compare Fig. 9D to Fig. 7D and Fig. 9E to Fig. 8D), likely because of SFR2’s ability to include MGDG-derived backbones in these pools. By contrast, levels of 16:3 fatty acids of PtdOH are only slightly increased relative to the uninduced condition (compare Fig. 9C to Fig. 7C), indicating these backbones are not stably accumulating in the PtdOH pool. These observations are consistent with the ability of N-DGD1-DAGK to access the pool of MGDG-derived DAG created by SFR2 activity but also with the previous observation that N-DGD1-DAGK activity does not cause a stable accumulation of PtdOH (Fig. 7, A and C). It is possible that the 16:3 fatty acid-PtdOH pool was further metabolized into other lipids, including PtdCho and TAG, which were observed to be increased in N-DGD1-DAGK plants (Fig. 8). Figure 9. Open in new tabDownload slide DAG produced by SFR2 is accessible to DAGK on the chloroplast outer envelope membrane. Levels of PtdOH, DAG, and TAG are given as a mole percentage of total fatty acids in the wild type or N-DGD1-DAGK-producing Arabidopsis before (A) or after (B) overnight treatment with 0.4 m MgCl2. The before treatment is reproduced from Figures 6 and 7 for easy comparison. Fatty acid profiles for PtdOH (C), DAG (D), and TAG (E) of treated plants are given as mole percentage of total fatty acids in each lipid. Error bars are sds from three or more biological replicates for all graphs. Levels statistically different from the wild type at a 95% confidence level are indicated by an asterisk. Those at a 99% confidence level are indicated by a double asterisk. Figure 9. Open in new tabDownload slide DAG produced by SFR2 is accessible to DAGK on the chloroplast outer envelope membrane. Levels of PtdOH, DAG, and TAG are given as a mole percentage of total fatty acids in the wild type or N-DGD1-DAGK-producing Arabidopsis before (A) or after (B) overnight treatment with 0.4 m MgCl2. The before treatment is reproduced from Figures 6 and 7 for easy comparison. Fatty acid profiles for PtdOH (C), DAG (D), and TAG (E) of treated plants are given as mole percentage of total fatty acids in each lipid. Error bars are sds from three or more biological replicates for all graphs. Levels statistically different from the wild type at a 95% confidence level are indicated by an asterisk. Those at a 99% confidence level are indicated by a double asterisk. DISCUSSION Because DAG is a low-abundance, rapidly metabolized intermediate, the role of different DAG pools in membranes has traditionally been difficult to determine in vivo. Here, we have described the generation of stable transgenic Arabidopsis lines in which DAGK was targeted to each lamella of the chloroplast envelopes. These lines represent tools in a strategy to investigate accessibility of different DAG pools in vivo. Analysis of steady-state lipid accumulation in DAGK-producing lines revealed a distinct pattern of PtdOH, DAG, TAG, and PtdCho levels in each line, which supports the hypothesis that DAG has individual roles in each membrane leaflet. Furthermore, comparison of the lines showed that PtdOH accumulation is apparently prevented in the outer leaflet of the outer envelope membrane as PtdOH is either rapidly converted to other lipids or removed to other membranes, while DAG reduction or redirection into other lipids in the inner leaflet of the inner envelope membrane is detrimental to plant growth. Usefulness of the lines was demonstrated by showing that N-DGD1-DAGK can access DAG pools generated by SFR2. In the outer envelope-targeted N-DGD1 lines, an unexpected lipid phenotype was observed: a significant reduction in PtdOH and DAG with coincident increases in TAG and PtdCho levels (Figs. 6 and 7). The implication is that the outer lamella of the chloroplast outer envelope membrane has mechanisms to convert PtdOH into other lipids and avoid accumulation of PtdOH; however, the enzymes involved are currently not defined. There are no known pathways for TAG or PtdCho synthesis from PtdOH in the chloroplast outer envelope membrane (Inoue, 2007; Joyard et al., 2010). Therefore, PtdCho and TAG are more likely synthesized in a different membrane system, perhaps directed by a transient increase in PtdOH at the outer plastid envelope membrane. Because PtdOH can be a precursor for both PtdCho and TAG production in the ER (Benning, 2009), it is tempting to hypothesize that the relevant metabolism occurs there. In that case, PtdOH would have to be transported to the ER. It was recently shown that TGD4, a protein implicated in lipid trafficking between the ER and chloroplast, specifically binds PtdOH (Wang et al., 2012). TGD4 activity does not depend on external energy sources and is therefore likely to direct PtdOH transport by principles of mass action. Reverse transport of PtdOH by TGD4 could therefore explain why PtdOH is not accumulated, but it would fail to explain why PtdOH is reduced. On the other hand, PtdOH is known to be a signaling lipid, and PtdOH signals are known to be transient and quickly dissipated (Testerink and Munnik, 2005). It is certainly possible that PtdOH has a signaling role when generated on the outside of the chloroplast, as it would be exposed to cytosolic enzymes. However, it is not clear what the direct target(s) of the signaling PtdOH would be. We briefly considered the possibility that it would activate SFR2, as phospholipase activity and SFR2 activity both occur in response to freezing (Welti et al., 2002; Moellering et al., 2010). However, no accumulation of diagnostic oligogalactolipids was observed. Another unexpected result was the growth phenotype of the tpTOC75 lines. Unlike the N-DGD1 and tpATS1 lines that were similar to the wild type, the tpTOC75 lines were dwarfed (Figs. 3A and 4). It seems unlikely that this is an effect on the endogenous TOC75 gene, as it is still expressed (Fig. 6A), and the plants are not pale (Figs. 3A and 6, B and C), as is typically observed for TOC75-attenuated lines. Therefore, it is most likely to be related to the presence of DAGK on the outside of the chloroplast inner and/or the inside of the chloroplast outer envelope membranes. Because the rates of DAG flip-flop across a membrane (approximately 50 s–1; Allan et al., 1978; Hamilton et al., 1991) and catalysis by DAGK (kcat, turnover number, approximately 26 s–1; Van Horn and Sanders, 2012) are similar and the flip-flop rate of PtdOH (approximately 0.02 h–1; Homan and Pownall, 1988) is much slower, it is possible that the tpTOC75 lines are pulling DAG to the intermembrane space side of the envelopes. If DAG was not present in the outside lamella of the inner envelope membrane, further metabolism to MGDG would be impaired and could cause the slow growth phenotype. However, mutants deficient in MGDG, such as mgd1, are pale (Jarvis et al., 2000), and tpTOC75 lines are not. Another possibility is the completion of a futile cycle associated with the inner envelope membrane. One has to assume that tpTOC75-DAGK is making PtdOH on the outside of the inner envelope membrane. Because the TGD1, TGD2, TGD3 complex in the chloroplast inner envelope is an ATP-dependent lipid transporter in the inner envelope membrane that specifically binds PtdOH (Lu et al., 2007; Lu and Benning, 2009; Roston et al., 2012), it could use ATP to return the PtdOH produced by DAGK to the inner leaflet of the inner envelope membrane. There, plastid PtdOH phosphatase hydrolyzes PtdOH to DAG, which can easily flip back through the inner envelope membrane and become the substrate for DAGK, thereby completing the futile cycle. This futile cycle effectively depletes the plant of energy resources and therefore could cause the slower growth phenotype seen. However, the PtdOH phosphatase portion of this futile cycle should also have been present in tpATS1, which shows no phenotype. Finally, the answer may be as simple as the fact that tpTOC75-DAGK is similar to a combination of DAGK targeted in tpATS1- and N-DGD1-DAGK lines. Because DAG can easily flip across the membranes, we may be observing an additive effect of DAGK acting on both chloroplast envelopes. Interestingly, both the visual and lipid phenotypes of stromally targeted tpATS1-DAGK-producing Arabidopsis differ from previously reported stromally targeted tpSSU-DAGK-producing tobacco (Fritz et al., 2007). Transformed tobacco plants had severely reduced growth, were pale, and showed alterations in levels and/or composition of thylakoid MGDG, DGDG, SQDG, PtdOH, and PtdGro. Specifically, the study focused on unique molecular species of PtdGro with 18:1 and 18:4 Ɗ3-trans acyl groups in the sn-2 position of the glycerol backbone. In the Arabidopsis tpATS1-DAGK lines discussed here, there was no altered growth phenotype and no chlorotic phenotype, and while fatty acid profiles of total PtdGro were closely examined for unique species, none were found. Because levels of 18:1 and 18:4 Ɗ3-trans were very small in the tobacco study and the increase in DAGK activity levels were not large enough for statistical significance in the tpATS1-DAGK lines, it seems likely that any increases in 18:1 from our whole-leaf PtdGro extracts were below the detection limit. However, our PtdGro profiles did show consistently reduced 16:3 and increased 18:2 acyl groups, which could indicate that some chloroplast PtdGro was derived from eukaryotic PtdOH precursors. Because of the large differences in the data sets, better comparisons can be made of lipids extracted from whole leaves, as these experimental conditions were more similar. At the whole-leaf level, tpSSU-DAGK production resulted in a relative increase of PtdCho levels and relative decreases of MGDG and DGDG levels. These changes are likely to be indirectly related to the paleness and loss of photosynthetic capacity of transgenic tobacco, as chloroplast-specific lipids are reduced while the major extraplastidic phospholipid is increased. By contrast, the corresponding transgenic Arabidopsis lines were not pale, indicating that their chloroplasts and thylakoid membranes were functional, and PtdCho, MGDG, and DGDG levels seemed to reflect that by remaining constant (Fig. 7B; Supplemental Fig. S1). Again, it is unclear why photosynthesis was affected in tobacco and not in Arabidopsis. One possibility is that there is an intrinsic difference in lipid metabolism between the two species, a possibility that is corroborated by increased phospholipase A activity observed for the Arabidopsis transgenic lines and not the tobacco lines. It is a possibility that additional metabolic activities involved in PtdOH turnover have been activated in Arabidopsis expressing tpATS1-DAGK but not in tobacco transgenic lines. Conversely, it may be equally possible that the absolute difference in DAGK activity between the two experimental systems could explain the difference in phenotypes. CONCLUSION In conclusion, the chloroplast-targeted DAGK lines generated in this study revealed multiple effects that are not necessarily explained by our current understanding of lipid synthesis in the chloroplast envelope membranes. Therefore, the lines are not only informative in the short term, but will also be valuable tools for understanding lipid metabolism in the future. Their usefulness was demonstrated by showing that DAG generated during activation of SFR2 is accessible in the membrane. MATERIALS AND METHODS DAGK Constructs All relevant primers are given in Table I and are therefore referenced in the text by number. All kits were used according to manufacturer protocols. The DAGK coding sequence was isolated from Escherichia coli genomic DNA using primers 1 and 2. Primer 1 included restriction sites for BglII and KpnI, and primer 2 included a HA tag. PCR products of DAGK-HA were inserted into Gateway vector pDONR-Zeo (Invitrogen) using BP clonase II (Invitrogen), and sequences were confirmed at the Michigan State University Research Technology Support Facility (http://rtsf.msu.edu/). Targeting sequences were isolated from complementary DNA (cDNA) produced using SuperScript III (Invitrogen) from total RNA purified from Arabidopsis (Arabidopsis thaliana) ecotype Columbia-2 with an RNeasy plant mini kit (Qiagen). Primers were as follows: N-DGD1, 3 and 4; tpTOC75, 5 and 6; and tpATS1, 7 and 8. All forward primers included a BglII site, and all reverse primers included a KpnI site. PCR products were cut with BglII and KpnI (New England Biolabs) and ligated into DAGK-HA-pDONR-Zeo, and the entire construct was confirmed by sequencing. They were then transferred to pMDC32 (Curtis and Grossniklaus, 2003) for plant transformation or pDEST24 (Invitrogen) for bacterial complementation using LR clonase II (Invitrogen). For chloroplast protein import, entire DAG constructs (i.e. N-DGD1-DAGK-HA) were inserted into pGEMTEASY (Promega) using forward primers 9, 10, 11, and 12 and reverse primer 13 and confirmed by sequencing to be properly inserted behind the Sp6 promoter. Construction of N-DGD1-DAGK-HA was challenging and required the use of the CopyCutter EPI400 bacterial host (Epicentre) to minimize expression. Primer sequences Table I. Primer sequences Primers used for cloning or RT-PCR experiments are listed in the 5′ to 3′ direction. They are identified in the text by the number assigned at left. No. . 5′ to 3′ Sequence . 1 AAAAAGCAGGCTAGATCTACTGGTACCATGGCCAATAATACCACTGG 2 AGAAAGCTGGGTCCTAAGCGTAATCTGGAACATCGTATGGGTATCCAAAATGCGACCATAAC 3 GGCAGATCTATGGTAAAGGAAACTCTAA 4 TACGGTACCCTCAGGCTTCACAAA 5 GGCAGATCTATGGCCGCCTTCTC 6 TACGGTACCAGAATCCCAATCCGG 7 GGCAGATCTATGACTCTCACGTTTTC 8 TACGGTACCATCCTGAACAAGCTC 9 TATGCCACTCATATGATGGCCAATAATACCACTGG 10 TATGCCACTCATATGGTAAAGGAAACTCTAATTCCTCC 11 TATGCCACTCATATGATGGCCGCCTTCTCC 12 TATGCCACTCATATGACTCTCACGTTTTCCTC 13 ACAGCAGCATGCCTAAGCGTAATCTGGAACAT 14 CGTATCTGGATGGTGTTTACAATC 15 GGAATTCTTAATACCTCTCTCCAAATCGGAAGAAC 16 ATGGCCAATAATACCACTGG 17 TCCAAAATGCGACCATAAC 18 ATGGCTGAGGCTGATGATATTCAACCAATC 19 AGGAGCAATACGAAGCTCATTGTAGAAAGT No. . 5′ to 3′ Sequence . 1 AAAAAGCAGGCTAGATCTACTGGTACCATGGCCAATAATACCACTGG 2 AGAAAGCTGGGTCCTAAGCGTAATCTGGAACATCGTATGGGTATCCAAAATGCGACCATAAC 3 GGCAGATCTATGGTAAAGGAAACTCTAA 4 TACGGTACCCTCAGGCTTCACAAA 5 GGCAGATCTATGGCCGCCTTCTC 6 TACGGTACCAGAATCCCAATCCGG 7 GGCAGATCTATGACTCTCACGTTTTC 8 TACGGTACCATCCTGAACAAGCTC 9 TATGCCACTCATATGATGGCCAATAATACCACTGG 10 TATGCCACTCATATGGTAAAGGAAACTCTAATTCCTCC 11 TATGCCACTCATATGATGGCCGCCTTCTCC 12 TATGCCACTCATATGACTCTCACGTTTTCCTC 13 ACAGCAGCATGCCTAAGCGTAATCTGGAACAT 14 CGTATCTGGATGGTGTTTACAATC 15 GGAATTCTTAATACCTCTCTCCAAATCGGAAGAAC 16 ATGGCCAATAATACCACTGG 17 TCCAAAATGCGACCATAAC 18 ATGGCTGAGGCTGATGATATTCAACCAATC 19 AGGAGCAATACGAAGCTCATTGTAGAAAGT Open in new tab Table I. Primer sequences Primers used for cloning or RT-PCR experiments are listed in the 5′ to 3′ direction. They are identified in the text by the number assigned at left. No. . 5′ to 3′ Sequence . 1 AAAAAGCAGGCTAGATCTACTGGTACCATGGCCAATAATACCACTGG 2 AGAAAGCTGGGTCCTAAGCGTAATCTGGAACATCGTATGGGTATCCAAAATGCGACCATAAC 3 GGCAGATCTATGGTAAAGGAAACTCTAA 4 TACGGTACCCTCAGGCTTCACAAA 5 GGCAGATCTATGGCCGCCTTCTC 6 TACGGTACCAGAATCCCAATCCGG 7 GGCAGATCTATGACTCTCACGTTTTC 8 TACGGTACCATCCTGAACAAGCTC 9 TATGCCACTCATATGATGGCCAATAATACCACTGG 10 TATGCCACTCATATGGTAAAGGAAACTCTAATTCCTCC 11 TATGCCACTCATATGATGGCCGCCTTCTCC 12 TATGCCACTCATATGACTCTCACGTTTTCCTC 13 ACAGCAGCATGCCTAAGCGTAATCTGGAACAT 14 CGTATCTGGATGGTGTTTACAATC 15 GGAATTCTTAATACCTCTCTCCAAATCGGAAGAAC 16 ATGGCCAATAATACCACTGG 17 TCCAAAATGCGACCATAAC 18 ATGGCTGAGGCTGATGATATTCAACCAATC 19 AGGAGCAATACGAAGCTCATTGTAGAAAGT No. . 5′ to 3′ Sequence . 1 AAAAAGCAGGCTAGATCTACTGGTACCATGGCCAATAATACCACTGG 2 AGAAAGCTGGGTCCTAAGCGTAATCTGGAACATCGTATGGGTATCCAAAATGCGACCATAAC 3 GGCAGATCTATGGTAAAGGAAACTCTAA 4 TACGGTACCCTCAGGCTTCACAAA 5 GGCAGATCTATGGCCGCCTTCTC 6 TACGGTACCAGAATCCCAATCCGG 7 GGCAGATCTATGACTCTCACGTTTTC 8 TACGGTACCATCCTGAACAAGCTC 9 TATGCCACTCATATGATGGCCAATAATACCACTGG 10 TATGCCACTCATATGGTAAAGGAAACTCTAATTCCTCC 11 TATGCCACTCATATGATGGCCGCCTTCTCC 12 TATGCCACTCATATGACTCTCACGTTTTCCTC 13 ACAGCAGCATGCCTAAGCGTAATCTGGAACAT 14 CGTATCTGGATGGTGTTTACAATC 15 GGAATTCTTAATACCTCTCTCCAAATCGGAAGAAC 16 ATGGCCAATAATACCACTGG 17 TCCAAAATGCGACCATAAC 18 ATGGCTGAGGCTGATGATATTCAACCAATC 19 AGGAGCAATACGAAGCTCATTGTAGAAAGT Open in new tab Complementation of the dgkA − Mutant of E. coli Bacterial strain FB21625 (Jerga et al., 2007) was T7 lysogenized using a λDE3 lysogenization kit (Merck,). Freshly lysogenized cells were transformed with pDEST24 constructs, including unrelated Plastid DiVision1 (PDV1)-pDEST24 as a vector control. After overnight selection on ampicillin and kanamycin, single colonies were grown in culture overnight, normalized for optical density, and then spotted onto ampicillin kanamycin media or media supplemented with 90 mm hydroquinone β-d-glucopyranoside (arbutin, Sigma) and 0.1 m isopropyl β-d-1-thiogalactopyranoside (Roche) and allowed to grow for 24 h at 37°C. Chloroplast Protein Import Control plasmids were as previously described for SSU (Olsen and Keegstra, 1992) and ARC6 (Vitha et al., 2003). TOC34 cDNA was obtained from Arabidopsis Biological Resource Center (Yamada et al., 2003), inserted into pENTR-SD-DTOPO, and then subsequently transferred into pDEST14 (Invitrogen) according to the manufacturer’s protocol. Intact chloroplasts were isolated from 8- to 12-d-old pea (Pisum sativum) seedlings and purified as previously described (Bruce et al., 1994). Intact pea chloroplasts were reisolated and resuspended in import buffer (330 mm sorbitol and 50 mm HEPES-KOH, pH 8.0) at a concentration of 1 mg chlorophyll mL–1. Import assays were performed essentially as described (Tranel et al., 1995). Briefly, large-scale import assays were performed in import buffer at 600 μL final volume containing 200 μL chloroplasts, 4 mm Mg-ATP, and either [3H]Leu or [35S]Met-labeled precursor proteins. After incubation for 40 min at room temperature, assays were divided into two 300-μL aliquots. One aliquot was mock treated with import buffer, while the second fraction was incubated with Thermolysin or trypsin as previously described (Jackson et al., 1998) for an additional 30 min on ice. After quenching, intact chloroplasts were recovered by centrifugation through a 40% (v/v) Percoll cushion, lysed, and then fractionated into total soluble and membrane fractions. All fractions were subsequently analyzed by SDS-PAGE. After electrophoresis, the gel was subjected to fluorography and exposed to x-ray film (Eastman Kodak). Plant Material Constructs in pMDC32 were transformed into Agrobacterium tumefaciens strain GV3101 and then introduced into the Arabidopsis wild type (ecotype Columbia-2) by floral dip (Clough and Bent, 1998). T1 seeds were screened for resistance to hygromycin (20 μg mL–1) on Murashige and Skoog medium (Murashige and Skoog, 1962) solidified with 0.6% (w/v) agar gel (Sigma), transferred to soil, and confirmed by RT-PCR (see below). The T3 generation of each transgenic line was habitually screened by growth on hygromycin for 3 to 4 weeks, except slow-growing tpTOC75, which was screened for 4 to 5 weeks, before transfer to soil for phenotypic and lipid analyses. Plants tested are therefore expected to be homo- or heterozygous with respect to the DAGK transgene. Growth on soil was in a controlled chamber with 16-h/8-h photoperiod at 22°C/18°C (day/night) with a photosynthetic photon flux density of 70 to 80 μmol m–2 s–1 during the day, and plants were supplemented with one-half-strength Hoagland’s solution (Hoagland and Arnon, 1950). RT-PCR Total RNA was isolated from T1 DAGK lines or wild-type Arabidopsis with the RNeasy plant mini kit. The cDNA was produced from 1 μg of RNA using SuperScript III. Expression of TOC75-III (At3g46740) was tested with primers 14 and 15. Expression of DAGK was tested with primers 16 and 17. Expression of ACTin2 (ACT2; At3g18780) was tested using primers 18 and 19. Initial tests confirmed that the reactions were not saturated at the cycle number used. DAGK Assay Measurement of DAGK activity was based on methods described previously (Loomis et al., 1985; Fritz et al., 2007). Briefly, 100 μg of chlorophyll-equivalent chloroplasts isolated as described (Bruce et al., 1994) were resuspended on ice with 50 μL of 50 mm HEPES, pH 6.8, 50 mm NaCl, 12.5 mm MgCl2, 2 mm dithiothreitol, 1% (w/v) decylmaltoside (Affymetrix), and 0.2 mg mL–1 1,2-dioleoyl-sn-glycerol (Avanti Polar Lipids). In specified experiments, commercial E. coli DAGK (Enzo Life Sciences) was serially diluted, and then a consistent volume (1 μL) of DAGK was added to chloroplasts before resuspension in the above buffer. Measurement of the DAGK reaction was started by addition of 1 μL of ATP and tetra (triethylammonium) salt [γ-32P] (5 mCi mL–1, American Radiolabeled Chemicals) and then incubated at room temperature for 20 min or as specified. Reactions were stopped by addition of 100 μL of methanol:chloroform (2:1, v/v) and 50 μL of 0.2 m H3PO4 and 1 m KCl. After centrifugation, the chloroform phase was loaded onto JT-Baker PA-Si250 plates, pretreated and baked as described (Wang and Benning, 2011), and then resolved with chloroform:methanol:7 m ammonium hydroxide (65:25:5, v/v). Radioactive PtdOH and l-PtdOH reaction products were visualized by autoradiography and quantified by scintillation counting of bands scraped from the thin layer chromatography (TLC) plate. For identification, lipids including a lipid standard generated by partial Rhizopus arrhizus lipase digestion (Li et al., 2012) were reversibly stained by exposure to iodine vapor. Lipid Analysis Lipids were extracted from leaves of 6- to 7-week-old soil-grown plants and used for TLC analysis as described (Wang and Benning, 2011). Resolving solvents used were as follows: DAG and TAG, petroleum ether:diethyl ether:glacial acetic acid (80:20:1, v/v); polar lipids excepting PtdOH, acetone:toluene:water (91:30:7.5, v/v). PtdOH was resolved using a two-dimensional TLC system, with the first dimension solvent chloroform:methanol:7 m ammonium hydroxide (130:80:8, v/v) and the second dimension chloroform:methanol:glacial acetic acid:water (170:25:25:6, v/v), similar to that described (Xu et al., 2005). Polar lipids were resolved on JT-Baker PA-Si250 plates and pretreated and baked as described (Wang and Benning, 2011). TAG, DAG, and PtdOH were resolved on TLC plates baked for 1 h at 120°C from EMD Millipore. Each resolved lipid band analyzed was compared to total lipid of a known fraction spotted onto the TLC plate outside the resolving area. These were scraped from the TLC plates, converted into fatty acid methyl esters, normalized to a 15:0 standard, and quantified by gas chromatographyflame ionization detection as described (Wang and Benning, 2011). Data Processing Using the unresolved total lipid fraction, gas chromatography data were converted to mole percentage of total fatty acids to allow comparison across experiments. When more than one form existed, various forms of 16:1 or 18:1 fatty acids were combined. To ensure that small changes were not misrepresented, when data sets were combined for graphical display, each data set was normalized to its wild-type samples run in the same experiment and displayed at the average wild-type value across all experiments. Statistical analyses used a two-tailed, paired, or unpaired Student’s t test as appropriate and only compared data sets to wild-type samples run in the same experiment. Antisera Generation of antipeptide antiserum recognizing E. coli DAGK was performed as described (Ramer and Bell, 1990), using a similar peptide ANNTTGFTRIIKAC, with the services of LifeTein. Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers E. coli DAGK: AAA24394; DGD1: NM_111999; TOC75: NM_114541; ATS1: NM_102953; PDV1: NM_124707; SSU: NM_105379; ARC6: NM_123613; TOC34: NM_120582; ACT2: NM_112764. Supplemental Data The following materials are available in the online version of this article. Supplemental Figure S1. Levels and composition of MGDG, DGDG, SQDG, PtdIns, and phosphatidylethanolamine in DAGK-producing plants. ACKNOWLEDGMENTS We thank Sanjaya (Michigan State University) for measurement of chlorophyll, Sanjaya and Bensheng Liu (Michigan State University) for investigative mass spectrometry-based lipidomics, Charles O. Rock (St. Jude Children’s Research Hospital) for the gift of bacterial strain FB21625, and Anna Hurlock (Michigan State University) for critical reading of the manuscript. Glossary DAG diacylglycerol TAG triacylglycerol ER endoplasmic reticulum l-PtdOH lyso-phosphatidic acid PtdOH phosphatidic acid PtdGro phosphatidylglycerol MGDG monogalactosyldiacylglycerol SQDG sulfoquinovosyldiacylglycerol PtdCho phosphatidylcholine DGDG digalactosyldiacylglycerol PtdIns phosphatidyl inositol RT reverse transcription TLC thin layer chromatography cDNA complementary DNA LITERATURE CITED Allan D Thomas P Michell RH ( 1978 ) Rapid transbilayer diffusion of 1,2-diacylglycerol and its relevance to control of membrane curvature . Nature 276 : 289 – 290 Google Scholar Crossref Search ADS PubMed WorldCat Andersson MX, Dörmann P (2009) Chloroplast membrane lipid biosynthesis and transport. In AS Sandelius, H Aronsson, eds, The Chloroplast: Interactions with the Environment. 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Plant Cell 17 : 3094 – 3110 Google Scholar Crossref Search ADS PubMed WorldCat Yamada K Lim J Dale JM Chen HM Shinn P Palm CJ Southwick AM Wu HC Kim C Nguyen M et al. ( 2003 ) Empirical analysis of transcriptional activity in the Arabidopsis genome . Science 302 : 842 – 846 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the U.S. Department of Energy (grant nos. DE–FG02–98ER20305 to C.B. and DE–FG02–91ER20021 to J.E.F.) and Michigan AgBioResearch (to C.B.). * Address correspondence to [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Christoph Benning ([email protected]). [C] Some figures in this article are displayed in color online but in black and white in the print edition. [W] The online version of this article contains Web-only data. [OPEN] Articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.113.222513 © 2013 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2013. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Ragel, Paula; Streb, Sebastian; Feil, Regina; Sahrawy, Mariam; Annunziata, Maria Grazia; Lunn, John E.; Zeeman, Samuel; Mérida, Ángel

doi: 10.1104/pp.113.223420pmid: 23872660

Abstract STARCH SYNTHASE4 (SS4) is required for proper starch granule initiation in Arabidopsis (Arabidopsis thaliana), although SS3 can partially replace its function. Unlike other starch-deficient mutants, ss4 and ss3/ss4 mutants grow poorly even under long-day conditions. They have less chlorophyll and carotenoids than the wild type and lower maximal rates of photosynthesis. There is evidence of photooxidative damage of the photosynthetic apparatus in the mutants from chlorophyll a fluorescence parameters and their high levels of malondialdehyde. Metabolite profiling revealed that ss3/ss4 accumulates over 170 times more ADP-glucose (Glc) than wild-type plants. Restricting ADP-Glc synthesis, by introducing mutations in the plastidial phosphoglucomutase (pgm1) or the small subunit of ADP-Glc pyrophosphorylase (aps1), largely restored photosynthetic capacity and growth in pgm1/ss3/ss4 and aps1/ss3/ss4 triple mutants. It is proposed that the accumulation of ADP-Glc in the ss3/ss4 mutant sequesters a large part of the plastidial pools of adenine nucleotides, which limits photophosphorylation, leading to photooxidative stress, causing the chlorotic and stunted growth phenotypes of the plants. The metabolism of starch plays an essential role in the physiology of plants. Starch breakdown provides the plant with carbon skeletons and energy when the photosynthetic machinery is inactive (transitory starch) or in the processes of germination and sprouting (storage starch). Deficiencies in the accumulation of transitory starch in Arabidopsis (Arabidopsis thaliana) have been described previously, specifically in mutants affected in the plastidial phosphoglucomutase (PGM1) or the small subunit (APS1) of the ADP-Glc pyrophosphorylase (AGPase). While they are described as “starchless,” they actually contain small amounts of starch (1%–2% of the wild-type levels; Streb et al., 2009) and share similar phenotypic alterations, such as growth retardation when cultivated under a short-day photoregime and increased levels of soluble sugars during the light phase and reduced levels during the night (Caspar et al., 1985; Lin et al., 1988b; Schulze et al., 1991). Carbon partitioning is altered in these plants. As photosynthate cannot be accumulated as starch, it is diverted via hexose phosphates in the cytosol to the synthesis of Suc, which accumulates together with the hexose sugars, Glc and Fru (Caspar et al., 1985). In Arabidopsis, there are five starch synthase isoforms: one granule-bound starch synthase and four soluble starch synthases: SS1, SS2, SS3, and SS4. We have described previously an Arabidopsis mutant plant lacking SS3 and SS4 that is also severely affected in the accumulation of starch (Szydlowski et al., 2009). SS4 is involved in the initiation of the starch granule and controls the number of granules per chloroplast (Roldán et al., 2007). The elimination of SS3 in an ss4 background leads to an absence of starch in most of the chloroplasts, despite the fact that SS1 and SS2 are still present and total starch synthase activity is only reduced by 35% (Szydlowski et al., 2009). However, a very small proportion of chloroplasts of this mutant plant contain a single huge starch granule, which is also a characteristic of chloroplasts in the ss4 single mutant (D’Hulst and Mérida, 2012). Thus, like aps1 and pgm1, ss3/ss4 plants contain only small amounts of starch. However, unlike aps1 or pgm1 plants, most of the cells of this mutant have empty chloroplasts, without starch (Szydlowski et al., 2009). In this work, we have analyzed the phenotypic effects of the impaired starch accumulation of ss3/ss4 plants. We show that this mutant displays phenotypic changes that are not found in other mutants with very low levels of starch, such as aps1 or pgm1 plants. We provide evidence that extremely high levels of ADP-Glc accumulate in the ss3/ss4 plants. Using reverse genetics to block the pathway of starch synthesis upstream of the starch synthases reduced the level of ADP-Glc in ss3/ss4 plants and reverted the other phenotypic traits. This suggests that ADP-Glc accumulation is the causal factor behind the chlorotic and stunted growth phenotypes of the ss3/ss4 mutant. RESULTS Phenotypic Characterization of an ss3/ss4 Double Mutant Arabidopsis plants mutated in both SS4 and SS3 genes accumulate very low levels of starch in leaves (around 12% of wild-type values; Fig. 1A) but have slightly higher levels of soluble sugars, particularly Suc (Fig. 1, B–D). However, this double mutant plant displays some phenotypic effects, such as poor growth rate under a long-day photoregime and pale green color (Fig. 2), that are not observed in other starch synthase-deficient mutants (Delvallé et al., 2005; Zhang et al., 2005) or in mutants that accumulate very low amounts of starch, such as those affected in APS1 (Lin et al., 1988a; Fig. 1A) or in the chloroplastic PGM1 (Caspar et al., 1985; Fig. 1A). We have carried out a comprehensive phenotypic analysis of the ss3/ss4 mutant in order to identify the metabolic cause of its distinctive growth and morphological traits. Figure 1. Open in new tabDownload slide Starch, sugar, and adenine nucleotide contents of wild-type and mutant Arabidopsis rosettes. Wild type (Col-0), ss3, ss4, ss3/ss4, aps1, pgm1, aps1/ss3/ss4, and pgm1/ss3/ss4 plants were grown under LD conditions (16 h of light, 20°C/8 h of dark, 18°C) and an irradiance of 150 µE m−2 s−1. Plants were harvested at midday at the nine-leaf stage and processed as described in “Materials and Methods.” Eight to 10 plants per sample were used for the ss3/ss4 mutant and four to five plants for the other genotypes. Values represent means ± sd of determinations on six independent samples (three independent samples for measurements of starch, sugars, and ADP-Glc in ss3s/ss4). Letters indicate samples that were not significantly different (P ≥ 0.05) according to one-way ANOVA (Holm-Sidak test). There were no significant differences in AMP content between the eight genotypes. ADPG, ADP-Glc; FW, fresh weight. Figure 1. Open in new tabDownload slide Starch, sugar, and adenine nucleotide contents of wild-type and mutant Arabidopsis rosettes. Wild type (Col-0), ss3, ss4, ss3/ss4, aps1, pgm1, aps1/ss3/ss4, and pgm1/ss3/ss4 plants were grown under LD conditions (16 h of light, 20°C/8 h of dark, 18°C) and an irradiance of 150 µE m−2 s−1. Plants were harvested at midday at the nine-leaf stage and processed as described in “Materials and Methods.” Eight to 10 plants per sample were used for the ss3/ss4 mutant and four to five plants for the other genotypes. Values represent means ± sd of determinations on six independent samples (three independent samples for measurements of starch, sugars, and ADP-Glc in ss3s/ss4). Letters indicate samples that were not significantly different (P ≥ 0.05) according to one-way ANOVA (Holm-Sidak test). There were no significant differences in AMP content between the eight genotypes. ADPG, ADP-Glc; FW, fresh weight. Figure 2. Open in new tabDownload slide Effects of photoperiod on growth phenotypes of starch-deficient mutants. Plants were grown in growth cabinets under different photoperiods: LD (16 h of light/8 h of dark), SD (8 h of light/16 h of dark), and LL. The growth of the different mutants was documented by photographs of representative 3-week-old plants. WT, Wild type. Figure 2. Open in new tabDownload slide Effects of photoperiod on growth phenotypes of starch-deficient mutants. Plants were grown in growth cabinets under different photoperiods: LD (16 h of light/8 h of dark), SD (8 h of light/16 h of dark), and LL. The growth of the different mutants was documented by photographs of representative 3-week-old plants. WT, Wild type. Growth of the ss3/ss4 Mutant Plants were cultivated in controlled conditions under different photoregimes: long day (LD; 16 h of light/8 h of dark), short day (SD; 8 h of light/16 h of dark), and continuous light (LL), and their growth was documented by photographs of 21-d-old plants (Fig. 2) and by a time course of the growth of plants cultivated under LD conditions (Fig. 3). The Arabidopsis aps1 mutant, which lacks the small subunit of AGPase and accumulates only very low amounts of starch, was used as a control, and its growth rate was compared with those of ss3, ss4, ss3/ss4, and wild-type plants. The pgm1 mutant also accumulated very low amounts of starch and showed essentially the same growth rate as aps1 (data not shown). Figure 2 shows that aps1, ss4, and ss3/ss4 plants all grew more slowly than wild-type plants in the SD condition, with growth being most severely restricted in the ss3/ss4 mutant. In contrast, the ss3 mutant grew slightly faster than wild-type plants. Increasing the length of the light period greatly increased the growth of the aps1 mutant, which was restored to nearly wild-type levels in LL conditions (Fig. 2). However, the growth of ss3/ss4 in LD and LL conditions remained severely limited, while the phenotype of ss4 was intermediate between that of ss3/ss4 and wild-type plants (Figs. 2 and 3). These results indicate that the poor growth of ss3/ss4, and to a lesser extent of ss4, is not caused solely by the impairment in starch accumulation and that some other metabolic perturbation, which is specific to these mutants, restricts their growth. Figure 3. Open in new tabDownload slide Growth rates of wild-type (WT) Col-0, ss3, ss4, and ss3/ss4 plants. Plants were grown in growth cabinets under a LD photoperiod (16 h of light/8 h of dark). Aerial parts of the plants (five plants per sample) were weighed on the days indicated. Each point represents the mean ± sd of three independent experiments. Figure 3. Open in new tabDownload slide Growth rates of wild-type (WT) Col-0, ss3, ss4, and ss3/ss4 plants. Plants were grown in growth cabinets under a LD photoperiod (16 h of light/8 h of dark). Aerial parts of the plants (five plants per sample) were weighed on the days indicated. Each point represents the mean ± sd of three independent experiments. Photosynthetic Activity The low growth rate observed in ss4 and ss3/ss4 plants suggests that their photosynthetic activity is negatively affected by the defects in starch synthesis. To investigate which aspects of photosynthesis are affected, CO2 assimilation rate versus photosynthetically active radiation were determined on attached leaves of plants grown under a LD photoregime at 130 µmol m−2 s−1 and ambient CO2 using an open gas-exchange system. The light-response curves are shown in Figure 4. The light-saturated photosynthesis rates of ss4 (6.87 ± 0.39 µmol CO2 m−2 s−1) and ss3/ss4 (2.51 ± 0.52 µmol CO2 m−2 s−1) in these conditions were substantially lower (65% and 24% of the control rate, respectively) than the maximum value obtained for wild-type plants (10.57 ± 0.64 µmol CO2 m−2 s−1). Values obtained for the ss3 mutant plants (10.63 ± 0.32 µmol CO2 m−2 s−1) were not significantly different from wild-type plants (Fig. 4). Measurements of modulated chlorophyll a fluorescence (see “Materials and Methods”) indicated that the maximum quantum efficiency of PSII photochemistry (F v/F m) was drastically reduced in the ss3/ss4 mutant (0.63 ± 0.040 and 0.82 ± 0.002 for ss3/ss4 and wild-type plants, respectively; Table I). A reduction was also observed in the PSII operating efficiency (ΦPSII), indicating a decrease in the relative quantum yield of linear electron transfer through the photosystems (Baker, 2008). The values of the parameters obtained for ss4 mutant plants were also significantly different from those obtained for wild-type plants (Table I), being intermediate between wild-type and ss3/ss4 double mutant plants. This suggests that the single ss4 mutation leads to negative effects on both PSII efficiency and the CO2 assimilation rate of the plant, which are exacerbated by loss of the SS3 gene function. Figure 4. Open in new tabDownload slide Photosynthetic assimilation rates. Light-response curves of CO2 assimilation rates (A) of wild-type (WT), ss3, ss4, and ss3/ss4 plants were measured in a leaf chamber. Three-week-old plants grown in growth cabinets under a LD photoperiod were used to determine the CO2 assimilation rates. Each point represents the mean ± sd of five independent determinations. Figure 4. Open in new tabDownload slide Photosynthetic assimilation rates. Light-response curves of CO2 assimilation rates (A) of wild-type (WT), ss3, ss4, and ss3/ss4 plants were measured in a leaf chamber. Three-week-old plants grown in growth cabinets under a LD photoperiod were used to determine the CO2 assimilation rates. Each point represents the mean ± sd of five independent determinations. Chlorophyll a fluorescence parameters of wild-type, ss3, ss4, ss3/ss4, aps1, and aps1/ss3/ss4 plants Table I. Chlorophyll a fluorescence parameters of wild-type, ss3, ss4, ss3/ss4, aps1, and aps1/ss3/ss4 plants F v/F m and ΦPSII at 130 µE m−2 s−1 were determined in dark-adapted plants. Values are means ± se of four independent determinations. Significance is indicated as follows: a, not significantly different from the wild type according to Student’s t test (P ≥ 0.01); b, significantly different from the wild type according to Student’s t test (P < 0.01). Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.816 ± 0.001 0.513 ± 0.000 ss3 0.801 ± 0.001 a 0.483 ± 0.008 a ss4 0.754 ± 0.008 b 0.350 ± 0.006 b ss3/ss4 0.631 ± 0.040 b 0.220 ± 0.044 b aps1 0.803 ± 0.003 a 0.480 ± 0.010 a aps1/ss3/ss4 0.802 ± 0.003 a 0.478 ± 0.027 a Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.816 ± 0.001 0.513 ± 0.000 ss3 0.801 ± 0.001 a 0.483 ± 0.008 a ss4 0.754 ± 0.008 b 0.350 ± 0.006 b ss3/ss4 0.631 ± 0.040 b 0.220 ± 0.044 b aps1 0.803 ± 0.003 a 0.480 ± 0.010 a aps1/ss3/ss4 0.802 ± 0.003 a 0.478 ± 0.027 a Open in new tab Table I. Chlorophyll a fluorescence parameters of wild-type, ss3, ss4, ss3/ss4, aps1, and aps1/ss3/ss4 plants F v/F m and ΦPSII at 130 µE m−2 s−1 were determined in dark-adapted plants. Values are means ± se of four independent determinations. Significance is indicated as follows: a, not significantly different from the wild type according to Student’s t test (P ≥ 0.01); b, significantly different from the wild type according to Student’s t test (P < 0.01). Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.816 ± 0.001 0.513 ± 0.000 ss3 0.801 ± 0.001 a 0.483 ± 0.008 a ss4 0.754 ± 0.008 b 0.350 ± 0.006 b ss3/ss4 0.631 ± 0.040 b 0.220 ± 0.044 b aps1 0.803 ± 0.003 a 0.480 ± 0.010 a aps1/ss3/ss4 0.802 ± 0.003 a 0.478 ± 0.027 a Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.816 ± 0.001 0.513 ± 0.000 ss3 0.801 ± 0.001 a 0.483 ± 0.008 a ss4 0.754 ± 0.008 b 0.350 ± 0.006 b ss3/ss4 0.631 ± 0.040 b 0.220 ± 0.044 b aps1 0.803 ± 0.003 a 0.480 ± 0.010 a aps1/ss3/ss4 0.802 ± 0.003 a 0.478 ± 0.027 a Open in new tab Pigment Content The lower yield of linear electron flux through the photosystems observed in ss3/ss4 and ss4 mutant plants could trigger some adaptive responses, such as a decrease in the light-harvesting antenna size, in order to prevent possible photooxidative damage (Niyogi, 1999). Such a response would account for the pale green color that is characteristic of ss4 and ss3/ss4 mutants compared with the other starch-deficient mutants. To test this hypothesis, we measured the levels of different pigments in ss3/ss4 plants and compared them with the pigment contents of both the single parental mutants and wild-type plants. Figure 5 shows that the levels of antenna and accessory pigments (chlorophylls a and b and carotenoids) were about 45% lower in the double mutant than in wild-type plants. There was a smaller decrease, to around 63% of the wild type, in the ss4 single mutant. While chlorophyll and carotenoid levels were decreased, the ss4 and ss3/ss4 mutants accumulated more anthocyanins than the controls, with 1.4- and 5.8-fold higher levels than wild-type plants, respectively (Fig. 5). The pigment content of the ss3 mutant was not significantly different from the wild type. Figure 5. Open in new tabDownload slide Photosynthetic pigments and anthocyanin contents. Plants were grown in growth cabinets under a LD photoperiod, and discs of rosette leaves from four different plants per sample were used to determine the chlorophyll a and b, carotenoid, and anthocyanin contents of wild-type (WT), ss3, ss4, and ss3/ss4 plants (A) and wild-type, aps1, pgm1, ss3/ss4, aps1/ss3/ss4, and pgm1/ss3/ss4 plants (B). Values are means ± sd of three independent experiments. Asterisks indicate values that are significantly different from the wild type according to Student’s t test (P < 0.05). fw, Fresh weight. Figure 5. Open in new tabDownload slide Photosynthetic pigments and anthocyanin contents. Plants were grown in growth cabinets under a LD photoperiod, and discs of rosette leaves from four different plants per sample were used to determine the chlorophyll a and b, carotenoid, and anthocyanin contents of wild-type (WT), ss3, ss4, and ss3/ss4 plants (A) and wild-type, aps1, pgm1, ss3/ss4, aps1/ss3/ss4, and pgm1/ss3/ss4 plants (B). Values are means ± sd of three independent experiments. Asterisks indicate values that are significantly different from the wild type according to Student’s t test (P < 0.05). fw, Fresh weight. Photooxidative Stress Reducing the levels of antenna pigments would be expected to help balance the input and utilization of light energy in the ss4 and ss3/ss4 mutants. Nevertheless, despite this adaptive response, blockage of the electron flux between photosystems could result in photooxidative damage in these plants. Photooxidative damage caused by reactive oxygen species (ROS) in ss4 and ss3/ss4 was confirmed by their higher content of malondialdehyde (MAD; Fig. 6), which is a marker of membrane lipid peroxidation caused by elevated levels of ROS (Janero, 1990). The MAD content was 2.5 times higher in ss3/ss4 than in wild-type plants, while ss4 plants had levels intermediate between ss3/ss4 and the wild type, being 1.5 times higher than wild-type plants (Fig. 6). Figure 6. Open in new tabDownload slide MAD accumulation. MAD content in leaves of wild-type (WT), ss3, ss4, and ss3/ss4 plants grown under a LD photoperiod was determined using the thiobarbituric acid method. Values are means ± sd of four independent experiments. Asterisks indicate values that are significantly different from the wild type according to Student’s t test (P < 0.05). Figure 6. Open in new tabDownload slide MAD accumulation. MAD content in leaves of wild-type (WT), ss3, ss4, and ss3/ss4 plants grown under a LD photoperiod was determined using the thiobarbituric acid method. Values are means ± sd of four independent experiments. Asterisks indicate values that are significantly different from the wild type according to Student’s t test (P < 0.05). The ss3/ss4 and ss4 Mutants Accumulate ADP-Glc ADP-Glc is the substrate for starch synthesis. Therefore, impairment of starch synthesis as found in ss4 and ss3/ss4 mutants (Roldán et al., 2007; Szydlowski et al., 2009) might be expected to affect the accumulation of ADP-Glc, especially in the ss3/ss4 mutant, which has very low levels of starch. ADP-Glc was measured in wild-type and mutant plants by high-performance anion-exchange chromatography coupled to tandem mass spectrometry (MS/MS; Lunn et al., 2006). This method provides greater specificity and sensitivity than the HPLC-based methods with UV detection that have generally been used to measure ADP-Glc in plants. The ss3/ss4 plants were found to have extremely high levels of ADP-Glc (341 nmol g−1 fresh weight), which was around 170-fold higher than in wild-type plants (Fig. 1E). The content of this compound in other mutants that accumulate only trace amounts of starch, such as aps1 and pgm1, was lower than in wild-type plants (Fig. 1E) as a consequence of the mutations in the plastidial enzymes that synthesize ADP-Glc and Glc-1-P, respectively. The levels of Suc, Fru, and Glc in ss3/ss4 plants were lower than in aps1 or pgm1, which contained 2- to 3-fold higher soluble sugar levels than wild-type plants (Fig. 1, B–D). Measurements of other metabolites by high-performance anion-exchange chromatography coupled to MS/MS showed a 2-fold higher level of trehalose 6-phosphate in ss3/ss4 compared with the wild type (Supplemental Fig. S1), presumably reflecting the 2-fold higher Suc content of the mutant (Fig. 1D). There were no striking differences between ss3/ss4 and the wild type in the levels of other sugar phosphates or glycolytic intermediates, apart from a decrease in Fru-1,6-bisP in the mutant (Supplemental Fig. S1). Citrate, aconitate, and isocitrate were slightly elevated in the mutant, while the other tricarboxylic acid cycle intermediates were either unchanged (2-oxoglutarate and succinate) or slightly lower (malate and fumarate) in the ss3/ss4 mutant (Supplemental Fig. S1). Restricting Plastidial ADP-Glc Synthesis Substantially Restores Growth in the ss3/ss4 Plants The high levels of ADP-Glc found in ss3/ss4 plants are presumably caused by an imbalance between the rates of ADP-Glc synthesis by AGPase and the consumption for starch synthesis, which is essentially blocked in this mutant. As a result, it seems likely that much of the adenine nucleotides in the chloroplasts will be effectively locked away in this large pool of ADP-Glc, restricting photophosphorylation, CO2 fixation, and other process that are dependent on ATP. To test this hypothesis, we crossed ss3/ss4 plants with mutant plants lacking the small subunit of AGPase (aps1) or the plastidial isoform of PGM (pgm1). Both mutants are greatly impaired in the synthesis of plastidial ADP-Glc and accumulate only very small amounts of starch (Stitt and Zeeman, 2012; Fig. 1A). Progeny of the respective crosses were analyzed to identify aps1/ss3/ss4 and pgm1/ss3/ss4 triple mutants (see “Materials and Methods”), and these plants were selected for subsequent analyses. Mutant plants cultured under a 16-h-light/8-h-dark (aps1/ss3/ss4) or a 12-h-light/12-h-dark (pgm1/ss3/ss4) photoregime showed the same growth rate as the aps1 or pgm1 parental plants, respectively, which were higher than the growth rate of ss3/ss4 parental plants (Fig. 7; Supplemental Fig. S2). Attached leaves of aps1/ss3/ss4 plants or whole pgm1/ss3/ss4 plants grown under the same conditions detailed above were used to determine their CO2 assimilation rates. Under saturating light, the CO2 assimilation rates of the ss3/ss4 (2.51 ± 0.52 µmol CO2 m−2 s−1) and aps1 (6.08 ± 0.23 µmol CO2 m−2 s−1) mutants were 61% and 36% lower, respectively, than in wild-type plants (9.29 ± 1.01 µmol CO2 m−2 s−1; Fig. 8). The CO2 assimilation rate of aps1/ss3/ss4 plants (6.69 ± 0.77 µmol CO2 m−2 s−1) was similar to that found for aps1 plants, indicating that the nearly complete loss of AGPase activity overrides the severe inhibition of photosynthesis in the ss3/ss4 mutant background (Fig. 8). Similarly, restricting ADP-Glc synthesis via the loss of plastidial PGM activity restored photosynthetic rates in the pgm1/ss3/ss4 mutant to wild-type levels (Fig. 9). Figure 7. Open in new tabDownload slide Growth phenotypes of wild-type (WT), aps1, pgm1, ss3/ss4, aps1/ss3/ss4, and pgm1/ss3/ss4 plants. The photographs show 3-week-old plants cultivated in growth cabinets under a 16-h-light/8-h-dark (aps1, ss3/ss4, and aps1/ss3/ss4) or a 12-h-light/12-h-dark (pgm1, ss3/ss4, and pgm1/ss3/ss4) photoperiod. Figure 7. Open in new tabDownload slide Growth phenotypes of wild-type (WT), aps1, pgm1, ss3/ss4, aps1/ss3/ss4, and pgm1/ss3/ss4 plants. The photographs show 3-week-old plants cultivated in growth cabinets under a 16-h-light/8-h-dark (aps1, ss3/ss4, and aps1/ss3/ss4) or a 12-h-light/12-h-dark (pgm1, ss3/ss4, and pgm1/ss3/ss4) photoperiod. Figure 8. Open in new tabDownload slide Photosynthetic assimilation rates. Light-response curves of CO2 assimilation rates (A) of wild-type (WT), aps1, ss3/ss4, and aps1/ss3/ss4 plants were measured in a leaf chamber. Three-week-old plants grown in growth cabinets under a LD photoperiod were used to determine the CO2 assimilation rates. Each point represents the mean ± sd of five independent determinations. Figure 8. Open in new tabDownload slide Photosynthetic assimilation rates. Light-response curves of CO2 assimilation rates (A) of wild-type (WT), aps1, ss3/ss4, and aps1/ss3/ss4 plants were measured in a leaf chamber. Three-week-old plants grown in growth cabinets under a LD photoperiod were used to determine the CO2 assimilation rates. Each point represents the mean ± sd of five independent determinations. Figure 9. Open in new tabDownload slide Photosynthetic assimilation rates. Whole, 3-week-old plants were used to determine the photosynthetic CO2 assimilation rates (A) in a multichamber system at an irradiance of 150 µE m−2 s−1. Values are means ± sd of four biological replicates. WT, Wild type. Figure 9. Open in new tabDownload slide Photosynthetic assimilation rates. Whole, 3-week-old plants were used to determine the photosynthetic CO2 assimilation rates (A) in a multichamber system at an irradiance of 150 µE m−2 s−1. Values are means ± sd of four biological replicates. WT, Wild type. Measurements of F v/F m and ΦPSII showed that both of these photosynthetic parameters in the aps1/ss3/ss4 and pgm1/ss3/ss4 triple mutants had reverted to wild-type values (Tables I and II). The levels of photosynthetic pigments (chlorophylls a and b and carotenoids) in aps1/ss3/ss4 were similar to those of wild-type plants (Fig. 5), providing further evidence that restricting ADP-Glc counteracted the negative effects of the loss of SS3 and SS4 activity on photosynthesis. Levels of anthocyanins in the aps1/ss3/ss4 mutant plants were much lower than in the parental ss3/ss4 mutant, being slightly higher than in wild-type plants but similar to the parental aps1 mutant (Fig. 5). Metabolite profiling of the triple mutants revealed that they accumulated 87% to 97% less ADP-Glc than the ss3/ss4 mutant (Fig. 1E). However, the levels of ADP-Glc in aps1/ss3/ss4 plants were still considerably higher (around 23-fold) than in wild-type plants but only about 5-fold higher than the wild type in the pgm1/ss3/ss4 mutant. Chlorophyll a fluorescence parameters of wild-type, ss3, ss4, ss3/ss4, pgm1, and pgm1/ss3/ss4 plants Table II. Chlorophyll a fluorescence parameters of wild-type, ss3, ss4, ss3/ss4, pgm1, and pgm1/ss3/ss4 plants F v/F m and ΦPSII at 130 µE m−2 s−1 were determined in dark-adapted plants. Values are means ± se of four independent determinations. Significance is indicated as follows: a, not significantly different from the wild type according to Student’s t test (P ≥ 0.01); b, significantly different from the wild type according to Student’s t test (P < 0.01). Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.808 ± 0.0005 0.394 ± 0.009 ss3 0.803 ± 0.0015 a 0.414 ± 0.003 a ss4 0.771 ± 0.0015 b 0.315 ± 0.002 b ss3/ss4 0.726 ± 0.005 b 0.204 ± 0.005 b pgm1 0.790 ± 0.000 a 0.381 ± 0.003 a pgm1/ss3/ss4 0.779 ± 0.0005 a 0.391 ± 0.006 a Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.808 ± 0.0005 0.394 ± 0.009 ss3 0.803 ± 0.0015 a 0.414 ± 0.003 a ss4 0.771 ± 0.0015 b 0.315 ± 0.002 b ss3/ss4 0.726 ± 0.005 b 0.204 ± 0.005 b pgm1 0.790 ± 0.000 a 0.381 ± 0.003 a pgm1/ss3/ss4 0.779 ± 0.0005 a 0.391 ± 0.006 a Open in new tab Table II. Chlorophyll a fluorescence parameters of wild-type, ss3, ss4, ss3/ss4, pgm1, and pgm1/ss3/ss4 plants F v/F m and ΦPSII at 130 µE m−2 s−1 were determined in dark-adapted plants. Values are means ± se of four independent determinations. Significance is indicated as follows: a, not significantly different from the wild type according to Student’s t test (P ≥ 0.01); b, significantly different from the wild type according to Student’s t test (P < 0.01). Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.808 ± 0.0005 0.394 ± 0.009 ss3 0.803 ± 0.0015 a 0.414 ± 0.003 a ss4 0.771 ± 0.0015 b 0.315 ± 0.002 b ss3/ss4 0.726 ± 0.005 b 0.204 ± 0.005 b pgm1 0.790 ± 0.000 a 0.381 ± 0.003 a pgm1/ss3/ss4 0.779 ± 0.0005 a 0.391 ± 0.006 a Genotype . F v/F m . ΦPSII . Wild type (Col-0) 0.808 ± 0.0005 0.394 ± 0.009 ss3 0.803 ± 0.0015 a 0.414 ± 0.003 a ss4 0.771 ± 0.0015 b 0.315 ± 0.002 b ss3/ss4 0.726 ± 0.005 b 0.204 ± 0.005 b pgm1 0.790 ± 0.000 a 0.381 ± 0.003 a pgm1/ss3/ss4 0.779 ± 0.0005 a 0.391 ± 0.006 a Open in new tab To assess the impact of the massive accumulation of ADP-Glc on other adenine nucleotides, we measured ATP, ADP, and AMP in mutant and wild-type plants. ATP was about 27% lower in the ss3/ss4 mutant than in wild-type plants (Fig. 1F). The difference was statistically significant (P = 0.047) according to Student’s t test but not significant according to one-way ANOVA (Holm-Sidak test; Fig. 1F). ADP also showed a tendency to be decreased in the ss3/ss4 mutant (Fig. 1G). Both ATP and ADP were significantly lower in the pgm mutant than in the wild type (Fig. 1F). The total adenine nucleotide pools (ATP + ADP + AMP + ADP-Glc) in the ss3/ss4 and ss4 mutants were about four times higher than in wild-type plants, with ADP-Glc constituting 81%, 77%, and 2% of the total adenylate pool, respectively (Fig. 1; Supplemental Fig. S3). The ATP, ADP, and total adenylate pool sizes in the aps1/ss3/ss4 and pgm/ss3/ss4 triple mutants were indistinguishable from the wild-type levels (Fig. 1, F and G; Supplemental Fig. S3). DISCUSSION We have previously reported that Arabidopsis requires SS4 or SS3 to synthesize starch (Szydlowski et al., 2009), strongly implicating SS4 and SS3 in the process of granule initiation. In subsequent analyses of the ss3/ss4 mutant, we have observed that a small number of chloroplasts do contain single starch granules with a distinctive shape, being more rounded than the discoid granules usually found in Arabidopsis leaves (D’Hulst and Mérida, 2012). This suggests that in the absence of SS4, stochastic initiation of starch granules can still occur, albeit very infrequently, even when SS3 is also missing (D’Hulst and Mérida, 2012). This would explain the trace amounts of starch in the ss3/ss4 plants (Fig. 1A). The most obvious phenotypic traits of the ss3/ss4 plants are their severely restricted growth and chlorotic leaves (Figs. 2 and 3). Transitory starch reserves provide the plant with a buffer itself against diurnal variation in carbon and energy supplies (Sun et al., 1999; Smith and Stitt, 2007). Mutants impaired in starch synthesis, such as aps1 and pgm1, accumulate high levels of Suc and hexoses (Caspar et al., 1985) as a consequence of their inability to store net photosynthate in starch. However, the total amount of carbohydrate accumulated as soluble sugars in starch-deficient mutants during the day is much less than the amount of starch accumulated by wild-type plants (Fig. 1, A–D). Furthermore, soluble sugars are directly available for export and respiration, so these reserves, which are already smaller than those in wild-type plants, are rapidly depleted during the first hours of the night. This leads to carbon starvation by the end of the night, which suppresses growth by mechanisms that are not yet fully understood but probably involve the inhibition of translation and other biosynthetic processes and wasteful turnover of proteins (Gibon et al., 2004). The carbon starvation diminishes as the length of the night period is reduced; hence, the growth rates of the aps1 (Fig. 2) and pgm1 mutant plants in LD or LL conditions were practically the same as that observed in wild-type control plants (Lin et al., 1988a). The ss3/ss4 plants show a severe dwarf phenotype even in LD or LL photoregimes (Figs. 2 and 3), suggesting that the impairment in starch synthesis on its own is not the underlying cause of the poor growth of these plants and that other metabolic changes must be responsible for their stunted phenotype. In chloroplasts, the substrate for starch synthesis is derived from the Calvin-Benson cycle in the form of Fru-6-P. This is converted to Glc-6-P and then Glc-1-P in two reversible reactions catalyzed by phosphoglucose isomerase and phosphoglucomutase, respectively. The next step in the pathway, the synthesis of ADP-Glc by AGPase, is rendered irreversible by the hydrolysis of the coproduct, pyrophosphate, by inorganic pyrophosphatase (Ballicora et al., 2003). Thus, loss of plastidial phosphoglucose isomerase, PGM, or AGPase activity essentially blocks both ADP-Glc and starch synthesis, although a small amount of starch may be synthesized from imported hexose phosphate or ADP-Glc in young developing leaves, bypassing the metabolic lesions in these mutants (Streb et al., 2009; Bahaji et al., 2011). The situation is different in the ss3/ss4 mutant, where starch synthesis is limited not by the availability of ADP-Glc but by the almost complete failure of starch granule initiation and the consequent lack of primers for glucan synthesis. Given the pyrophosphatase-driven irreversibility of the AGPase reaction, there is no obvious reason why ADP-Glc should not continue to be synthesized. Starch synthesis is the main route for catabolizing ADP-Glc in the chloroplasts (Ball et al., 2011), but as this route is blocked in the ss3/ss4 mutant, there is likely to be an imbalance between ADP-Glc production and consumption, leading to its accumulation in the chloroplast stroma. Metabolite profiling confirmed that the ss3/ss4 mutant does indeed accumulate massive amounts of ADP-Glc (341 ± 16 nmol g−1 fresh weight), reaching levels that are over 170 times higher than in wild-type plants (Fig. 1E). In addition, levels of soluble sugars are only slightly increased in the ss3/ss4 mutant (Fig. 1, B–D), likely caused by the severe decrease of CO2 fixation in this mutant, which determines that the carbon diverted to soluble sugars, such as Suc, as consequence of the blockage of starch synthesis is lower than in aps1 or pgm1 mutants. The unprecedented accumulation of ADP-Glc in the ss3/ss4 plants will have important knock-on effects in the chloroplasts, as the adenosine and phosphate moieties within the ADP-Glc pool will in effect be locked away and unavailable for other metabolic processes. In wild-type plants, ATP and ADP are the dominant adenine nucleotides, with AMP and ADP-Glc forming only a small percentage (about 2% each) of the total adenine nucleotide pool (Fig. 1, E–H; Supplemental Fig. S3). Our measurements of ATP and ADP in wild-type plants indicate a total pool size of around 100 nmol g−1 fresh weight, close to the value of 120 nmol g−1 fresh weight reported by Strand et al. (2000). This is actually lower than the amount of ADP-Glc alone that we found in the ss3/ss4 and ss4 mutants, and the total adenylate pools in these two mutants are four times higher than in wild-type plants (Supplemental Fig. S3). This suggests that there has been a massive compensatory increase in the total adenine nucleotide pool size in the mutants, but despite this increase, the vast majority (77%–81%) of the enlarged pools are still sequestered in the form of ADP-Glc. As a consequence, there will be very little ADP available in the chloroplast stroma for photophosphorylation, which in turn will have severely negative effects on many metabolic and other processes in the chloroplasts. The most obvious effect of impaired ATP synthesis will be the limitation of photosynthetic CO2 assimilation, as observed in the ss3/ss4 mutant (Fig. 4). Within the Calvin-Benson cycle, ATP is required not only for the regeneration of ribulose-1,5-bisphosphate but also for the synthesis of triose phosphates from 3-phosphoglycerate (Heldt and Heldt, 2006; MacRae and Lunn, 2006). Limiting the synthesis of triose phosphates will, in turn, restrict their export to the cytosol for the synthesis and export of Suc to sink organs such as developing leaves, roots, flowers, and seeds. Low stromal ATP levels will also inhibit many of the biosynthetic pathways that occur in the chloroplasts (e.g. for the production of lipids, amino acids, nucleotides, isoprenoids, phenylpropanoids, and vitamins; Lunn, 2007). Presumably, at some point, ATP will become limiting for the further synthesis of ADP-Glc itself. However, the K m (ATP) of the Arabidopsis AGPase is around 70 µm (Crevillén et al., 2003), so it is capable of drawing down the stromal concentration of ATP to very low levels. ATP is also required for many housekeeping functions in the chloroplasts, including DNA, RNA, and protein synthesis, as well as the import, processing, and assembly of nucleus-encoded proteins. Altogether, the profound disturbance of chloroplast metabolism caused by low ATP availability is bound to have far-reaching effects beyond the chloroplasts, reducing the energy and carbon supplies available for cell maintenance