w

# **ENDOPLASMIC RETICULUM – SHAPE AND FUNCTION IN STRESS TRANSLATION**

**Topic Editors Lorenzo Frigerio, Federica Brandizzi, Stephen H. Howell and Patrick Schäfer**

#### *FRONTIERS COPYRIGHT STATEMENT*

© Copyright 2007-2015 Frontiers Media SA. All rights reserved.

All content included on this site, such as text, graphics, logos, button icons, images, video/audio clips, downloads, data compilations and software, is the property of or is licensed to Frontiers Media SA ("Frontiers") or its licensees and/or subcontractors. The copyright in the text of individual articles is the property of their respective authors, subject to a license granted to Frontiers.

The compilation of articles constituting this e-book, wherever published, as well as the compilation of all other content on this site, is the exclusive property of Frontiers. For the conditions for downloading and copying of e-books from Frontiers' website, please see the Terms for Website Use. If purchasing Frontiers e-books from other websites or sources, the conditions of the website concerned apply.

Images and graphics not forming part of user-contributed materials may not be downloaded or copied without permission.

Individual articles may be downloaded and reproduced in accordance with the principles of the CC-BY licence subject to any copyright or other notices. They may not be re-sold as an e-book.

As author or other contributor you grant a CC-BY licence to others to reproduce your articles, including any graphics and third-party materials supplied by you, in accordance with the Conditions for Website Use and subject to any copyright notices which you include in connection with your articles and materials.

All copyright, and all rights therein, are protected by national and international copyright laws.

The above represents a summary only. For the full conditions see the Conditions for Authors and the Conditions for Website Use.

**ISSN** 1664-8714 **ISBN** 978-2-88919-344-8 **DOI** 10.3389/978-2-88919-344-8

## *ABOUT FRONTIERS*

Frontiers is more than just an open-access publisher of scholarly articles: it is a pioneering approach to the world of academia, radically improving the way scholarly research is managed. The grand vision of Frontiers is a world where all people have an equal opportunity to seek, share and generate knowledge. Frontiers provides immediate and permanent online open access to all its publications, but this alone is not enough to realize our grand goals.

## *FRONTIERS JOURNAL SERIES*

The Frontiers Journal Series is a multi-tier and interdisciplinary set of open-access, online journals, promising a paradigm shift from the current review, selection and dissemination processes in academic publishing.

All Frontiers journals are driven by researchers for researchers; therefore, they constitute a service to the scholarly community. At the same time, the Frontiers Journal Series operates on a revolutionary invention, the tiered publishing system, initially addressing specific communities of scholars, and gradually climbing up to broader public understanding, thus serving the interests of the lay society, too.

## *DEDICATION TO QUALITY*

Each Frontiers article is a landmark of the highest quality, thanks to genuinely collaborative interactions between authors and review editors, who include some of the world's best academicians. Research must be certified by peers before entering a stream of knowledge that may eventually reach the public - and shape society; therefore, Frontiers only applies the most rigorous and unbiased reviews.

Frontiers revolutionizes research publishing by freely delivering the most outstanding research, evaluated with no bias from both the academic and social point of view.

By applying the most advanced information technologies, Frontiers is catapulting scholarly publishing into a new generation.

## *WHAT ARE FRONTIERS RESEARCH TOPICS?*

Frontiers Research Topics are very popular trademarks of the Frontiers Journals Series: they are collections of at least ten articles, all centered on a particular subject. With their unique mix of varied contributions from Original Research to Review Articles, Frontiers Research Topics unify the most influential researchers, the latest key findings and historical advances in a hot research area!