and growth. Low ADP availability in the chloroplasts could also result in damage to existing chloroplast structures due to overenergization of the thylakoid membranes. The reduced rate of photophosphorylation and, therefore, dissipation of the proton-motive force across the thylakoid membranes will lead to hyperpolarization of the membranes, blocking further electron transport between the photosystems (Heldt and Heldt, 2006). This will lower the relative quantum yield of the linear electron transport, leading to a reduction in ΦPSII, as observed in the ss3/ss4 plants (Table I; Baker, 2008). This situation favors the formation of ROS that can cause oxidative damage to the photosynthetic apparatus (Niyogi, 1999). This is corroborated by the low value of F v/F m found in the ss3/ss4 plants, which is indicative of photoinhibition (Maxwell and Johnson, 2000). Further evidence of oxidative stress in the ss3/ss4 plants came from the finding of high levels of MAD in these plants, which is a characteristic product of membrane lipid peroxidation by ROS (Janero, 1990). One of the adaptive responses of plants to photoinhibitory damage is the adjustment of the light-harvesting antenna size in order to balance light absorption and utilization (Niyogi, 1999). This response explains the reduction in the amounts of chlorophylls a and b and carotenoids found in the ss3/ss4 plants (Fig. 2). Finally, the high levels of anthocyanins detected in the ss3/ss4 plants provide further support for the conclusion that the ss3/ss4 plants are subject to photoinhibition. Anthocyanin accumulation generally coincides with situations where there is an imbalance between light capture, CO2 assimilation, and carbohydrate utilization (Wand et al., 2002). All of the phenotypic traits described for the ss3/ss4 plants were also detected in the parental ss4 single mutant, such as pale color (Fig. 2; Roldán et al., 2007), lower growth and CO2 assimilation rates (Figs. 3 and 4), and low values of F v/F m and ΦPSII (Table I), although these effects were less severe than in the ss3/ss4 plants. Plants lacking SS4 accumulate around 60% to 80% of the starch found in wild-type plants (Roldán et al., 2007), suggesting that there is greater turnover of the ADP-Glc pool in ss4 than in the ss3/ss4 double mutant. Nevertheless, we observed a similarly large accumulation of ADP-Glc in both mutants, indicating that the limitation of starch synthesis in the single ss4 mutant is sufficient to have a major impact on ADP-Glc levels and to trigger a similar compensatory increase in the total adenylate pool to that seen in the ss3/ss4 mutant. We tested the hypothesis that the chlorotic and growth phenotypes of the ss3/ss4 mutant are attributable to the massive ADP-Glc accumulation by introducing further mutations (aps1 and pgm1) to limit ADP-Glc synthesis. The pgm1 mutation blocks the production of Glc-1-P in the chloroplasts, so there is no substrate available for ADP-Glc synthesis, except for a very small amount imported from the cytosol by a Glc-1-P transporter (Fettke et al., 2011). Introduction of the pgm1 mutation into the ss3/ss4 background lowered the amount of ADP-Glc accumulated in the leaves by 97% and restored the photosynthetic capacity and growth of the plants to nearly wild-type levels (Table II; Figs. 1E, 5, and 7–9). Knocking out the APS1 gene, encoding the small subunit of AGPase, led to an 87% decrease in the accumulation of ADP-Glc in the ss3/ss4 background but was less effective than the loss of PGM activity (Fig. 1E). Nevertheless, the aps1/ss3/ss4 triple mutant grew much better than the ss3/ss4 double mutant and was phenotypically similar to the aps1 parent (Table I; Figs. 5, 7, and 8). Loss of the APS1 subunit does not appear to abolish AGPase activity entirely, because the aps1 mutant was found to contain low but detectable traces of ADP-Glc and a small amount of starch (about 2% of the wild-type level; Fig. 1A). The residual activity can be ascribed to the remaining AGPase large subunits (APL1–APL4), several of which have been demonstrated to have a low level of catalytic activity (Ventriglia et al., 2008). Metabolites were measured in plants harvested after 8 h of illumination. If we assume that all of the ADP-Glc in the aps1/ss3/ss4 mutant (44 nmol g−1 fresh weight; Table I) had been synthesized during this period and that there was negligible consumption of ADP-Glc for starch synthesis, we can calculate that as little as 0.09 nmol min−1 g−1 fresh weight of AGPase activity would be sufficient to account for the observed accumulation of ADP-Glc in the triple mutant. This level of activity would represent less than 0.01% of the maximal catalytic activity of AGPase in wild-type Arabidopsis rosettes (1.0–1.5 µmol min−1 g−1 fresh weight; Hädrich et al., 2012). The amount of ADP-Glc in the aps1/ss3/ss4 mutant constitutes 34% of the total adenylate pool (Fig. 1E; Supplemental Fig. S3). Thus, by limiting the accumulation of ADP-Glc to a moderate level, a substantial pool of ATP and ADP should remain available for photophosphorylation and other biochemical reactions in the aps1/ss3/ss4 mutant. The photosynthetic capacities and growth rates of the aps1/ss3/ss4 and aps1 mutants were very similar, indicating that even such a partial rebalancing of the adenine nucleotide pool was sufficient to support higher rates of CO2 fixation and growth. Loss of plastidial PGM activity was even more effective at restoring the adenine nucleotide and ADP-Glc pools of the pgm/ss3/ss4 mutant to nearly wild-type levels (Fig. 1, E–H). In conclusion, the loss of SS3 and SS4 activity in Arabidopsis leaves almost totally abolishes starch synthesis due to the plant’s inability to initiate starch granules in most of the cells. The block in starch synthesis was found to result in an unprecedented accumulation of ADP-Glc, thereby sequestering much of the plastidial adenine nucleotide in a metabolically inaccessible form. This will severely limit the synthesis and availability of ATP for photosynthetic CO2 fixation and many other processes and will also trigger ROS production, leading to photooxidative damage of the photosynthetic apparatus within the chloroplasts. Restricting the synthesis of ADP-Glc in the chloroplasts by introducing the pgm1 or aps1 mutation into ss3/ss4 greatly reduced the accumulation of ADP-Glc and restored both the photosynthetic capacity and growth of the plants. This confirmed that the massive accumulation of ADP-Glc in the ss3/ss4 mutant underlies the chlorotic and stunted growth phenotype of this mutant and explains why it differs so markedly from other starch-deficient mutants. MATERIALS AND METHODS Plant Materials and Growth Conditions All Arabidopsis (Arabidopsis thaliana) mutants were in the Columbia-0 (Col-0) background. The ss3, ss4, and ss3/ss4 mutants were as described by Szydlowski et al. (2009), and the pgm1 mutant was from Caspar et al. (1985). The aps1 mutant (SALK_ 040155) contains a transfer DNA insertion in the APS1 gene (At5g48300; Alonso et al., 2003), and homozygous knockout plants were selected using PCR-based genotyping. Lines carrying triple mutations were obtained by crossing and selecting homozygous triple mutant plants from the segregating F2 populations using PCR-based genotyping. All primers used are described in Supplemental Table S1. Unless otherwise specified, plants were grown in growth cabinets at 23°C (day)/20°C (night), 70% humidity, and an irradiance of 120 µE m−2 s−1 supplied by white fluorescent lamps. Seeds were sown in soil and irrigated with 0.5× Murashige and Skoog medium (Murashige and Skoog, 1962). Photosynthesis Measurements Photosynthetic gas exchange was measured using a portable infrared gas analyzer (model LI-6400; LI-COR Biosciences), which allows environmental conditions inside a standard leaf clamp of the chamber to be precisely controlled. Air temperature in the chamber was set at 25°C, and the relative humidity was maintained at 50%. Light-response curves were determined on one of the upper leaves of the plants, progressively increasing the irradiance from 0 to 50 and then to 100, 250, 500, 750, 1,000, 1,500, and 2,000 μmol quanta m−2 s−1 in stepwise changes every 3 min. The CO2 assimilation rate was determined on 3-week-old plants grown in a controlled environment under LD conditions (16 h of light/8 h of dark) with an irradiance of 100 μmol quanta m−2 s−1 at a constant 22°C temperature. Photosyn Assistant software, developed by Dundee Scientific (R. Parsons and S.A. Ogston), has been used to determine parameters such as CO2 assimilation rates to help in the comparison between the mutants. Gas exchange of whole Arabidopsis rosettes was measured with a custom-built multichamber system connected in parallel to an infrared gas analyzer (Li-7000; LI-COR). Individual plants were introduced into each of eight chambers, and after an adaptation period of 2 d, gas exchange was measured for a complete 24-h light/dark period. During the measurement, air with 380 µL L−1 CO2 and a relative humidity of 60% was directed through the system. Each chamber was measured consecutively for 6 min, and the average value was taken for the light period. At the end of the experiment, a photograph was taken to calculate the projected leaf area for each plant. Based on the area, photosynthetic and respiration rates were calculated with the ƊCO2 and ƊH2O values gained from the gas-exchange system. Chlorophyll fluorescence emission parameters were measured at 22°C with a PAM 2000 fluorometer (Walz). The F v/F m was calculated from the measured parameters using the following equation: F v/F m = (F m − F o)/F m, where F o is the initial minimal fluorescence emitted from leaves dark adapted for 1 h and F m is the maximal fluorescence elicited by saturating actinic light. The ΦPSII was calculated from the measured parameters using the following equation: ΦPSII = F m′ − F′/F m′, where F m′ is the maximal fluorescence from light-adapted leaves and F′ is the fluorescence emission from light-adapted leaves. Plant Growth Measurement Plant growth was determined by weighing the aerial part of the plants. In the case of pgm1 and pgm1/ss3/ss4 mutant plants, the plant growth was measured based on whole rosette area. Digital images were taken every 2 or 3 d at the same time. The digital images were converted with Photoshop (Adobe Photoshop CS4 Extended; Adobe Systems) to black-and-white images, with the rosette being black and the background being white. ImageJ 1.42q (National Institutes of Health) was used to calculate the leaf area, based on the scale, from the black-and-white images. Analytical Methods Seven discs of rosette leaves (5–10 mg) from independent plants were used for the determination of chlorophylls a and b and carotenoids. Pigments were extracted with methanol and quantified according to the methods described by Porra et al. (1989) for chlorophylls and by Lichtenthaler and Buschmann (2001) for carotenoids. The extraction and quantitation of anthocyanins were performed as described (Rabino and Mancinelli, 1986) using seven rosette leaves from independent plants. The concentration of MAD in plants was determined using the thiobarbituric acid method described by Buege and Aust (1978). Soluble sugars and starch were measured enzymatically in ethanolic extracts and the ethanol-insoluble residue as described by Hendriks et al. (2003). ADP-Glc, phosphorylated intermediates, and organic acids were measured in chloroform-methanol extracts using high-pressure anion-exchange chromatography coupled to MS/MS as described (Lunn et al., 2006). ATP, ADP, and AMP were measured enzymatically in freshly prepared TCA extracts as described (Weiner et al., 1987; Trethewey et al., 1998). Supplemental Data The following materials are available in the online version of this article. Supplemental Figure S1 . Determination of levels of phosphorylated intermediates and organic acids of wild-type, ss3, ss4, ss3/ss4, aps1, pgm1, aps1/ss3/ss4, and pgm1/ss3/ss4 Arabidopsis plants grown in 16-h-light/8-h-dark conditions (20°C/18°C, irradiance of 150 µE m−2 s−1) and harvested at 8 h into the light period. Supplemental Figure S2. Growth rates of Col-0, aps1/ss3/ss4, pgm1/ss3/ss4, and their parental lines. Supplemental Figure S3. Total adenine nucleotide pool size (ATP + ADP + AMP + ADP-Glc) was measured in wild-type, ss3, ss4, ss3/ss4, aps1, aps1/ss3/ss4, pgm, and pgm/ss3/ss4 Arabidopsis plants grown in 16-h-light/8-h-dark conditions (20°C/18°C, irradiance at 150 µE m−2 s−1) and harvested at 8 h into the light period. Supplemental Table S1. Primers used for the identification of the triple mutant plants. Glossary LD long day SD short day LL continuous light F v/F m maximum quantum efficiency of PSII photochemistry ΦPSII PSII operating efficiency ROS reactive oxygen species MAD malondialdehyde MS/MS tandem mass spectrometry Col-0 Columbia-0 LITERATURE CITED Alonso JM Stepanova AN Leisse TJ Kim CJ Chen H Shinn P Stevenson DK Zimmerman J Barajas P Cheuk R et al. ( 2003 ) Genome-wide insertional mutagenesis of Arabidopsis thaliana . 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Plant Physiol 148 : 65 – 76 Google Scholar Crossref Search ADS PubMed WorldCat Wand S Holcroft D Jacobs G ( 2002 ) Anthocyanins in vegetative tissues: a proposed unified function in photoprotection . New Phytol 155 : 349 – 361 Google Scholar OpenURL Placeholder Text WorldCat Weiner H Stitt M Heldt HW ( 1987 ) Subcellular compartmentation of pyrophosphate and alkaline pyrophosphatase in leaves . Biochim Biophys Acta 893 : 13 – 21 Google Scholar Crossref Search ADS WorldCat Zhang X Myers AM James MG ( 2005 ) Mutations affecting starch synthase III in Arabidopsis alter leaf starch structure and increase the rate of starch synthesis . Plant Physiol 138 : 663 – 674 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the Comisión Interministerial de Ciencia y Tecnología and the European Union-FEDER (grant no. BIO2009–07040), by the Junta de Andalucía (grant nos. P07–CVI–02795 and P09–CVI–4704), and by the Spanish Ministry of Education (Formación de Personal Universitario grant to P.R.). * Address correspondence to [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Ángel Mérida ([email protected]). [W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.113.223420 © 2013 American Society of Plant Biologists. All Rights Reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Schultink, Alex; Cheng, Kun; Park, Yong Bum; Cosgrove, Daniel J.; Pauly, Markus

doi: 10.1104/pp.113.221788pmid: 23893172

Abstract Xyloglucan (XyG) is the dominant hemicellulose present in the primary cell walls of dicotyledonous plants. Unlike Arabidopsis (Arabidopsis thaliana) XyG, which contains galactosyl and fucosyl substituents, tomato (Solanum lycopersicum) XyG contains arabinofuranosyl residues. To investigate the biological function of these differing substituents, we used a functional complementation approach. Candidate glycosyltransferases were identified from tomato by using comparative genomics with known XyG galactosyltransferase genes from Arabidopsis. These candidate genes were expressed in an Arabidopsis mutant lacking XyG galactosylation, and two of them resulted in the production of arabinosylated XyG, a structure not previously found in this plant species. These genes may therefore encode XyG arabinofuranosyltransferases. Moreover, the addition of arabinofuranosyl residues to the XyG of this Arabidopsis mutant rescued a growth and cell wall biomechanics phenotype, demonstrating that the function of XyG in plant growth, development, and mechanics has considerable flexibility in terms of the specific residues in the side chains. These experiments also highlight the potential of reengineering the sugar substituents on plant wall polysaccharides without compromising growth or viability. The cell wall of higher plants represents a composite material consisting of various polymers including cellulose, hemicellulose, lignin, pectin, and glycoproteins (Somerville et al., 2004). The quantity and fine structure of each of these components varies based on the tissue type and plant species (Pauly and Keegstra, 2008). One of the major components of the dicot primary wall (the wall of growing cells) is the hemicellulose xyloglucan (XyG), whose structure and biosynthesis are relatively well described (Zabotina, 2012). The glycan backbone of XyG consists of β-1,4-linked glucosyl residues, which are substituted with a regular pattern of xylosyl residues that can be further decorated with a diverse array of carbohydrate and noncarbohydrate substituents. A one-letter code nomenclature has been developed to specify the substituents of a particular backbone glucosyl residue (Fry et al., 1993). An unsubstituted Glc residue is depicted as G while a Glc substituted with a xylosyl residue is depicted as X. Further substitution of the Xyl with a β-galactosyl or α-arabinofuranosyl-residue is abbreviated as L or S, respectively (Fig. 1). In addition, an L side chain may contain an α-fucosyl moiety on the Gal (abbreviated F) or an acetyl group (underlined L). More than 10 additional side chain structures have been identified in various plant species (Hantus et al., 1997; Jia et al., 2003; Ray et al., 2004; Peña et al., 2008, 2012). Figure 1. Open in new tabDownload slide Xyloglucan motifs present in the walls of Arabidopsis (XXXG, XXLG, and XLXG) and tomato (XSGG) with nomenclature indicated below the structure. AtMUR3 and AtXLT2 are galactosyltransferases required to produce XXLG and XLXG, respectively. Glc, d-Glucopyranose; Xyl, d-xylopyranose; Gal, d-galactopyranose; Ara, l-arabinofuranose. Figure 1. Open in new tabDownload slide Xyloglucan motifs present in the walls of Arabidopsis (XXXG, XXLG, and XLXG) and tomato (XSGG) with nomenclature indicated below the structure. AtMUR3 and AtXLT2 are galactosyltransferases required to produce XXLG and XLXG, respectively. Glc, d-Glucopyranose; Xyl, d-xylopyranose; Gal, d-galactopyranose; Ara, l-arabinofuranose. The analysis of XyG structure is facilitated by the availability of a XyG-specific endoglucanase (XEG) that can release XyG oligosaccharides from plant cell wall preparations (Pauly et al., 1999). When XyG is enzymatically released from the walls of the plant model species Arabidopsis (Arabidopsis thaliana), the oligosaccharides XXXG, XXLG, XXLG, XXFG, XXFG, XLFG, and XLFG are observed (Scheller and Ulvskov, 2010), the structures of several of which are shown in Figure 1. Arabinosylated side chains (S) have not been observed in Arabidopsis walls but are abundant in Solanaceous species such as tomato (Solanum lycopersicum). Tomato XyG consists primarily of the subunits LSGG, XSGG, LXGG, LLGG, and XXGG (Jia et al., 2003, 2005). Unlike the “XXXG” type motif in Arabidopsis, XyG in tomato is less xylosylated, with a repeating “XXGG” type motif. In addition, the glucan backbone of tomato XyG is O-acetylated, a modification that in Arabidopsis is only observed on the side chain galactosyl moiety. The functional significance and genetic basis of these structural differences are not understood. Numerous genes, mainly identified from Arabidopsis, are known to be involved in the biosynthesis of XyG (Pauly et al., 2013). These include a glucan synthase, a member of the Cellulose Synthase-Like C gene family (Cocuron et al., 2007), several XyG xylosyltransferases (XXTs; Faik et al., 2002; Cavalier and Keegstra, 2006), the galactosyltransferases MURUS3 (MUR3; Madson et al., 2003) and XyG Galactose Transferase at Position 2 (XLT2; Jensen et al., 2012), a XyG-specific galacturonosyltransferase (XUT1; Peña et al., 2012), the fucosyltransferase MUR2 (Perrin et al., 1999; Vanzin et al., 2002), and the XyG O-acetyltransferases Altered XyG4 (AXY4) and AXY4-Like (AXY4L; Gille et al., 2011). The Cellulose Synthase-Like C from nasturtium (Tropaeolum majus; Cocuron et al., 2007), where XyG is produced as a seed storage polymer, and MUR3 from eucalyptus (Eucalyptus grandis; Lopes et al., 2010) have both been investigated and appear to have specificities similar to their Arabidopsis orthologs. Glycosyltransferases (GTs) with novel specificities required for the diversity of XyG substitution found in various non-Arabidopsis species, including a XyG arabinosyltransferase, have not been identified to date. XyG figures prominently in many models of the plant cell wall, where it is thought to cross link cellulose microfibrils and have a mechanistic role in cell elongation (Somerville et al., 2004; Hayashi and Kaida, 2011). However, the recent discovery that an Arabidopsis xxt1 xxt2 double mutant lacks detectable XyG but only has relatively minor growth phenotypes questions the structural significance of this polysaccharide (Cavalier et al., 2008; Park and Cosgrove, 2012b). XyG substitution has been shown to influence polymer solubility and binding affinity in vitro, with the enzymatic removal of side chains leading to decreased polymer solubility (Sims et al., 1998; Lima et al., 