Find out more on how to host your own Frontiers Research Topic or contribute to one as an author by contacting the Frontiers Editorial Office<EMAIL_ADDRESS>
## **ENDOPLASMIC RETICULUM - SHAPE AND FUNCTION IN STRESS TRANSLATION**

Topic Editors:

**Lorenzo Frigerio,** University of Warwick, United Kingdom **Federica Brandizzi,** Michigan State University, East Lansing, USA **Stephen H. Howell,** Iowa State University, USA **Patrick Schäfer,** University of Warwick, United Kingdom

Schematic representation of topics covered in the Research Topic. Numbers correspond to articles as follows: (1) Griffing et al. (2014), ER network dynamics are differentially controlled by myosins XI-K, XI-C, XI-E, XI-I, XI-1, and XI-2, (2) Viotti (2014), ER and vacuoles: never been closer, (3) Li and Liu (2014), Endoplasmic reticulum-mediated protein quality control in Arabidopsis, (4) Ruberti and Brandizzi (2014), Conserved and plant-unique strategies for overcoming endoplasmic reticulum stress, (5) Srivastava et al. (2014), Stress sensing in plants by an ER stress sensor/transducer, bZIP28, (6) Tintor and Saijo (2014), ER-mediated control for abundance, quality, and signaling of transmembrane immune receptors in plants, (7) Verchot (2014), The ER quality control and ER associated degradation machineries are vital for viral pathogenesis, (8) Nakano et al. (2014), ER bodies in plants of the Brassicales order: biogenesis and association with innate immunity, (9) Höwing et al. (2014), Endoplasmic reticulum KDEL-tailed cysteine endopeptidase 1 of Arabidopsis (AtCEP1) is involved in pathogen defense, (10) Williams et al. (2014), When supply does not meet demand-ER stress and plant programmed cell death, (11) Cai et al. (2014), Endoplasmic reticulum stress-induced PCD and caspaselike activities involved.

The endoplasmic reticulum (ER) is a manufacturing unit in eukaryotic cells required for the synthesis of proteins, lipids, metabolites and hormones. Besides supporting cellular signalling networks by its anabolic function, the ER on its own or in communication with other organelles directly initiates signalling processes of physiological significance. Based on the intimate and immediate involvement in stress signalling the ER is considered as sensory organelle on which cells strongly rely to effectively translate environmental cues into adaptive stress responses. The transcellular distribution of the ER providing comprehensive cell-to-cell connections in multicellular organisms probably allows a concerted action of cell alliances and tissue areas towards environmental constraints. At the cellular level, stress adaptation correlates with the capability of the ER machinery to synthesise proteins participating in stress signalling as well as in the activation of ER membrane localised proteins to start cellprotective signalling processes. Importantly, depending on the stress insult, the ER either supports protective strategies or initiates cell death programmes. Recent, genetic, molecular and cell biological studies have drawn an initial picture of underlying signalling events activated by ER membrane localised proteins. In this Research Topic, we provided a platform for articles describing research on ER morphology and metabolism with a focus on stress translation. The Research Topic is sub-divided into the following sections:

# Table of Contents

Lawrence R. Griffing, Hongbo T. Gao and Imogen Sparkes

Cristina Ruberti and Federica Brandizzi

Nico Tintor and Yusuke Saijo

*57 The ER Quality Control and ER Associated Degradation Machineries are Vital for Viral Pathogenesis* Jeanmarie Verchot

*68 ER Bodies in Plants of the Brassicales Order: Biogenesis and Association with Innate Immunity*

Ryohei T.Nakano, Kenji Yamada, Paweł Bednarek, Mikio Nishimura and Ikuko Hara-Nishimura

*85 Endoplasmic Reticulum KDEL-Tailed Cysteine Endopeptidase 1 of Arabidopsis (AtCEP1) is Involved in Pathogen Defense*

Timo Höwing, Christina Huesmann, Caroline Hoefle, Marie-Kristin Nagel, Erika Isono, Ralph Hückelhoven and Christine Gietl

*96 When Supply does not Meet Demand-ER Stress and Plant Programmed Cell Death*

Brett Williams, Jeanmarie Verchot and Martin B. Dickman

*105 Endoplasmic Reticulum Stress-Induced PCD and Caspase-Like Activities Involved*

Yao-min Cai, Jia Yu and Patrick Gallois

## Endoplasmic reticulum—shape and function in stress translation

#### *Federica Brandizzi 1 \*, Lorenzo Frigerio2 \*, Stephen H. Howell 3 \* and Patrick Schäfer 2 \**

*1 Plant Research Laboratory, Department of Energy, Michigan State University, East Lansing, MI, USA*

*2 School of Life Sciences, University of Warwick, Coventry, UK*

*3 Department of Genetics, Development and Cell Biology, Plant Sciences Institute, Iowa State University, Ames, IA, USA*

*\*Correspondence<EMAIL_ADDRESS><EMAIL_ADDRESS><EMAIL_ADDRESS><EMAIL_ADDRESS>
#### *Edited and reviewed by:*

*Simon Gilroy, University of Wisconsin - Madison, USA*

**Keywords: unfolded protein response (UPR), ER associated degradation (ERAD), ER stress, bZIP transcription factors, myosins, caspases, ER bodies, cysteine endopeptidase, programmed cell death**

The endoplasmic reticulum (ER) is a very versatile organelle. Besides its major role as the gateway to the secretory pathway the ER is central to adaptation against abiotic and biotic stress. Here, we summarize the current knowledge on ER dynamics and architecture, the association and interaction of the ER with other organelles as well as its role in stress translation and adaptation (**Figure 1**).

The ER is a dynamic network of membrane tubules and sheets. Movement of the ER along the actin cytoskeleton has a significant influence on both its architecture and dynamics. Members of class XI myosin motor proteins have recently been shown to control the movement of the ER and other organelles such as Golgi bodies, peroxisomes and mitochondria. Griffing et al. (2014) explore a subset of class XI myosins with a focus on their role in regulating ER network remodeling by affecting ER tubulation, sheet formation and the persistence of these structures. In addition to the importance of these ER network dynamics for organelle communication, the ER may further directly provide the source membrane for vacuole biogenesis, as hypothesized by Viotti (2014). The majority of soluble vacuolar proteins travel from the ER to the Golgi complex. A growing number of reports however indicate that some proteins, including some vacuolar membrane proteins appear to reach the vacuole without visiting intermediate compartments of the endomembrane system.

An essential function of the ER is the synthesis of secreted proteins. To insure that only correctly folded proteins are exported from the ER, eukaryotes have evolved ER quality control (ERQC) mechanisms, which supervise the folding process. Protein folding is monitored through the stepwise modification of oligosaccharide side chains on glycoproteins, and proteins that fail to fold correctly are extracted from the folding process and subjected to ER associated degradation (ERAD). Li and Liu (2014) describe the protein folding process and how client proteins are chosen for ERAD. Proper function of the ER is particular relevant under stress where the demand for secreted proteins exceed the ER working load capacity. Under such conditions, ER homeostasis requires cellular communication between the ER and the nucleus. Therefore, eukaryotes have signaling proteins on the ER membrane that sense impaired ER function (ER stress) through the accumulation of misfolded proteins in the ER and that employ different strategies to signal the nucleus. Some of these communication strategies are common to all eukaryotes, while others are unique to plants. Ruberti and Brandizzi (2014) compare the ER stress signal pathways between yeast, plants and animals and review the responses in plants, which vary from adaptive measures to cell death. The bZIP transcription factor bZIP28 represents an ER stress signaling factor that is tethered