2004). However, mutants deficient for MUR2, XLT2, or AXY4 show minor, if any, growth phenotypes under laboratory conditions (Vanzin et al., 2002; Gille et al., 2011; Jensen et al., 2012). A point mutant in MUR3 (mur3.1) shows minor growth phenotypes (Madson et al., 2003), while a transfer DNA mutant in MUR3 shows impaired growth and altered Golgi structure (Tamura et al., 2005). The difference between these alleles was attributed to a role of the MUR3 protein interacting with actin to help organize Golgi structure independent from its function as a GT (Tamura et al., 2005). XyG oligosaccharides, which can be liberated by endogenous or exogenous glycosyl hydrolases, have been suggested to have a role in signaling (Aldington et al., 1991), though the biological significance of this is unclear and a specific pathway has not been identified. A complimentary approach to using mutant lines lacking certain XyG substitutions to investigate the function of XyG substitution would be to introduce exogenous side chain structures by the expression of XyG biosynthetic genes from other species. This functional complementation approach requires the identification of the genes responsible for exogenous substitution patterns. The MUR3, XLT2, and XUT1 genes are in the same subclade of the inverting GT family 47 (Li et al., 2004). These three transferases all add β-glycosyl groups to the O2-position of a xylosyl group on XyG but differ in donor specificity, with MUR3 and XLT2 utilizing UDP-Gal and XUT1 utilizing UDP-GalA. The diversity of donor substrate specificity present in this subclade of GTs suggests that similar enzymes may represent good candidate genes for unidentified XyG GTs responsible for XyG side chain diversity in other species. Here, we report the identification of several tomato genes involved in XyG biosynthesis. Candidate genes were constitutively expressed in the Arabidopsis mur3.1 xlt2 double mutant, which contains mostly nonsubstituted XyG, for functional characterization. Two putative XyG arabinofuranosyltransferases were identified, and the evolutionary history of these genes was investigated using phylogenetics. The expression of these genes rescued the growth and petiole extensibility phenotypes of the mutant, demonstrating partial functional redundancy of XyG galactosylation and arabinosylation. RESULTS Phylogenetic Analysis A phylogenetic tree was constructed of the GT family 47 subclade containing MUR3 and XLT2 (Li et al., 2004) using available angiosperm protein sequences from Phytozome and the Medical Plant Genomics Resource (Fig. 2). Nine tomato genes were identified, one of which, Sl09g064470, was most similar to AtMUR3, whereas three (Sl02g092840, Sl07g044960, and Sl07g049610) were most similar to AtXLT2. The four genes were cloned and transformed into the Arabidopsis mur3.1 xlt2 double mutant for ubiquitous expression. Quantitative reverse transcription-PCR was performed on leaf tissue harvested from the T2 generation and indicated that all genes were successfully expressed in the lines analyzed (Supplemental Fig. S1). Figure 2. Open in new tabDownload slide Phylogenetic analysis of the MUR3 and XLT2 subclade of GT family 47. A, Maximum-likelihood tree of Arabidopsis (At) and tomato (Sl) protein sequences. B, The XLT2 clade was expanded with sequences from additional species. The protein sequences are labeled with abbreviations for species (see below) and the gene model (Phytozome) or EST identifier (Medical Plants Genomics Resource). Abel, Atropa belladonna; Alyr, Arabidopsis lyrata; At, Arabidopsis; Brap, Brassica rapa; Cacu, Camptotheca acuminata; Ccle, Citrus clementina; Cpap, Carica papaya; Cros, Catharanthus roseus; Crub, Capsella rubella; Csat, Cannabis sativa; Csin, Citrus sinensis; Dpur, Digitalis purpurea; Dvil, Dioscorea villosa; Egra, Eucalyptus grandis; Epur, Echinacea purpurea; Fves, Fragaria vesca; Gbil, Ginkgo biloba; Gmax, Glycine max; Grai, Gossypium raimondii; Hgor, Hoodia gordonii; Hper, Hypericum perforatum; Lusi, Linum usitatissimum; Mdom, Malus domestica; Mesc, Manihot esculenta; Mgut, Mimulus guttatus; Mtru, Medicago truncatula; Pper, Prunus persica; Pqui, Panax quinquefolius; Ptri, Populus trichocarpa; Pvul, Phaseolus vulgaris; Rcom, Ricinus communis; Roff, Rosmarinus officinalis; Rser, Rauvolfia serpentina; Sl, tomato; Stub, Solanum tuberosum; Tcac, Theobroma cacao; Thal, Thellungiella halophila; Voff, Valeriana officinalis. Figure 2. Open in new tabDownload slide Phylogenetic analysis of the MUR3 and XLT2 subclade of GT family 47. A, Maximum-likelihood tree of Arabidopsis (At) and tomato (Sl) protein sequences. B, The XLT2 clade was expanded with sequences from additional species. The protein sequences are labeled with abbreviations for species (see below) and the gene model (Phytozome) or EST identifier (Medical Plants Genomics Resource). Abel, Atropa belladonna; Alyr, Arabidopsis lyrata; At, Arabidopsis; Brap, Brassica rapa; Cacu, Camptotheca acuminata; Ccle, Citrus clementina; Cpap, Carica papaya; Cros, Catharanthus roseus; Crub, Capsella rubella; Csat, Cannabis sativa; Csin, Citrus sinensis; Dpur, Digitalis purpurea; Dvil, Dioscorea villosa; Egra, Eucalyptus grandis; Epur, Echinacea purpurea; Fves, Fragaria vesca; Gbil, Ginkgo biloba; Gmax, Glycine max; Grai, Gossypium raimondii; Hgor, Hoodia gordonii; Hper, Hypericum perforatum; Lusi, Linum usitatissimum; Mdom, Malus domestica; Mesc, Manihot esculenta; Mgut, Mimulus guttatus; Mtru, Medicago truncatula; Pper, Prunus persica; Pqui, Panax quinquefolius; Ptri, Populus trichocarpa; Pvul, Phaseolus vulgaris; Rcom, Ricinus communis; Roff, Rosmarinus officinalis; Rser, Rauvolfia serpentina; Sl, tomato; Stub, Solanum tuberosum; Tcac, Theobroma cacao; Thal, Thellungiella halophila; Voff, Valeriana officinalis. Effect on XyG structure The XyG structure in several independently transformed lines for each transgene was analyzed by XyG oligosaccharide profiling utilizing matrix-assisted laser-desorption ionization-time-of-flight (MALDI-TOF) mass spectrometry (MS) and high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD; Fig. 3; Supplemental Figs. S2 and S3; Lerouxel et al., 2002). As previously described, the oligosaccharides detected in the mur3.1 xlt2 mutant consisted primarily of XXXG (Jensen et al., 2012). Expression of Sl09g064470 in this mutant resulted in the generation of additional oligosaccharides consistent by mass and retention time with XXLG, XXFG, and minor amounts of XLFG, as well as their O-acetylated forms (Fig. 3; Supplemental Fig. S3). This XyG structure resembles that of the Arabidopsis AtXLT2 single mutant, indicating that Sl09g064470 is functionally equivalent to AtMUR3 and mediates the galactosylation of XXXG on the third xylosyl residue from the nonreducing end. Sl09g064470 was therefore named SlMUR3. Figure 3. Open in new tabDownload slide XyG oligosaccharide profile by MALDI-TOF MS from leaf tissue of the Arabidopsis wild type (WT; ecotype Columbia), the double mutant mur3.1 xlt2, and transgenic lines of mur3.1 xlt2 expressing the indicated genes from tomato. Assignment of oligosaccharide structures by one-letter code (Fig. 1) as described in the text. Figure 3. Open in new tabDownload slide XyG oligosaccharide profile by MALDI-TOF MS from leaf tissue of the Arabidopsis wild type (WT; ecotype Columbia), the double mutant mur3.1 xlt2, and transgenic lines of mur3.1 xlt2 expressing the indicated genes from tomato. Assignment of oligosaccharide structures by one-letter code (Fig. 1) as described in the text. Expression of Sl02g092840 in the Arabidopsis mur3.1 xlt2 mutant did not change the XyG oligosaccharide profile (Fig. 3; Supplemental Fig. S3), suggesting that Sl02g092840 is either inactive, not involved in XyG biosynthesis or the appropriate donor or acceptor substrates are not present in the Arabidopsis double mutant. Expression of either Sl07g044960 or Sl07g049610 resulted in the appearance of an XyG oligosaccharide with a mass-to-charge ratio (m/z) of 1,217 (Fig. 3), consistent with an oligosaccharide containing four pentoses and four hexoses. Linkage analysis of this oligosaccharide (purified by HPAEC) revealed the presence of terminal arabinofuranose, possibly attached to the O2-position of a xylosyl residue, in addition to linkages associated with the XXXG oligosaccharide (Supplemental Fig. S4). To determine which xylosyl residue had the attached arabinofuranose, the purified oligosaccharide was digested with the oligoxyloglucan reducing end-specific xyloglucanbiohydrolase (OREX) enzyme (Bauer et al., 2005), which can hydrolyze XXXG to the cellobiose backbone subunits. This digest resulted in the disappearance of the 1,217 m/z ion and the appearance of a 629 m/z ion (Supplemental Fig. S5), representing an oligosaccharide consisting of two pentoses and two hexoses. This data are consistent with an OREX digestion of XXSG to XX and SG. The XyG oligomer XXSG had been previously isolated and characterized from olive (Olea europaea) pulp (Vierhuis et al., 2001). HPAEC analysis of XXSG prepared from olive pulp has the same retention time and mass as the unusual XyG oligosaccharide found in the mur3.1 xtl2 Arabidopsis mutant expressing Sl07g044960 (Supplemental Fig. S6). In addition, 1H-nuclear magnetic resonance (NMR) was performed on the purified unusual oligosaccharide, and the observed chemical shifts were identical to those of the previously published NMR spectrum for XXSG derived from olive pulp (Fig. 4; Supplemental Fig. S7; Vierhuis et al., 2001). Hence, the addition of either Sl07g044960 or Sl07g049610 to the mur3.1 xlt2 double mutant leads to arabinosylation of XyG in Arabidopsis. The genes were hence named XyG “S”-side chain transferase1 (XST1) and XST2, respectively. The abundance of XXSG was higher in the XST1 lines compared with the XST2 lines (Supplemental Figs. S2 and S3). The XST1 lines also had a minor ion in the MALDI-TOF spectra with an m/z of 1,349, which would be consistent with an XXSG oligosaccharide containing an additional pentose (Fig. 3). Future experiments are needed to establish the nature and position of this additional pentose. Figure 4. Open in new tabDownload slide Structure of the novel oligosaccharide. A, 1H-NMR spectrum of the oligosaccharide with an m/z of 1,217 produced in the XST1 overexpression line. B, Proposed structure of the oligosaccharide (reduced to alditol prior to NMR). Superscripts in the spectrum in A refer to the glycosyl units in B. See Supplemental Figure S5 for detailed assignments. Glc, d-Glucopyranose; Xyl, d-xylopyranose; Gal, d-galactopyranose; Ara, l-arabinofuranose. Figure 4. Open in new tabDownload slide Structure of the novel oligosaccharide. A, 1H-NMR spectrum of the oligosaccharide with an m/z of 1,217 produced in the XST1 overexpression line. B, Proposed structure of the oligosaccharide (reduced to alditol prior to NMR). Superscripts in the spectrum in A refer to the glycosyl units in B. See Supplemental Figure S5 for detailed assignments. Glc, d-Glucopyranose; Xyl, d-xylopyranose; Gal, d-galactopyranose; Ara, l-arabinofuranose. Effect on Plant Morphology and Cell Wall Biomechanics The Arabidopsis mur3.1 xlt2 double mutant exhibits dwarfism compared with wild-type or single mutant plants (Jensen et al., 2012). To investigate how this growth phenotype is affected by alteration of XyG structure the heights of wild-type and mur3.1 xlt2 plants as well as plants from two independently transformed lines for each heterologously expressed tomato gene were measured. Expression of Sl02g092840, which did not affect the XyG structure, resulted in the dwarfism phenotype being retained, while the expression of SlMUR3, XST1, or XST2 resulted in an increase in plant height compared with the nontransgenic double mutant (P ≤ 0.02; Fig. 5; Supplemental Fig. S8). Despite a significant increase, the average heights of plants from XST1- and XST2-expressing lines were less than that of the wild-type plants for three of the four lines examined. This variability may be the result of differential gene expression between the transgenic lines as observed by quantitative PCR analysis (Supplemental Fig. S1) and differing quantities of XyG substitution (Supplemental Fig. S2). Expression of SlMUR3, resulting in a galactosylated, fucosylated XyG, restored plant height completely in both lines examined (Fig. 5B). Figure 5. Open in new tabDownload slide Plant phenotypes and biomechanics. A, Representative 7-week-old Arabidopsis plants for the indicated lines (see Supplemental Fig. S6 for images of all lines). B, Average plant height at 7 weeks with two independent lines for each construct. Error bars indicate sd (n ≥ 4). C, Acid-induced extensibility of leaf petioles under constant force for indicated lines. Error bars show se (n ≥ 9). Asterisk indicates statistically significant difference from the wild type (WT; P < 0.05). Figure 5. Open in new tabDownload slide Plant phenotypes and biomechanics. A, Representative 7-week-old Arabidopsis plants for the indicated lines (see Supplemental Fig. S6 for images of all lines). B, Average plant height at 7 weeks with two independent lines for each construct. Error bars indicate sd (n ≥ 4). C, Acid-induced extensibility of leaf petioles under constant force for indicated lines. Error bars show se (n ≥ 9). Asterisk indicates statistically significant difference from the wild type (WT; P < 0.05). Biomechanical tests on cell wall specimens from leaf petioles of the various lines showed the mur3.1 xlt2 double mutant to be less extensible as assessed by acid-induced creep, which is mediated by the wall-loosening protein α-expansin (Cosgrove, 2000; Sampedro and Cosgrove, 2005; Park and Cosgrove, 2012a). Expression of XST1 restored acid-induced cell wall creep back to wild-type level (Fig. 5C). However, this reduction in extensibility in the mur3.1 xlt2 double mutant and its reversal by XST1 expression were evidently not due to simple changes in mechanical compliance as assessed by stress strain analyses, which was the same in both lines (Supplemental Fig. S9); instead, XyG substitution appeared to be important for the wall-loosening action of endogenous α-expansins. DISCUSSION SlMUR3: A Tomato XyG Galactosyltransferase In vitro activity assays have previously demonstrated that the AtMUR3 protein acts as a XyG galactosyltransferase specific for the third position of the XXXG motif (Madson et al., 2003). Putative orthologs to AtMUR3 can be found in many angiosperm species including nasturtium (Jensen et al., 2012) and eucalyptus, where the MUR3 gene was cloned and shown to functionally complement the Arabidopsis mur3 mutant (Lopes et al., 2010). Unlike Arabidopsis, eucalyptus, and nasturtium, which have XXXG type XyG, tomato XyG contains primarily the XXGG motif. It was therefore unknown if tomato would require a gene with orthologous function to MUR3 for XyG biosynthesis. Through the genetic complementation approach described above, an ortholog to AtMUR3 was identified in tomato (SlMUR3) representing a XyG galactosyltransferase with a similar specificity as AtMUR3. Interestingly, while the XyG structure of the Arabidopsis lines expressing SlMUR3 consisted primarily of XXXG, XXLG, XXFG, and XXFG (consistent with a plant with a defect only in the XLT2), minor amounts of XLFG and XLFG were detected. This may indicate that SlMUR3 also mediates galactosylation of the second position of the XXXG motif, which has not been observed for the AtMUR3 gene. While AtMUR3 has been shown to be active on XXXG, it is unclear if it can act on XXGG. That tomato has retained a gene with MUR3-like activity despite apparently lacking the XXXG motif suggests that SlMUR3 may be able to act on the XXGG motif to form XLGG, a structure that is found in tomato. Alternatively, the XXXG motif may be transiently present in tomato, and a xylosidase may process it to the XXGG form. Future work to test MUR3 activity on various substrates in vitro or a crystal structure of the MUR3 protein bound to substrate would help to reveal the specific recognition motif of this enzyme. XST1 and XST2: Tomato XyG Arabinofuranosyltransferases In addition to the more common substituents found in XyG side chains, such as galactosyl and fucosyl residues, other identified sugar moieties include arabinopyranose (Selaginella kraussiana and Equisetum hyemale), galacturonic acid (Arabidopsis root hair and Physcomitrella patens; Peña et al., 2008, 2012), xylopyranose (Argania spinosa; Ray et al., 2004), and arabinofuranose (Ceratopteris richardii, tomato, and Olea europaea; Vierhuis et al., 2001; Jia et al., 2003; Peña et al., 2008). The differences in XyG structure among these plants are presumably caused by sequence differences in the GTs. The identification of such XyG GTs has been hampered by difficulties in generating or obtaining mutants in nonmodel species as well as a lack of genome sequence information. The recent completion of the tomato genome (Sato et al., 2012) allowed for the identification of candidate XyG arabinofuranosyltransferase genes. The overexpression of either XST1 or XST2 in the Arabidopsis mur3.1 xlt2 mutant background resulted in the production of arabinosylated XyG. While the activity of purified XST1 or XST2 was not demonstrated in vitro, the result that these proteins are sufficient for arabinosylation of XyG in planta and that the in vitro activity of the closely related AtMUR3 protein has been shown suggests that both XST1 and XST2 represent XyG arabinofuranosyltransferases. Interestingly, despite being more closely related to AtXLT2, which acts at the second position of the XXXG motif, both XST1 and XST2 resulted in the addition of arabinofuranose to the third position of the XXXG motif. The diversity of activities of various proteins within the MUR3 subclade of GT47, which can act at the first, second, or third position of the XXXG motif and can add Gal, galacturonic acid, arabinofuranose, and likely additional glycosyl groups, make it an interesting enzyme family to study for insight into the evolution of GT acceptor and donor specificities. Plant GTs believed to utilize UDP-arabinofuranose have previously been identified from the GT47 (Harholt et al., 2006), GT61 (Anders et al., 2012), and GT77 (Egelund et al., 2007; Gille et al., 2009) families, indicating that this donor specificity has evolved multiple times. Origin of XST1 and XST2 A phylogenetic tree of approximately 400 angiosperm protein sequences from the GT47 family subclade including MUR3 and XLT2 (Li et al., 2004) suggests a specific subgroup of proteins (XST clade, which includes both XST1 and XST2; Fig. 2B) could represent XyG arabinofuranosyltransferases. This putative XST clade contains proteins from species within the Solanales, Lamiales, and Gentianales orders, all of which contain species with arabinosylated XyG (in Lamiales, O. europaea [Vierhuis et al., 2001] and in Gentianales, Nerium oleander [Hoffman et al., 2005]). While Gentianales and Lamiales species contain putative orthologs to AtXLT2, such a gene was not identified in any of the Solanales species. The absence of an AtXLT2 ortholog in the Solanales is consistent with the finding that none of the tomato genes expressed in the mur3.1 xlt2 mutant complemented the defect in XLT2 by resulting in the addition of a galactosyl residue to the middle xylosyl position of the XXXG motif. Together this suggests that a gene with XST activity arose following a gene duplication event in a common ancestor to the Lamiales, Gentianales, and Solanales and that the original XLT2 gene was subsequently lost in the Solanales lineage. Implications for the Function of XyG Substitution The mur3.1 xlt2 double mutant has approximately a 30% decrease in plant height at maturity and reduced cell wall creep. These phenotypes can be partially or fully rescued by expression of SlMUR3, XST1, or XST2. This demonstrates that for the function of XyG, the presence of a glycosyl substituent on the xylosyl residues is more important than the identity of that substituent. Previous studies showed that the enzymatic removal of side chains from XyG polysaccharides results in the formation of aggregates in vitro (Sims et al., 1998), demonstrating that XyG substitution can affect polymer solubility. XyG substitution may therefore help regulate the solubility of the XyG polysaccharide during biosynthesis and transport while still allowing for appropriate interactions with other polymers once deposited in the apoplast. Substitution is also a factor in XyG binding to cellulose and affects cellulose crystallinity (Lima et al., 2004; Whitney et al., 2006). Both of these processes affect cell wall structure and likely resulted in a wall with altered sensitivity to α-expansin-mediated creep, as seen in Figure 5C and also found previously in the xxt1/xxt2 line entirely lacking XyG (Park and Cosgrove, 2012a). CONCLUSION Identifying the genetic basis for and the functional significance of the structural diversity of plant cell wall