**FIGURE 1 | Schematic representation of topics covered in the special issue.** Numbers correspond to review articles as follows: (1) Griffing et al. (2014), ER network dynamics are differentially controlled by myosins XI-K, XI-C, XI-E, XI-I, XI-1, and XI-2, (2) Viotti (2014), ER and vacuoles: never been closer, (3) Li and Liu (2014), Endoplasmic reticulum-mediated protein quality control in Arabidopsis, (4) Ruberti and Brandizzi (2014), Conserved and plant-unique strategies for overcoming endoplasmic reticulum stress, (5) Srivastava et al. (2014), Stress sensing in plants by an ER stress sensor/transducer, bZIP28, (6) Tintor and Saijo (2014), ER-mediated control for abundance, quality, and signaling of transmembrane immune receptors in plants, (7) Verchot (2014), The ER quality control and ER associated degradation machineries are vital for viral pathogenesis, (8) Nakano et al. (2014), ER bodies in plants of the Brassicales order: biogenesis and association with innate immunity, (9) Höwing et al. (2014), Endoplasmic reticulum KDEL-tailed cysteine endopeptidase 1 of Arabidopsis (AtCEP1) is involved in pathogen defense, (10) Williams et al. (2014), When supply does not meet demand-ER stress and plant programmed cell death, (11) Cai et al. (2014), Endoplasmic reticulum stress-induced PCD and caspase-like activities involved.

to the ER membrane under unstressed conditions but is mobilized and transferred to the nucleus upon stress. Srivastava et al. (2014) describe the structure of bZIP28 and underlying principles of bZIP28 mobilization in response to stress.

ER stress activates the unfolded proteins response (UPR) and the ERAD system to eliminate misfolded proteins, which is vital for the establishment of an effective immune system. Tintor and Saijo (2014) introduce the significance of the ERQC in the synthesis of immune receptors and provide further insights into the causalities of ER function (UPR) and plant immunity. Despite its role in immunity, the ER has been hijacked by viruses to promote viral pathogenesis. As reviewed by Verchot (2014), viruses have developed sophisticated strategies to overload the ER protein folding machinery with viral encoded proteins in infected cells. Interestingly, this activates the UPR and up-regulates cellular chaperones that further aid in virus protein folding. The overall significance of the ER in stress adaption is however apparent and underlined by the review of Nakano et al. (2014) which, in addition to the biogenesis and evolution, discusses the putative function of ER bodies in abiotic and biotic stress. As ER-derived compartments ER bodies contain stress-associated proteins (e.g., various β-glucosidases of the PYK10 family) that are thought to enhance stress resilience by activating glucosinolates and glucosylated phytohormones. The study of Höwing et al. (2014) further identified an immune-active function of the ER-localized cysteine endopeptidase AtCEP1. In addition to its function in developmental processes, AtCEP1 is expressed and localized to the ER network, which condenses around haustoria during powdery mildew infection. Knockout of AtCEP1 results in enhanced susceptibility, suggesting that AtCEP1 is involved in restricting fungal infection, possibly by controlling a defenserelated programmed cell death (PCD) at the late stages of the interaction.

Cells mount adaptive responses (e.g., UPR) to mitigate the damage caused by ER stress. However, when stress becomes so excessive that the ERQC system cannot meet the demands, then PCD ensues. Williams et al. (2014) review the possible links between the UPR and PCD and discuss the involvement of calcium signaling and N-rich proteins in promoting PCD and the role of ER chaperones in limiting it as well as the connection of pathogens to ER-mediated cell death. Cai et al. (2014) finally compare the activation and/or regulation of ER stress-induced PCD in animals and plants and highlight the significance of caspase and caspase-like activities in underlying PCD processes.

The articles published in this e-book summarize our current knowledge of the multi-functionality of the ER in stress adaptation. By providing a stable microenvironment for the synthesis of metabolites and secreted proteins, the ER functions as an intracellular stress sensory organelle and, accordingly, initiates and regulates adaptive responses to environmental stress. Understanding the molecular basis of these processes and the role of ER architectural dynamics therein is of high relevance for sustainable crop production. We hope our e-book will stimulate research in the field to further enhance our knowledge of ER biology in plants.