polysaccharides remains a significant challenge due to the complex nature of their structures and the large number of putative carbohydrate active enzymes present in the typical plant genome, the majority of which are uncharacterized. When available, genetic approaches can be used to create mutants with reduced polysaccharide complexity. These mutants can then be used as platforms for the heterologous expression of GTs for identification and characterization as demonstrated here. Additionally, this approach allows for study of the in vivo function of particular cell wall structures. Plant cell wall materials are important for a variety of applications including fuels, feed, food, textiles, and paper (Mishra and Malhotra, 2009), but native wall structures may not be ideal for these uses. Plant engineering approaches as presented here can allow for the tailoring of polysaccharide structures to produce glycans and wall materials with properties uniquely suited for particular applications. The naturally occurring diversity of wall components found in various plant species and tissues demonstrates the potential for altering wall structures for these purposes without detriment. MATERIALS AND METHODS Gene Identification and Phylogenetic Analysis A BLAST (Altschul et al., 1997) search was performed using the Arabidopsis (Arabidopsis thaliana) protein sequences from the MUR3 and XLT2 subclade of GT family 47 to identify homologous genes from tomato (Solanum lycopersicum; Bombarely et al., 2011). Additional sequences were obtained from Phytozome (http://www.phytozome.net) and the Medicinal Plant Genomics Resource (http://medicinalplantgenomics.msu.edu/). A multiple sequence alignment was performed using Clustal (Sievers et al., 2011) and a maximum-likelihood tree was constructed using PhyML (Gouy et al., 2010; Guindon et al., 2010; Fig. 1). Cloning and Transformation The coding sequences of the GT47 candidate genes were amplified from genomic DNA using PCR with the primers from Supplemental Table S1 and cloned into the pCR8 TOPO TA vector with TA cloning (Life Technologies). LR reactions were performed to move the genes into a Gateway-compatible version of the plant overexpression plasmid pORE E4 (Coutu et al., 2007). These plasmids were transformed into Agrobacterium tumefaciens strain GV3101, which was subsequently used to transform the mur3.1 xlt2 Arabidopsis double mutant using the dip infiltration method (Clough and Bent, 1998). Transformed Arabidopsis plants were selected by plating the resulting seeds on one-half-strength Murashige and Skoog (Murashige and Skoog, 1962), 1% (w/v) Suc, and 60 µg mL–1 kanamycin agar plates. After 2 weeks, resistant plants were moved to soil and genotyped to confirm the presence of the transgene. Xyloglucan Analysis Analysis of XyG structure by XEG digestion of cell wall material followed by MALDI-TOF MS as previously described (Jensen et al., 2012). Purification of the XXSG Oligosaccharide To obtain large amounts of the initially unknown XyG oligosaccharide, 1-week-old etiolated hypocotyls were dried, ground, and extracted three times each with 70% (v/v) ethanol and chloroform:methanol (1:1, v/v). The pellet, approximately 150 mg, was then extracted with 4 m potassium hydroxide containing 10 mm sodium borohydride for 4 h under vigorous shaking. The supernatant was removed and neutralized with acetic acid and hydrochloric acid. Polysaccharides were precipitated from the neutralized extract by adding ethanol to 70% (v/v) and incubation of the solution at –20°C overnight. The precipitated material was washed four times with 70% (v/v) ethanol to remove remaining salts. The pellet was subsequently dried and digested with 4 mL of 50 mm ammonium formate, pH 4.5, containing 16 Units XEG. The supernatant from the digest was dried, and the oligosaccharides were reduced by incubation with 200 µL 1 m ammonium hydroxide containing 10 mg mL–1 sodium borohydride for 1 h. The reaction was neutralized with acetic acid followed by three additions and evaporations of methanol:acetic acid (9:1, v/v) and four additions of methanol. The reduced oligosaccharides were resuspended in 400 µL water and then separated using HPAEC-PAD. The fraction containing the dominant novel oligosaccharide (pooled from multiple injections) was desalted using a SupelClean ENVI-Carb 57109-U reversed-phase column (Supelco). The column was equilibrated with 50% (v/v) acetonitrile and washed four times with water. The oligosaccharide was bound to the column and then washed with water until the flow through showed neutral pH. The oligosaccharide was eluted from the column with 50% (v/v) acetonitrile and dried. NMR The purified oligosaccharide was dissolved in D2O (deuterium 99.99%), freeze dried, and dissolved in 0.3 mL D2O (deuterium 99.96%) with 3-(Trimethylsilyl)-1-propanesulfonic acid internal standard (0.01 mg mL–1). 1H-NMR spectra were recorded on a Bruker AVANCE 600 MHz NMR spectrometer equipped with an inverse gradient 5-mm TXI 1H/13C/15N CryoProbe at 285 K and 315 K, respectively. A two-dimensional Total Correlation Spectroscopy (TOCSY) NMR experiment was recorded using standard Bruker pulse program “dipsi2ph” for additional peak assignments with the following parameters: 2048 × 256 data points for F2 × F1 dimensions, 48 scans; four dummy scans; interscan delay of 1.5 s; prescan delay of 69.6 µs; and mixing time of 200 ms. All chemical shifts were referenced relative to 3-(Trimethylsilyl)-1-propanesulfonic acid (0.00 ppm for 1H). The NMR data processing and analysis were performed using Bruker’s Topspin 3.1 software. Plant Growth Conditions and Phenotyping The mur3.1 xlt2 mutant was generated as described in Jensen et al. (2012). Arabidopsis plants were grown on soil in a Percival growth chamber at 22°C with a repeating cycle of 16-h light and 8-h dark. Six-week-old plants were used for analysis of the growth phenotype. Plant height was measured from the base of the stem to the most distal part of the plant. Wall Extension (Creep) Measurement Eight-millimeter sections from the middle of petioles of the 4-week-old wild type and mutants were collected and prepared for an acid-induced wall extension (creep) assay and a stress strain assay as described (Park and Cosgrove, 2012a). For the creep assays, the clamped segments were initially incubated in neutral buffer (250 µL of 20 mm HEPES, pH 6.8) for 30 min with a constant load of 7.5 g. The incubation buffer was then replaced with 250 µL of 20 mm sodium acetate (pH 4.5) containing 5 mm dithiothreitol. Wall creep extension was measured with a position transducer, and the data were recorded every 30 s. Sequence data referred to in this article can be found in the GenBank/EMBL data libraries under the following accession numbers: SlMUR3 (Sl09g064470; XP_004247129), XST1 (Sl07g044960; XP_004243615), XST2 (Sl07g049610; XP_004243499), Sl02g092840 (XP_004231889). Supplemental Data The following materials are available in the online version of this article. Supplemental Figure S1. Quantification of transgene expression. Supplemental Figure S2. XyG oligosaccharide mass profiles of independent transformants. Supplemental Figure S3. HPAEC-PAD analysis of XEG-released XyG oligosaccharides. Supplemental Figure S4. Glycosidic linkage analysis of the purified oligosaccharide. Supplemental Figure S5. OREX digestion of the purified oligosaccharide. Supplemental Figure S6. HPAEC-PAD analysis of XEG-released xyloglucan oligosaccharides from olive and the Arabidopsis mur3.1 xlt2 double mutant expressing XST1. Supplemental Figure S7. 1H NMR peak assignments for the purified oligosaccharide. Supplemental Figure S8. Representative pictures of Arabidopsis plants with altered xyloglucan structures. Supplemental Figure S9. Extensibility of leaf petioles. Supplemental Table S1. Primer sequences. ACKNOWLEDGMENTS We thank Kirk Schnorr (Novozymes) for the generous gift of the XEG and Stefan Bauer (Energy Biosciences Institute, UC Berkeley) for a Pichia pastoris culture expressing the OREX enzyme. 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The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Markus Pauly ([email protected]). [W] The online version of this article contains Web-only data. [OPEN] Articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.113.221788 © 2013 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2013. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Lovegrove, Alison; Wilkinson, Mark D.; Freeman, Jackie; Pellny, Till K.; Tosi, Paola; Saulnier, Luc; Shewry, Peter R.; Mitchell, Rowan A.C.

doi: 10.1104/pp.113.222653pmid: 23878080

Abstract The cell walls of wheat (Triticum aestivum) starchy endosperm are dominated by arabinoxylan (AX), accounting for 65% to 70% of the polysaccharide content. Genes within two glycosyl transferase (GT) families, GT43 (IRREGULAR XYLEM9 [IRX9] and IRX14) and GT47 (IRX10), have previously been shown to be involved in the synthesis of the xylan backbone in Arabidopsis, and close homologs of these have been implicated in the synthesis of xylan in other species. Here, homologs of IRX10 TaGT47_2 and IRX9 TaGT43_2, which are highly expressed in wheat starchy endosperm cells, were suppressed by RNA interference (RNAi) constructs driven by a starchy endosperm-specific promoter. The total amount of AX was decreased by 40% to 50% and the degree of arabinosylation was increased by 25% to 30% in transgenic lines carrying either of the transgenes. The cell walls of starchy endosperm in sections of grain from TaGT43_2 and TaGT47_2 RNAi transgenics showed decreased immunolabeling for xylan and arabinoxylan epitopes and approximately 50% decreased cell wall thickness compared with controls. The proportion of AX that was water soluble was not significantly affected, but average AX polymer chain length was decreased in both TaGT43_2 and TaGT47_2 RNAi transgenics. However, the long AX chains seen in controls were absent in TaGT43_2 RNAi transgenics but still present in TaGT47_2 RNAi transgenics. The results support an emerging picture of IRX9-like and IRX10-like proteins acting as key components in the xylan synthesis machinery in both dicots and grasses. Since AX is the main component of dietary fiber in wheat foods, the TaGT43_2 and TaGT47_2 genes are of major importance to human nutrition. Xylan is a hemicellulosic component of cell walls and one of the most abundant polysaccharides in nature (Ebringerova et al., 2005; Scheller and Ulvskov, 2010). Its prevalence and structure differ markedly between dicots and grasses; in the former, it constitutes only about 5% of the polysaccharide of primary cell walls, whereas it is typically 30% of the polysaccharide of grass primary cell walls (Carpita, 1996; Scheller and Ulvskov, 2010). Arabinofuranose (Araf) substitution of the xylan backbone is more common in grasses than in dicots, and some of the Araf sugars are ester linked to ferulic acid, which confers a cross-linking functionality to the xylan chains that is absent in dicots. Furthermore, the reducing end of dicot xylan has a characteristic 4-β-d-Xylp-(1→4)-β-d-Xylp-(1→3)-α-l-Rhap(1→2)-α-d-GalpA-(1→4)-d-Xylp oligosaccharide that has not been detected in xylans from grasses (Pena et al., 2007; York and O’Neill, 2008). The candidate genes responsible for xylan synthesis in Arabidopsis (Arabidopsis thaliana) have been identified by studies of knockout mutants in glycosyl transferase (GT) families. Plants carrying mutations in the GT43 family genes IRREGULAR XYLEM9 (IRX9) and IRX14 and in the GT47 family gene IRX10 all have decreased xylan content, with the remaining xylan having a shorter chain length compared with the wild type but the reducing end oligosaccharide still being present (Brown et al., 2007, 2009; Pena et al., 2007; Wu et al., 2009). Closely related genes exist for all these (IRX9-L, IRX10-L, and IRX14-L) but are less expressed and are functionally redundant with the more highly expressed counterpart (Brown et al., 2009; Wu et al., 2010). It is unclear how the products from the very different GT43 and GT47 gene families, which appear to be equally essential in Arabidopsis, cooperate to synthesize the xylan backbone. By overexpressing IRX9 and IRX14 together in tobacco (Nicotiana tabacum) BY2 cells, xylan xylosyl transferase activity was stimulated, suggesting that they encode the key proteins mediating the activity, although an endogenous IRX10 ortholog would presumably also be present in these cells (Lee et al., 2012). On the other hand, abundant xylan is synthesized in psyllium (Plantago afra) seed mucilage, which has abundant transcripts of an IRX10 ortholog but only very low levels of transcripts of IRX9 and IRX14 orthologs (Jensen et al., 2011). It is possible that different xylan synthetic mechanisms exist, for example between grasses and dicots (York and O’Neill, 2008). There is less direct evidence for the genes involved in xylan backbone synthesis in grasses. Recently, a rice (Oryza sativa) line carrying a mutation in the ortholog of IRX10 was shown to have decreased xylan content, although the decrease was modest (10% decrease in cell wall Xyl). The mutation also resulted in smaller stature and increased ease of biomass saccharification (Chen et al., 2013). A microsomal complex isolated from wheat (Triticum aestivum) seedlings that exhibited all the main activities required for synthesizing glucuronoarabinoxylan (i.e. xylan backbone extension and addition of Araf and GlcA sugars to the backbone) was shown to contain three wheat proteins that were homologous to the GT43 IRX14, the GT47 IRX10, and a GT75 protein (Zeng et al., 2010). This GT75 protein is orthologous to OsUAM1 and presumably has the same UDP-Ara mutase activity found for the protein in rice (Konishi et al., 2007), converting the supplied UDP-Arap into the UDP-Araf found in the products. Since the observed xylan GlcA transferase activity is retaining and the three identified proteins are all from inverting families, other active proteins must be present. In Arabidopsis, xylan GlcA transferase proteins from GT8 have been shown to confer this activity (Mortimer et al., 2010). Also, GT61 proteins have been shown to be xylan arabinosyl transferases in grasses, including wheat (Anders et al., 2012). While only a IRX14 homolog was detected in this complex, of great interest is whether a IRX9 homolog is also required in grasses for xylan backbone extension, as appears to be the case in woody and herbaceous dicots (Brown et al., 2007; Pena et al., 2007; Lee et al., 2011, 2012). The wheat starchy endosperm cell wall has been established as an excellent system for studying grass cell walls due to the prevalence of arabinoxylan (AX) and 1,3;1,4-β-d-glucan, which constitute approximately 70% and 25% of the total cell wall polysaccharide, respectively. Using a strong starchy endosperm-specific promoter to drive RNA interference (RNAi) to suppress specific transcripts, we have demonstrated the key role of TaCSLF6 in synthesizing 1,3;1,4-β-d-glucan (Nemeth et al., 2010) and that the GT61 gene TaXAT1 is responsible for nearly all the monosubstituted Ara in AX in endosperm (Anders et al., 2012). The amount and structure of AX in wheat starchy endosperm are also of great practical importance; as it is the main component of dietary fiber in wheat foods, it is a major contributor of dietary fiber in the human diet (Topping, 2007), and it also affects the processing properties of wheat flour for different end uses (Saulnier et al., 2007). The most abundant transcripts for the genes implicated in xylan backbone extension (xylan synthases) in wheat starchy endosperm are TaGT47_2, TaGT43_1, and TaGT43_2, which are homologous to IRX10, IRX14, and IRX9, respectively, in Arabidopsis (Pellny et al., 2012). In fact, GT47_2 is the most abundantly expressed of all GT genes in this cell type (excluding the genes involved in starch synthesis), in keeping with the dominance of AX in the cell wall. Here, we report the effects of specifically suppressing the expression of TaGT47_2 and TaGT43_2 by RNAi in transgenic wheat lines on the amount and structure of AX and the wheat starchy endosperm cell wall. The effects of the suppression are similar for these two diverse genes, suggesting an analogous mechanism for xylan extension in wheat to that in Arabidopsis. RESULTS Phylogenetic trees for predicted protein sequences for the whole GT43 family and for the IRX10 clade of the GT47 family from the fully sequenced genomes of Arabidopsis, poplar (Populus trichocarpa), rice, Brachypodium distachyon, Physcomitrella patens, and Selaginella moellendorffii (www.phytozome.org; Goodstein et al., 2012) and from wheat transcripts present in the starchy endosperm (Pellny et al., 2012) are shown in Figure 1. Three rice genes in the GT43 family are indicated that have recently been demonstrated to be functional orthologs of IRX9, IRX9-L, and IRX14, being able to complement mutations in these Arabidopsis genes (Chiniquy et al., 2013). While there is considerable diversity between the IRX9 and IRX14 homologs, the IRX10 homologs are highly conserved (note the different scale bars for the two trees). The two wheat genes studied here are the IRX9 homolog TaGT43_2 and the IRX10 homolog TaGT47_2. Figure 1. Open in new tabDownload slide Phylogenetic trees of the GT43 family and the IRX10 clade within the GT47 family. Trees were derived from protein alignments of all genes of fully sequenced plants plus wheat genes expressed in starchy endosperm. Lower plants are represented by S. moellendorffii (dark red) and P. patens (orange); monocots by rice (dark green), B. distachyon (light green), and wheat genes (black); and dicots by Arabidopsis (dark blue) and poplar (light blue). Figure 1. Open in new tabDownload slide Phylogenetic trees of the GT43 family and the IRX10 clade within the GT47 family. Trees were derived from protein alignments of all genes of fully sequenced plants plus wheat genes expressed in starchy endosperm. Lower plants are represented by S. moellendorffii (dark red) and P. patens (orange); monocots by rice (dark green), B. distachyon (light green), and wheat genes (black); and dicots by Arabidopsis (dark blue) and poplar (light blue). Bread wheat is a hexaploid with three related genomes (A, B, and D). The term “gene,” therefore, is used to encompass the three homeologous forms from the three genomes; these typically have 95% to 97% nucleotide identity of transcripts and can be assumed to have the same molecular function. The recent availability of the chromosome-sorted genomic survey sequences from the International Wheat Genome Sequencing Consortium (www.wheatgenome.org) makes it possible to unequivocally assign the three variants in complementary DNA sequence that we identify for each of the two genes to chromosomes. Therefore, TaGT43_2 and TaGT47_2 genes can be assigned to chromosomes 4 and 3 (i.e. 4A, 4B, 4D and 3A, 3B, 3D), respectively, with all three homeologs of both being expressed in starchy endosperm. The transcript abundances differ somewhat between the three homeologs but all show the same pattern through endosperm development for both genes (Fig. 2). These sequences and the RNAi constructs designed to suppress all three variants are shown in Supplemental Figures S1 and S2. The RNAi construct for TaGT47_2 had sufficient identity to potentially also suppress the very similar TaGT47_1 and TaGT47_4 genes (whose longest identical fragments are 29 and 38 bp, respectively), whereas the RNAi construct for TaGT43_2 was specific for this gene. Figure 2. Open in new tabDownload slide Transcript abundance of TaGT43_2 and TaGT47_2 in developing starchy endosperm. Estimates were based on reanalysis of transcript abundance for three homeologs of TaGT43_2 and TaGT47_2 from RNA-Seq libraries (Pellny et al., 2012). Figure 2. Open in new tabDownload slide Transcript abundance of TaGT43_2 and TaGT47_2 in developing starchy endosperm. Estimates were based on reanalysis of transcript abundance for three homeologs of TaGT43_2 and TaGT47_2 from RNA-Seq libraries (Pellny et al., 2012). We identified six wheat transgenic lines carrying the TaGT43_2 RNAi transgene and six lines carrying the TaGT47_2 RNAi. All the TaGT43_2 RNAi lines had clear effects on the amounts of arabinoxylan oligosaccharide (AXOS) released by xylanase digestion in samples from T1 grain, but three of the GT47_2 RNAi lines did not show any clear effects on AXOS in T1 samples and were not pursued further (data not shown). Such an absence of effect is sometimes encountered in cereal transformants due to the partial integration of transgenes or integration into a nonexpressed part of the genome. Data are presented for four GT43_2 RNAi lines and the three GT47_2 RNAi lines that had an effect; all these exhibited a segregation of transgenes consistent with a single insertion site. For all measurements, samples for homozygous transgenic plants were compared with corresponding azygous null segregants from the same line. Transcript abundance within the developing starchy endosperm of the transgenic lines was determined by quantitative reverse transcription-PCR, showing strong suppression of the target genes in all four GT43_2 RNAi lines and the three GT47_2 RNAi lines (Fig. 3). However, there was also suppression of the closely related GT47_1 gene in the GT47_2 RNAi lines and some evidence of GT47_4 suppression. The phenotypes presented for these GT47_2 RNAi lines, therefore, may partially result from the suppression of GT47_1 and GT47_4, although these genes are expressed at only approximately 10% of the level of the GT47_2 genes in starchy endosperm (Pellny et al., 2012). These three very similar genes appear to have arisen from duplication after divergence from the common ancestor with IRX10, and it is not possible to say which of them is the true IRX10 ortholog (Fig. 1). Figure 3. Open in new tabDownload slide Effects of RNAi transgenes on endogenous transcript abundance. Transcript abundance was estimated by quantitative reverse transcription-PCR from pure starchy endosperm dissected at 21 DPA. Error bars represent se, and stars denote significant differences at P < 0.05. A, Four replicate samples from homozygous plants carrying the TaGT43_2 RNAi transgene for lines 3, 4, 5, and 6 (H3, H4, H5, and H6) or the corresponding azygous controls (A3, A4, A5, and A6). B, As in A, but for the TaGT47_2 RNAi lines 1, 4, and 6. Figure 3. Open in new tabDownload slide Effects of RNAi transgenes on endogenous transcript abundance. Transcript abundance was estimated by quantitative reverse transcription-PCR from pure starchy endosperm dissected at 21 DPA. Error bars represent se, and stars denote significant differences at P < 0.05. A, Four replicate samples from homozygous plants carrying the TaGT43_2 RNAi transgene for lines 3, 4, 5, and 6 (H3, H4, H5, and H6) or the corresponding azygous controls (A3, A4, A5, and A6). B, As in A, but for the TaGT47_2 RNAi lines 1, 4, and 6. Monosaccharide analyses of nonstarch polysaccharides (NSP; composed of cell wall polysaccharides, oligosaccharides, and arabinogalactan peptide) from white flour showed that suppression of TaGT43_2 and TaGT47_2 decreased the total amount of NSP Xyl, which is virtually all in the AX backbone (Ordaz-Ortiz and Saulnier, 2005), by 45% and 48%, respectively, averaged across three lines (Fig. 4A). There was an increase in the ratio of AX Ara to Xyl in the transgenic lines, indicating greater Ara substitution of the remaining Xyl residues (Fig. 4B). Thus, total AX was decreased by 40% and 43% (Fig. 4C). Similar effects were observed for the water-unextractable portion of AX (Fig. 4D), showing that there was no overall effect on the solubility of AX, with water-extractable (WE) AX being about 35% of total AX in all lines. There was a small increase in the amount of Glc in the NSP in all of the transgenic lines relative to the controls (Fig. 4E). Although this was not significant when averaged across the three lines for each gene, there was a clear trend, with the lines showing the greatest decrease in AX also having the greatest increase in NSP Glc (Fig. 4F). However, this only had a minor influence on the total amount of cell wall polysaccharide, the lines with a decrease of approximately 50% AX having an increase in Glc of approximately 20%, which would give an overall decrease in cell wall sugars of approximately 30%. There was no significant trend in the amount of Man that was present in the NSP, suggesting that glucomannan was not affected (Supplemental Table S1). Figure 4. Open in new tabDownload slide Monosaccharide analyses of NSP fractions from white flour. All data are from samples from three TaGT43_2 RNAi lines (lines 1, 3, and 6) and three TaGT47_2 RNAi lines (lines 1, 4, and 6). A to E, Average of azygous controls (A) and homozygous transgenics (H) for three lines; error bars represent se, and asterisks denote significant differences at P < 0.05 from Student’s t test. A, Total Xyl, assumed to be all in AX. B, Ratio of Ara to Xyl in AX. Ara in AX estimated as (Ara \x{2013} 0.7 × Gal), since it is assumed all Gal is in AGP, with AGP Ara/Gal = 0.7 (Ordaz-Ortiz and Saulnier, 2005). C, Total AX, estimated as Xyl + Ara in AX, estimated as in B. D, Water-unextractable (WU) AX, estimated as for total AX, using monosaccharide values from the water-unextractable fraction. E, Total Glc. F, Effect of transgenes on total AX versus effect on NSP Glc, expressed as values for transgenics relative to corresponding controls in individual lines. Gray symbols represent TaGT43_2 lines, and white symbols represent TaGT47_2 lines. Figure 4. Open in new tabDownload slide Monosaccharide analyses of NSP fractions from white flour. All data are from samples from three TaGT43_2 RNAi lines (lines 1, 3, and 6) and three TaGT47_2 RNAi lines (lines 1, 4, and 6). A to E, Average of azygous controls (A) and homozygous transgenics (H) for three lines; error bars represent se, and asterisks denote significant differences at P < 0.05 from Student’s t test. A, Total Xyl, assumed to be all in AX. B, Ratio of Ara to Xyl in AX. Ara in AX estimated as (Ara \x{2013} 0.7 × Gal), since it is assumed all Gal is in AGP, with AGP Ara/Gal = 0.7 (Ordaz-Ortiz and Saulnier, 2005). C, Total AX, estimated as Xyl + Ara in AX, estimated as in B. D, Water-unextractable (WU) AX, estimated as for total AX, using monosaccharide values from the water-unextractable fraction. E, Total Glc. F, Effect of transgenes on total AX versus effect on NSP Glc, expressed as values for transgenics relative to corresponding controls in individual lines. Gray symbols represent TaGT43_2 lines, and white symbols represent TaGT47_2 lines. AX structure was characterized using simultaneous digestion of NSP preparations of white flour by xylanase and lichenase and analysis of the resulting oligosaccharides as described previously (Ordaz-Ortiz et al., 2005; Nemeth et al., 2010; Pellny et al., 2012). Large decreases (significant at P < 0.001) were observed in the abundances of AXOS in all of the transgenic lines (Fig. 5A). However, the magnitude of effects differed between individual fragments, with unsubstituted Xyl and xylobiose (X and XX) being affected less than substituted AXOS, and XA2+3XX being affected slightly less than other substituted AXOS. The effects on glucan oligosaccharides (G3 and G4) released by lichenase digestion were more variable: in some lines, significant increases were observed, while in others, no effect or small decreases were seen (Fig. 5B); these variable effects were much the same as those for NSP Glc (Fig. 5D), indicating that the effects on Glc are largely or solely due to changes in the 1,3;1,4-β-d-glucan from which the G3 and G4 oligosaccharides are derived. The magnitude of the effects on AXOS across the lines was correlated with the size of the effect on total AX estimated from monosaccharide analysis, with correlation coefficients of 0.83, 0.87, and 0.90 for X, XA3XX, and XA2+3XX, respectively. However, for all lines, the relative decrease in AXOS peaks was greater than the decrease in total AX measured as a monosaccharide (e.g. a line with approximately 50% decrease in total AX had a decrease of approximately 90% in XA3XX; Fig. 5D). Virtually all the water-unextractable portion of AX was solubilized by xylanase action, and this was unchanged in the transgenics (Supplemental Table S1). However, our high-performance anion-exchange chromatography (HPAEC) methodology measures the abundance of AXOS containing up to nine pentoses, accounting for about 75% of the total in wild-type wheat, but matrix-assisted laser-desorption ionization (MALDI)-mass spectrometry (MS) analysis of digested AX shows that in transgenic samples, this fraction was much smaller (Fig. 5E), explaining the greater decrease in measured AXOS. This is presumably caused by the greater arabinosylation of AX in transgenic samples (Fig. 4B), which results in fewer consecutive unsubstituted Xyl residues that are required for GH11 xylanase cleavage. Figure 5. Open in new tabDownload slide Relative abundance of oligosaccharides released by simultaneous digestion of endosperm NSP samples by xylanase and lichenase. A to D, Oligosaccharide abundance from HPAEC peak area for transgenics relative to corresponding controls in individual lines. A and B, Relative abundance (average of three biological replicates ± se). Solid colors represent GT43_2 lines, and striped colors represent GT47_2 lines. Columns are sorted by decreasing relative abundance of XA3XX: GT43_2 lines 2, 6, and 1, GT47_2 lines 1 and 6, GT43_2 line 3, and GT47_2 line 4. A, AXOS. Nomenclature is from Faure et al. (2009). Columns are colored according to substitution with Araf: unsubstituted (green), monosubstituted only (red), disubstituted only (blue), and monosubstituted and disubstituted (purple). B, (1,3);(1,4)-β-Glucan oligosaccharides [G3, (Glc)3; G4, (Glc)4]. C and D, Comparison of relative oligosaccharide and monosaccharide abundances for individual lines. Colored symbols represent GT43_2 lines, and white symbols represent GT47_2 lines. C, Relative total AX abundance from monosaccharide analyses versus relative AXOS abundance released by xylanase. Squares represent X, triangles represent XA3XX, and circles represent XA2+3XX; colors are as in A. D, Relative NSP Glc abundance versus relative (1,3);(1,4)-β-glucan oligosaccharide abundance. Squares represent G3, and triangles represent G4. E, Relative distribution of AXOS released by xylanase estimated by MALDI-MS. Number of pentoses (DP) was identified from mass-to-charge ratio = DP × 132.114 + 18.016 + 23. Data are from transgenic and corresponding null samples of GT43_2 line 3. Peaks at DP 11, 16, and 27 are omitted because of overlap with hexose oligosaccharides; AXOS of DP < 5 was not analyzed. The dotted line indicates DP below which AXOS was analyzed by the HPAEC method. Figure 5. Open in new tabDownload slide Relative abundance of oligosaccharides released by simultaneous digestion of endosperm NSP samples by xylanase and lichenase. A to D, Oligosaccharide abundance from HPAEC peak area for transgenics relative to corresponding controls in individual lines. A and B, Relative abundance (average of three biological replicates ± se). Solid colors represent GT43_2 lines, and striped colors represent GT47_2 lines. Columns are sorted by decreasing relative abundance of XA3XX: GT43_2 lines 2, 6, and 1, GT47_2 lines 1 and 6, GT43_2 line 3, and GT47_2 line 4. A, AXOS. Nomenclature is from Faure et al. (2009). Columns are colored according to substitution with Araf: unsubstituted (green), monosubstituted only (red), disubstituted only (blue), and monosubstituted and disubstituted (purple). B, (1,3);(1,4)-β-Glucan oligosaccharides [G3, (Glc)3; G4, (Glc)4]. C and D, Comparison of relative oligosaccharide and monosaccharide abundances for individual lines. Colored symbols represent GT43_2 lines, and white symbols represent GT47_2 lines. C, Relative total AX abundance from monosaccharide analyses versus relative AXOS abundance released by xylanase. Squares represent X, triangles represent XA3XX, and circles represent XA2+3XX; colors are as in A. D, Relative NSP Glc abundance versus relative (1,3);(1,4)-β-glucan oligosaccharide abundance. Squares represent G3, and triangles represent G4. E, Relative distribution of AXOS released by xylanase estimated by MALDI-MS. Number of pentoses (DP) was identified from mass-to-charge ratio = DP × 132.114 + 18.016 + 23. Data are from transgenic and corresponding null samples of GT43_2 line 3. Peaks at DP 11, 16, and 27 are omitted because of overlap with hexose oligosaccharides; AXOS of DP < 5 was not analyzed. The dotted line indicates DP below which AXOS was analyzed by the HPAEC method. Since Arabidopsis mutants lacking functional IRX9 or IRX10 have decreased glucuronoxylan chain length (Brown et al., 2007, 2009; Pena et al., 2007), we determined the distribution of WE-AX chain size by measuring the concentration and intrinsic viscosity ([η]) profiles of WE-AX separated by size-exclusion HPLC. The concentration profiles for samples from two homozygous transgenic lines each of GT43_2 RNAi (lines 3 and 5) and GT47_2 RNAi (lines 1 and 4) showed shifts to higher retention volumes compared with azygous controls, indicating that, in all cases, a greater fraction of the WE-AX was made up of small molecules (Fig. 6). This reflects a shift in M r distribution, which depends both on chain length and Ara substitution, but [η] shows a close direct relationship with AX chain length, as log [η] is linearly related to log chain length (Dervilly-Pinel et al., 2001). The profiles of the transgenic samples show much greater relative abundance of AX at low [η], consistent with a shift to shorter AX chains (Fig. 6) and resulting in a pronounced decrease in average [η] (Table I). However, comparison of the log [η] value in the control and transgenic plants shows different effects between the GT43_2 RNAi and GT47_2 RNAi constructs. In particular, the peak in log [η] of about 2.8 to 2.9 in the controls is shifted to a shoulder at 2.5 to 2.6 in the GT43_2 RNAi transgenics (Fig. 6, A and B) but not in the GT47_2 RNAi transgenics (Fig. 6, C and D). Based on the relationship between log [η] and log chain length reported by Dervilly-Pinel et al. (2001), the effect of the GT43_2 RNAi construct corresponds to a decrease in AX chain length at this peak/shoulder from 1,900 to 2,300 down to 750 to 950 Xyl residues. Figure 6. Open in new tabDownload slide Profiles of concentration and [η] from size-exclusion chromatography for WE-AX from mature endosperm of transgenic and control wheat lines. Left panels show original profiles against retention volume, with void volume (Vo) and total volume (Vt) of the column indicated. Right panels show log [η], which is linearly related to log AX chain length, versus concentration. Data are shown for TaGT43_2 RNAi lines 3 and 5 and for TaGT47_2 RNAi lines 1 and 4 azygous control (A) and homozygous transgenic (H) samples. Figure 6. Open in new tabDownload slide Profiles of concentration and [η] from size-exclusion chromatography for WE-AX from mature endosperm of transgenic and control wheat lines. Left panels show original profiles against retention volume, with void volume (Vo) and total volume (Vt) of the column indicated. Right panels show log [η], which is linearly related to log AX chain length, versus concentration. Data are shown for TaGT43_2 RNAi lines 3 and 5 and for TaGT47_2 RNAi lines 1 and 4 azygous control (A) and homozygous transgenic (H) samples. [η] and amount (percentage of white flour) of WE-AX from homozygous and azygous wheat lines determined by HPSEC Table I. [η] and amount (percentage of white flour) of WE-AX from homozygous and azygous wheat lines determined by HPSEC The values are averages of the profiles against retention volume shown in Figure 6. Results are presented for each of two extractions (1 and 2). Line . WE-AX . [η] . 1 . 2 . 1 . 2 . % of white flour mL g−1 GT43_2-3A 0.304 0.291 527 572 GT43_2-3H 0.103 0.104 283 294 GT43_2-5A 0.384 0.326 454 569 GT43_2-5H 0.111 0.137 332 297 GT47_2-1A 0.358 0.334 495 616 GT47_2-1H 0.132 0.165 310 432 GT47_2-4A 0.368 0.477 527 525 GT47_2-4H 0.164 0.098 438 499 Line . WE-AX . [η] . 1 . 2 . 1 . 2 . % of white flour mL g−1 GT43_2-3A 0.304 0.291 527 572 GT43_2-3H 0.103 0.104 283 294 GT43_2-5A 0.384 0.326 454 569 GT43_2-5H 0.111 0.137 332 297 GT47_2-1A 0.358 0.334 495 616 GT47_2-1H 0.132 0.165 310 432 GT47_2-4A 0.368 0.477 527 525 GT47_2-4H 0.164 0.098 438 499 Open in new tab Table I. [η] and amount (percentage of white flour) of WE-AX from homozygous and azygous wheat lines determined by HPSEC The values are averages of the profiles against retention volume shown in Figure 6. Results are presented for each of two extractions (1 and 2). Line . WE-AX . [η] . 1 . 2 . 1 . 2 . % of white flour mL g−1 GT43_2-3A 0.304 0.291 527 572 GT43_2-3H 0.103 0.104 283 294 GT43_2-5A 0.384 0.326 454 569 GT43_2-5H 0.111 0.137 332 297 GT47_2-1A 0.358 0.334 495 616 GT47_2-1H 0.132 0.165 310 432 GT47_2-4A 0.368 0.477 527 525 GT47_2-4H 0.164 0.098 438 499 Line . WE-AX . [η] . 1 . 2 . 1 . 2 . % of white flour mL g−1 GT43_2-3A 0.304 0.291 527 572 GT43_2-3H 0.103 0.104 283 294 GT43_2-5A 0.384 0.326 454 569 GT43_2-5H 0.111 0.137 332 297 GT47_2-1A 0.358 0.334 495 616 GT47_2-1H 0.132 0.165 310 432 GT47_2-4A 0.368 0.477 527 525 GT47_2-4H 0.164 0.098 438 499 Open in new tab Despite these massive changes in the principal polysaccharide of the endosperm cell wall, the transgenic seed appeared normal and grain weight and germination rates showed no consistent effects (Supplemental Fig. S3). Immunolabeling with the LM11 monoclonal antibody, which labels xylan that has little or no substitution with Ara (McCartney et al., 2005), showed little labeling of the starchy endosperm at 11 DPA, but by 18 DPA, there was substantial labeling in the cell walls of the controls but almost no labeling in large parts of the starchy endosperm in transgenic grain (Fig. 7). In both the TaGT43_2 and TaGT47_2 RNAi samples, labeling tended to be retained in the transfer cell region close to the dorsal crease and lost elsewhere. By 28 DPA, labeling with LM11 appeared to be abolished in the starchy endosperm in both the TaGT43_2 and TaGT47_2 RNAi samples, apart from a small area around the dorsal crease. Immunolabeling with the anti-AX1 monoclonal antibody raised against a mixture of AXOS (Guillon et al., 2004) gave more labeling at 11 DPA but smaller differences between transgenic and control samples (Fig. 8). However, the intensity of labeling was consistently lower in transgenics compared with controls, particularly in the lobe regions of the starchy endosperm; the label also showed apparently thinner cell walls at 28 DPA in transgenics compared with controls (Figs. 7 and 8). Measurements carried out on sections stained with toluidine blue (Supplemental Fig. S4) showed that the cell wall thickness was reduced by about 50% in the transgenic lines (Table II). Figure 7. Open in new tabDownload slide Sections of developing grain from TaGT43_2 RNAi line 6 and TaGT47_2 RNAi line 4 showing comparison of homozygous grain and corresponding azygous controls. Immunolabeling is with the LM11 monoclonal antibody raised against unsubstituted xylan oligosaccharide (McCartney et al., 2005). [See online article for color version of this figure.] Figure 7. Open in new tabDownload slide Sections of developing grain from TaGT43_2 RNAi line 6 and TaGT47_2 RNAi line 4 showing comparison of homozygous grain and corresponding azygous controls. Immunolabeling is with the LM11 monoclonal antibody raised against unsubstituted xylan oligosaccharide (McCartney et al., 2005). [See online article for color version of this figure.] Figure 8. Open in new tabDownload slide As for Figure 7, but immunolabeling was with the anti-AX1 monoclonal antibody raised against a mixture of AXOS (Guillon et al., 2004). [See online article for color version of this figure.] Figure 8. Open in new tabDownload slide As for Figure 7, but immunolabeling was with the anti-AX1 monoclonal antibody raised against a mixture of AXOS (Guillon et al., 2004). [See online article for color version of this figure.] Cell wall width (in µm) estimated by electronic calipers and a 100× objective on grain sections at 28 DPA stained with toluidine blue (see examples in Supplemental Fig. S4) Table II. Cell wall width (in µm) estimated by electronic calipers and a 100× objective on grain sections at 28 DPA stained with toluidine blue (see examples in Supplemental Fig. S4) Values are averages of multiple observations (n = number of observations) on single grain sections from replicate plants. Lines used were GT43_2 line 6 and GT47_2 line 4. Tissue . Construct . Replicate . Control A . Transgenic H . H/A . P . Mean ± sd n Mean ± sd n Central endosperm GT43_2 A 2.38 ± 1.22 38 1.02 ± 0.16 24 43% GT43_2 B 1.24 ± 0.39 13 GT47_2 A 2.39 ± 0.43 24 0.97 ± 0.17 25 41% GT47_2 B 2.08 ± 0.44 25 1.03 ± 0.17 25 49% Average 2.28 ± 0.45a 3 1.06 ± 0.11a 4 47% 0.0001 Aleurone periclinal GT43_2 A 3.30 ± 0.98 12 GT43_2 B 2.06 ± 0.28 12 GT47_2 A 3.45 ± 0.88 13 2.18 ± 0.60 13 63% GT47_2 B 4.09 ± 0.86 12 1.88 ± 0.49 14 46% Average 3.61 ± 0.06a 3 2.04 ± 0.16a 3 56% 0.0035 Aleurone anticlinal GT43_2 A 4.57 ± 0.33 8 GT43_2 B 2.29 ± 0.50 12 GT47_2 A 4.51 ± 0.74 12 3.99 ± 0.78 13 88% GT47_2 B 4.61 ± 0.86 12 3.10 ± 0.49 14 67% Average 4.56 ± 0.08a 2 3.49 ± 0.19a 4 76% 0.2268 Tissue . Construct . Replicate . Control A . Transgenic H . H/A . P . Mean ± sd n Mean ± sd n Central endosperm GT43_2 A 2.38 ± 1.22 38 1.02 ± 0.16 24 43% GT43_2 B 1.24 ± 0.39 13 GT47_2 A 2.39 ± 0.43 24 0.97 ± 0.17 25 41% GT47_2 B 2.08 ± 0.44 25 1.03 ± 0.17 25 49% Average 2.28 ± 0.45a 3 1.06 ± 0.11a 4 47% 0.0001 Aleurone periclinal GT43_2 A 3.30 ± 0.98 12 GT43_2 B 2.06 ± 0.28 12 GT47_2 A 3.45 ± 0.88 13 2.18 ± 0.60 13 63% GT47_2 B 4.09 ± 0.86 12 1.88 ± 0.49 14 46% Average 3.61 ± 0.06a 3 2.04 ± 0.16a 