## **ACKNOWLEDGMENTS**

Research in the lab of Federica Brandizzi is supported by the NIH, NASA, NSF, and DOE. Research in the lab of Lorenzo Frigerio is supported by the BBSRC and the Leverhulme Trust. Research in the lab of Patrick Schäfer is supported by the Deutsche Forschungsgemeinschaft (DFG), BASF and the University of Warwick.

### **REFERENCES**

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 06 August 2014; accepted: 11 August 2014; published online: 01 September 2014.*

*Citation: Brandizzi F, Frigerio L, Howell SH and Schäfer P (2014) Endoplasmic reticulum—shape and function in stress translation. Front. Plant Sci. 5:425. doi: 10.3389/fpls.2014.00425*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Brandizzi, Frigerio, Howell and Schäfer. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## ER network dynamics are differentially controlled by myosins XI-K, XI-C, XI-E, XI-I, XI-1, and XI-2

#### *Lawrence R. Griffing1, Hongbo T. Gao2 and Imogen Sparkes 2 \**

*1 Biology Department, Texas A&M University, College Station, TX, USA*

*2 Biosciences, College of Life and Environmental Sciences, Exeter University, Exeter, UK*

#### *Edited by:*

*Christopher J. Staiger, Purdue University, USA*

#### *Reviewed by:*

*Andreas Nebenführ, University of Tennessee, USA Valerian V. Dolja, Oregon State University, USA*

#### *\*Correspondence:*

*Imogen Sparkes, Biosciences, College of Life and Environmental Sciences, Exeter University, Geoffrey Pope Building, Stocker Rd., Exeter, EX4 4QD, UK e-mail<EMAIL_ADDRESS>
The endoplasmic reticulum (ER) of higher plants is a complex network of tubules and cisternae. Some of the tubules and cisternae are relatively persistent, while others are dynamically moving and remodeling through growth and shrinkage, cycles of tubule elongation and retraction, and cisternal expansion and diminution. Previous work showed that transient expression in tobacco leaves of the motor-less, truncated tail of myosin XI-K increases the relative area of both persistent cisternae and tubules in the ER. Likewise, transient expression of XI-K tail diminishes the movement of organelles such as Golgi and peroxisomes. To examine whether other class XI myosins are involved in the remodeling and movement of the ER, other myosin XIs implicated in organelle movement, XI-1 (MYA1),XI-2 (MYA2), XI-C, XI-E, XI-I, and one not, XI-A, were expressed as motor-less tail constructs and their effect on ER persistent structures determined. Here, we indicate a differential effect on ER dynamics whereby certain class XI myosins may have more influence over controlling cisternalization rather than tubulation.