3 56% 0.0035 Aleurone anticlinal GT43_2 A 4.57 ± 0.33 8 GT43_2 B 2.29 ± 0.50 12 GT47_2 A 4.51 ± 0.74 12 3.99 ± 0.78 13 88% GT47_2 B 4.61 ± 0.86 12 3.10 ± 0.49 14 67% Average 4.56 ± 0.08a 2 3.49 ± 0.19a 4 76% 0.2268 a Mean and sd of means for each grain section shown above. Open in new tab Table II. Cell wall width (in µm) estimated by electronic calipers and a 100× objective on grain sections at 28 DPA stained with toluidine blue (see examples in Supplemental Fig. S4) Values are averages of multiple observations (n = number of observations) on single grain sections from replicate plants. Lines used were GT43_2 line 6 and GT47_2 line 4. Tissue . Construct . Replicate . Control A . Transgenic H . H/A . P . Mean ± sd n Mean ± sd n Central endosperm GT43_2 A 2.38 ± 1.22 38 1.02 ± 0.16 24 43% GT43_2 B 1.24 ± 0.39 13 GT47_2 A 2.39 ± 0.43 24 0.97 ± 0.17 25 41% GT47_2 B 2.08 ± 0.44 25 1.03 ± 0.17 25 49% Average 2.28 ± 0.45a 3 1.06 ± 0.11a 4 47% 0.0001 Aleurone periclinal GT43_2 A 3.30 ± 0.98 12 GT43_2 B 2.06 ± 0.28 12 GT47_2 A 3.45 ± 0.88 13 2.18 ± 0.60 13 63% GT47_2 B 4.09 ± 0.86 12 1.88 ± 0.49 14 46% Average 3.61 ± 0.06a 3 2.04 ± 0.16a 3 56% 0.0035 Aleurone anticlinal GT43_2 A 4.57 ± 0.33 8 GT43_2 B 2.29 ± 0.50 12 GT47_2 A 4.51 ± 0.74 12 3.99 ± 0.78 13 88% GT47_2 B 4.61 ± 0.86 12 3.10 ± 0.49 14 67% Average 4.56 ± 0.08a 2 3.49 ± 0.19a 4 76% 0.2268 Tissue . Construct . Replicate . Control A . Transgenic H . H/A . P . Mean ± sd n Mean ± sd n Central endosperm GT43_2 A 2.38 ± 1.22 38 1.02 ± 0.16 24 43% GT43_2 B 1.24 ± 0.39 13 GT47_2 A 2.39 ± 0.43 24 0.97 ± 0.17 25 41% GT47_2 B 2.08 ± 0.44 25 1.03 ± 0.17 25 49% Average 2.28 ± 0.45a 3 1.06 ± 0.11a 4 47% 0.0001 Aleurone periclinal GT43_2 A 3.30 ± 0.98 12 GT43_2 B 2.06 ± 0.28 12 GT47_2 A 3.45 ± 0.88 13 2.18 ± 0.60 13 63% GT47_2 B 4.09 ± 0.86 12 1.88 ± 0.49 14 46% Average 3.61 ± 0.06a 3 2.04 ± 0.16a 3 56% 0.0035 Aleurone anticlinal GT43_2 A 4.57 ± 0.33 8 GT43_2 B 2.29 ± 0.50 12 GT47_2 A 4.51 ± 0.74 12 3.99 ± 0.78 13 88% GT47_2 B 4.61 ± 0.86 12 3.10 ± 0.49 14 67% Average 4.56 ± 0.08a 2 3.49 ± 0.19a 4 76% 0.2268 a Mean and sd of means for each grain section shown above. Open in new tab DISCUSSION The suppression of TaGT43_2 and TaGT47_2 genes by RNAi resulted in the largest reported decrease of xylan in a grass, and in a cell type where this is the dominant polysaccharide. The effects can be compared with those in Arabidopsis xylan due to mutations in the respective homologs of TaGT43_2 and TaGT47_2, IRX9 and IRX10. Arabidopsis plants carrying these mutations also continue to produce xylan (due to the presence of functionally redundant genes), but in severely decreased amounts and with shorter average chain length (Brown et al., 2007, 2009; Pena et al., 2007; Wu et al., 2010; Lee et al., 2012). However, important differences were identified here: (1) Ara substitution was increased in the transgenic wheat lines, whereas GlcA substitution remains constant in the Arabidopsis mutants; (2) a clear difference in phenotype was identified in the loss of long AX chains in TaGT43_2 but not TaGT47_2 RNAi endosperm, whereas no clear difference between the xylan from irx9 and irx10 Arabidopsis mutants has been reported. These results are discussed in more detail below. Effect of TaGT43_2 and TaGT47_2 Suppression on Cell Wall Polysaccharides RNAi suppression of the TaGT43_2 and TaGT47_2 genes decreased the total amount by 50% in some lines, altering its structure such that the Ara-to-Xyl ratio was increased by approximately 25% (Fig. 4, A–C). There was some variation in effects between lines shown by both AXOS abundance and monosaccharide analysis (Fig. 5C); this variation was not explained by the variation between lines of measured effects on transcript abundance (Fig. 2), but this is only a snapshot of abundance for transcripts that vary widely with development. Allelic variation in the TaGT43_2 and TaGT47_2 genes, or in cis-elements controlling their expression, might be expected to have a large influence on AX and traits dependent on this, such as dietary fiber content and composition and flour extract viscosity. The genes are located on chromosomes 4A, 4B, 4D (TaGT43_2) and 3A, 3B, 3D (TaGT47_2). A consensus quantitative trait locus for AX and viscosity has been mapped to a distal part of the long arm of chromosome 3D (Quraishi et al., 2011). TaGT47_2 D is on the long arm of chromosome 3; greater resolution of its position is not yet possible due to the lack of a physical map for this chromosome, but the ortholog in barley (Hordeum vulgare) MLOC_64806 is localized toward the telomere of 3HL (identified using barley ENSEMBL at http://plants.ensembl.org based on the barley physical map; IBS Consortium, 2012), which suggests that this gene may colocalize with the quantitative trait locus in wheat. The effects of RNAi suppression on the amounts of oligosaccharides released by xylanase digestion were even more dramatic (Fig. 5A), indicating a decrease in the fraction of AX that was fully digestible to small oligosaccharides by xylanase, as confirmed by MALDI-MS (Fig. 5E). This is explained by the greater Ara substitution in the AX in transgenic lines: cleavage by the GH11 enzyme that was used requires three consecutive unsubstituted Xyl residues, so the more densely substituted AX chains present in the transgenics will have lower frequencies of such cleavage sites. There was a small increase in 1,3;1,4-β-d-glucan in the transgenic lines that had the greatest decreases in AX (Figs. 4F and 5D). This could be a compensatory effect in response to the depletion of AX in the cell wall; alternatively, it could be due to an accumulation of UDP-Glc in the Golgi lumen caused by the decreased use of substrates for xylan synthesis. Spatial and Temporal Variations of Effects Although the HMW1Dx5 promoter drives strong expression at 10 DPA under our standard conditions (Pellny et al., 2012), we have previously observed that the effects of RNAi transgenes driven by this promoter on cell walls do not become evident until 14 DPA (Nemeth et al., 2010); this delay is expected due to the time required for the RNAi machinery to operate and for existing enzymes to turn over. Here, we observed complete abolition of LM11 labeling in most of the starchy endosperm by 18 DPA in the TaGT47_2 line and a similar effect in the most distal layers of the dorsal region and of the lobes in the TaGT43_2 line (Fig. 7). This absence of LM11 labeling is likely due both to the decreased amount of AX and to increased Ara substitution in AX of transgenic lines, since this monoclonal antibody was raised using an unsubstituted penta-1,4-xylanoside glycoprotein (McCartney et al., 2005). This explanation is supported by the labeling pattern of the anti-AX1 monoclonal antibody (which recognizes substituted AXOS; Guillon et al., 2004), as this was never abolished in transgenics, although a decrease in labeling intensity was observable (Fig. 8). LM11 labeling was never abolished in transfer cells and immediately surrounding region cells, which show similar labeling to controls (Fig. 7); this region contains some of the earliest developing cells within the starchy endosperm, and the HMW1Dx5 promoter does not drive expression strongly in transfer cells themselves. In general, the cellular organization and structure of the starchy endosperm of the transgenic lines was remarkably unaffected by the loss of up to 50% of the major cell wall polysaccharide, even though the cell walls were about 50% thinner by 28 DPA (Table II). Although the HMW1Dx5 promoter is specific to the starchy endosperm cells, the aleurone periclinal cell walls were also approximately 40% thinner (Table II); this could be due to the movement of small interfering RNA molecules from the starchy endosperm to the aleurone. Participation of TaGT43_2 and TaGT47_2 in Xylan Synthetic Machinery The effects of suppressing TaGT43_2 and TaGT47_2 were similar for most parameters measured (Figs. 4, 5, 7, and 8; Table I). This is in accordance with findings in Arabidopsis, where the effects on xylan of mutations in IRX9, IRX10, and IRX14 (sometimes combined with mutations in close homologs to address functional redundancy) are essentially the same: substantial decreases in xylan content and in xylan chain length. Here, we studied the effects of suppressing homologs of only two of these genes: IRX9 and IRX10. However, the third component, the IRX14 homolog, which we call TaGT43_1, is expressed in wheat endosperm at similar levels to IRX9 (Pellny et al., 2012). The TaGT43_1 protein and another IRX10 homolog were also found in a microsomal complex from wheat seedlings with xylan synthase activity (Zeng et al., 2010). From studies in Arabidopsis, it appears that there is a requirement for all three components to maintain xylan synthase activity (Brown et al., 2007, 2009; Pena et al., 2007; Wu et al., 2010; Lee et al., 2012). Since homologous poplar genes are able to complement for the lack of functional IRX9 or IRX14 (Lee et al., 2011), their function seems to have been conserved in evolution. All the plants that have been fully sequenced have genes for the three components, with the lycophyte S. moellendorffii having the minimal set of three genes (Fig. 1). Our results are consistent with such a xylan synthase complex existing in the starchy endosperm cells of wheat, with components encoded by TaGT43_2 and TaGT47_2 being responsible for the majority of the AX backbone synthesis in this tissue. It seems likely that a third component corresponding to IRX14 would also be essential for normal AX synthesis and that TaGT43_1 probably encodes this component, but this was not tested here. AX Arabinosylation The degree of AX substitution was increased as indicated by increases in the AX Ara-to-Xyl ratio of up to 40% (Fig. 4B). This contrasts strongly with irx9 and irx10/irx10-L knockout mutants in Arabidopsis, which maintained the same degree of substitution by methylated or unmethylated GlcA as in the wild-type plants, about 11% of Xyl residues (Brown et al., 2007, 2009), when xylan synthesis was strongly suppressed. It appears that the excess arabinosylation capacity when AX backbone synthesis was suppressed led to regions of highly substituted AX that were not fully digestible by xylanase, leading to even greater decreases in abundance of small AXOS (Fig. 5A). The observation that the unsubstituted X and XX released by xylanase was decreased to about the same degree as total AX, but substituted AXOS was decreased much more (Fig. 5C), may suggest the existence of fractions of AX with low substitution that were not much altered in the transgenics, whereas other fractions with higher Ara substitution tended to become even more highly substituted and, therefore, gave rise to a greater proportion of higher M r AXOS upon digestion with GH11 (Fig. 5E). WE-AX Chain Length Profiles from size-exclusion chromatography suggest that the average chain length of xylan was substantially decreased in the Arabidopsis irx9, irx10, and irx14 mutants (Pena et al., 2007; Brown et al., 2009; Wu et al., 2010). Whereas a rice mutant with inactive OsIRX10 showed no change in the M r profile of a fraction solubilized by 1 m KOH (Chen et al., 2013), only one of the six IRX10-L rice genes was inactivated, resulting in a 10% decrease in cell wall Xyl, compared with a 60% decrease in the Arabidopsis IRX10/IRX10-L double mutant. Effects on chain length cannot be inferred from profiles of concentration against retention volume alone when substitution is changed, as is the case here, because this will also affect M r. However, it has been shown that the chain length of wheat grain WE-AX is directly related to [η] for a wide range of Ara-to-Xyl ratios (Dervilly-Pinel et al., 2001). When this relationship was applied, it was clear that both GT43_2 and GT47_2 RNAi transgenic lines did indeed show a substantial decrease in average chain length of WE-AX (Table I; Fig. 6). However, a shoulder in the profiles of [η] for the GT43_2 RNAi transgenic lines, which corresponds to a peak in the control, shows a decrease in the chain length of WE-AX by a factor of more than 2 (Fig. 6, A and B), whereas there is apparently no effect in the GT47_2 RNAi lines (Fig. 6, C and D); similar differences are also seen in the maximum chain length. This suggests that xylan synthase complexes partially deficient in TaGT43_2 lack the capacity to make long AX chains. By contrast, xylan synthase complexes partially deficient in TaGT47_2 can make such long chains, albeit at a much decreased rate. To our knowledge, this is the first finding of a clear difference between the roles of IRX9 and IRX10 homologs, and it is possible that they are only apparent in wheat endosperm WE-AX because of the much greater average chain length of 1,000 to 1,500 Xyl (Saulnier et al., 2007) compared with Arabidopsis stem GX at around 100 Xyl (Pena et al., 2007). CONCLUSION RNAi suppression of TaGT43_2 and TaGT47_2 in wheat induces massive changes in the starchy endosperm AX composition. Therefore, variation in the expression of these genes, or the activity of their encoded proteins, will affect important traits such as dietary fiber content and composition and the viscosity of extracts from wheat grain. The effect on AX is consistent with the TaGT43_2 and TaGT47_2 proteins playing key roles in the synthesis of the xylan backbone, but differences in the effects on WE-AX chain length give intriguing clues to the differing roles of these two proteins within the xylan synthesis machinery. MATERIALS AND METHODS Plant Growth Bread wheat (Triticum aestivum ‘Cadenza’) plants were grown in temperature-controlled glasshouse rooms as described (Nemeth et al., 2010). RNAi Construct Preparation and Transformation Full-length TaGT43_2 and TaGT47_2 clones were obtained from endosperm complementary DNA as described previously (Anders et al., 2012) but using primers GT43_2F, GT43_2R, GT47_2F, and GT47_2R (Supplemental Table S2). Multiple clones were sequenced as described previously to obtain the consensus sequences shown in Supplemental Figures S1 and S2, which correspond exactly to regions of chromosome-sorted contigs available at www.wheatgenome.org, confirming that the three isoforms originate from homologous chromosomes of the A, B, and D genomes. These sequences have been deposited in the EMBL European Nucleotide Sequence database with the following accession numbers: TaGT43_2D, HF913567; TaGT43_2B, HF913568; TaGT43_2A, HF913569; TaGT47_2B, HF913570; TaGT47_2D, HF913571; TaGT47_2A, HF913572. RNAi constructs with the starchy endosperm-specific HMW1Dx5 promoter were created as described by Nemeth et al. (2010) but using the 446- and 563-bp fragments indicated in Supplemental Figures S1 and S2, respectively, using the PCR primers indicated in Supplemental Table S2. Wheat transformation was carried out by particle bombardment (PDS1000; Bio-Rad) of immature scutella (10–14 DPA) of cv Cadenza according to Sparks and Jones (2009), and zygosity testing was carried out as described previously (Nemeth et al., 2010). For each construct, transgenic lines with segregation consistent with a single insertion locus were identified. In subsequent generations, homozygous and azygous segregants descended from the same original transformant were identified and grown as three replicate pots in block design experiments. Analyses were conducted on T3 or T4 seed from such experiments. Transcript Analysis Total RNA was extracted as described (Nemeth et al., 2010). Down-regulation of the transcript was measured as described previously (Anders et al., 2012). In short, developing T4 seeds (T3 for GT47_2 line 6) were harvested at 21 DPA, and RNA from pure endosperm was extracted. In the GT43_2 RNAi lines, the transcript down-regulation was tested using primers prTYW343 and prTYW348, resulting in a 117-bp amplicon. For the GT47_2 transgenics, in addition to the targeted gene (primers prTYW460 and prTYW462; 94-bp fragment), the transcript levels for GT47_1 and GT47_4 were determined, as they also showed some sequence homology to the RNAi fragment used. The primers were prTYW273 and prTYW449 (61 bp) for GT47_1 and prTYW464 and prTYW465 (96 bp) for GT47_4. Three reference genes were used to normalize expression: Ta2526, a stably expressed EST from grain (primers prTYW19 and TYW20), glyceraldehyde-3-phosphate dehydrogenase (primers prTYW270 and prTYW271), and succinate dehydrogenase (primers prTYW209 and prTYW210). All primer sequences are given in Supplemental Table S2. Cell Wall Analyses White flour fractions (pure starchy endosperm) were obtained from mature seed as described previously (Anders et al., 2012). Analyses of nonstarch monosaccharide content of white flour samples were conducted following the protocol of Englyst et al. (1994). The methodology for analysis of endosperm AX and 1,3;1,4-β-d-glucan by digestion with endoxylanase and lichenase followed by HPAEC of resultant AXOS was originally developed by Ordaz-Ortiz et al. (2005), and the procedure followed here was as described by Nemeth et al. (2010). The monosaccharide contents of the products of endoxylanase and lichenase digestion (Nemeth et al., 2010; xylanase extractable) were determined as described (Englyst et al., 1994), except that total hydrolysate volume was 7.85 mL. MALDI-MS analyses of AXOS released by digestion were performed on an Autoflex III MALDI-time of flight/time of flight mass spectrometer (Bruker Daltonics) equipped with a Smartbeam laser (355 nm) in positive ion mode with linear detection. Acquisition parameters (laser power, pulsed ion extraction, etc.) were optimized for each sample. Mass spectra were automatically processed with Flex Analysis 3.0 software (Bruker Daltonics). A mixture of 2,5-dihydroxybenzoic acid and N-dimethylaniline was used as ionic matrix, and samples were prepared as described previously (Ropartz et al., 2011). High-Performance Size-Exclusion Chromatography For extraction of WE-AX, 500 mg of flour was suspended in 2 mL of water and agitated for 20 min at room temperature; after centrifugation, supernatants were heated for 5 min in a boiling-water bath, filtrated over 0.45 µm, and stabilized to pH 2 with Gly-HCl buffer. Extracts (0.05 mL) were injected on the high-performance size-exclusion chromatography (HPSEC) system. HPSEC was performed at room temperature on a system consisting of a Shodex OH SB-G guard column (Showa Denko) and Shodex OH-Pak SB-805 HQ a columns eluted at 1 mL min−1 with 50 mm sodium nitrate buffer. The Viscotek tri-SEC model 270 was used for light scattering and differential pressure detection, and a Viscotek VE 3580 RI detector was used for the determination of polymer concentration. A refractive index increment per unit concentration increment (dn/dc) value of 0.146 mL g−1 was used for concentration determination. Data were collected with Omnisec 4.5 software (Viscotek), and all calculations on polymer peaks (concentration, [η]) were carried out using Omnisec software. Due to the presence of aggregates in some of the samples, average M r values from light-scattering measurements were overestimated and, therefore, not used. Conversely, the viscosity detector is not affected by the presence of aggregates; therefore, [η] values were used for comparison of samples. Microscopy Transverse slices, approximately 1 mm thick, were cut from developing wheat grains at 11, 18, and 28 DPA and fixed and embedded as described (Pellny et al., 2012). The LM11 antibody was used at a dilution of 1:5; the monoclonal anti-AX1 antibody was used at a 1:25 dilution; secondary antibodies (anti-rat Alexa 633 conjugated and anti-mouse Alexa 568 conjugated [Invitrogen]) were used at a 1:100 dilution. Images were taken using a Zeiss 780 confocal microscope. Cell wall width measurements were done on 1-µm-thick transverse sections stained with toluidine blue prepared from 28-DPA samples. Imaging was done on a Zeiss Axiophot microscope equipped with a QImaging Retiga Exi CCD digital camera using a 100× objective. Measurements of width were done in MetaMorph software (Molecular Devices) using digital calipers. Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers HF913567, HF913568, HF913569, HF913570, HF913571, and HF913572. Supplemental Data The following materials are available in the online version of this article. Supplemental Figure S1. Alignment of TaGT43_2A, B and D cDNA sequences. Supplemental Figure S2. Alignment of TaGT47_2A, B and D cDNA sequences. Supplemental Figure S3. Grain dry weight and germination rates. Supplemental Figure S4. Example grain sections stained with Toluidine Blue. Supplemental Table S1. Monosaccharide analyses of endosperm cell walls. Supplemental Table S2. List of PCR primers. ACKNOWLEDGMENTS We thank Dr. Stephen Powers (Rothamsted Research) for statistical analyses. We thank D. Ropartz from the Biopolymères, Biologie Structurale platform (INRA Angers-Nantes Center) for mass spectrometry analyses. 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