**Keywords: endoplasmic reticulum, movement, myosin, persistency mapping, dynamics, remodeling**

## **INTRODUCTION**

The endoplasmic reticulum (ER) is a polygonal network that rapidly interconverts between tubular and cisternal forms, with tubules also growing, shrinking, and undergoing lateral sliding to form junctions, and cisternae undergoing expansion and shrinkage (Griffing, 2010; Sparkes et al., 2011). Whilst ER network dynamics are mainly controlled by actin and myosin (Quader et al., 1987; Liebe and Menzel, 1995; Sparkes et al., 2009a; Ueda et al., 2010), ER morphology and structure are also governed by additional factors such as reticulons and RHD3 implicated in ER membrane curvature and homotypic fusion (Sparkes et al., 2011; Zhang and Hu, 2013). Quantifying network dynamics is challenging because there are at least two levels at which movement takes place; (1) flow of protein within the lumen and on the surface of ER tubules and cisternae, as well as (2) remodeling through translocation and transformation of the tubular and cisternal elements themselves. Note, membrane flow can occur even when there is no remodeling (Sparkes et al., 2009a). The former movement, flow within and on the membrane is measured with fluorescent recovery after photobleaching (FRAP) and photoactivation (Sparkes et al., 2009a) as well as with optical flow image processing (Ueda et al., 2010; Stefano et al., 2014). But it is the latter form of movement, ER remodeling, which is analyzed here, starting from the perspective that there are relatively persistent features of the ER upon which change agents act. Hence, an approach to visualize and measure the relatively static or persistent regions in the network has been developed, persistency mapping (Sparkes et al., 2009a). Persistency mapping is an image processing technique that includes morphometric processing to separate tubules from cisternae and visualizes the relative movement and change of these ER elements as the ER remodels. The less-moving, more persistent fraction of the ER tubules and cisternae is then quantified. It is tempting to assign functions of some of the persistent structures of the ER, such as the potential anchor sites (Sparkes et al., 2009a,b) which may be regions where traffic between the plasma membrane and the ER can take place or/and could act as anchors to tether the ER required for spread throughout the cortex. A role in tethering or anchoring the ER to the plasma membrane was alluded to by optical tweezer experiments whereby a trapped Golgi body was used to stably remodel the ER around small islands of ER (Sparkes et al., 2009b). These experiments also highlighted that there is a physical connection between Golgi bodies and the ER bringing into question whether Golgi and ER dynamics are coordinated or are independent of one another? The common network upon which these interactions play out is the actin cytoskeleton.

Organelle movement in interphase plant cells is largely driven by actin and myosin, however the specificity of myosin-organelle interactions is poorly understood (Madison and Nebenführ, 2013 and references therein). In plants, organelle positioning and movement are correlated with responses to extracellular stresses such as pathogen ingress, wounding, and cadmium toxicity (Takemoto et al., 2003; Lipka et al., 2005; Hardham et al., 2008; Rodriguez-Serrano et al., 2009). At present, the precise role that organelle dynamics play in such processes is unclear. The best characterized study of the functional role of organelle movement is that of chloroplast repositioning upon high light, a response required to prevent photodamage (Wada et al., 2003). In addition there is a correlation between the morphology of the ER and the level of secretion with cisternal ER being more prevalent in cells producing more protein (Stephenson and Hawes, 1986; Ridge et al., 1999). Treating cells with latrunculin B to depolymerize actin results in a cessation of movement of spheroid organelles such as Golgi (Boevink et al., 1998; Nebenführ et al., 1999), mitochondria (Van Gestel et al., 2002; Zheng et al., 2009), and peroxisomes (Jedd and Chua, 2002; Mano et al., 2002; Mathur et al., 2002), and produces a relatively static ER network (Quader et al., 1987; Liebe and Menzel, 1995; Sparkes et al., 2009a) with larger cisternae, indicating that an actively remodeling system affects the global morphology of the ER. The movement of Golgi, peroxisomes and mitochondria appear to be controlled by a subset of class XI myosins; XI-C, XI-E, XI-I, XI-K, XI-1, and XI-2 (Avisar et al., 2008, 2009, 2012; Peremyslov et al., 2008, 2010; Prokhnevsky et al., 2008; Sparkes et al., 2008, 2009a). Dominant negative tail domain expression of these different myosins shows some quantitative differences in the reduction of movement produced by each member of this subset, but the results do not preclude the simple interpretation that one myosin form may control the movement of several different organelles. Double, triple and quadruple *Arabidopsis* mutants in myosins XI-K, XI-1, XI-2, and XI-I have reduced organelle dynamics and display gross morphological defects (Prokhnevsky et al., 2008; Peremyslov et al., 2010; Ojangu et al., 2012). Although myosin XI-K has been shown to change ER form and dynamics (Sparkes et al., 2009a; Ueda et al., 2010), work with mutants of XI-1, and XI-2, shows that they have little effect on their own, but enhance the effect of XI-K when double or triple mutants are analyzed. Here, we further explore whether these different subclasses of myosin XI that reduce spheroid organelle mobility differentially affect the movement and remodeling of the ER network. Remodeling is assessed by quantifying the static elements in the network which should increase if motor activity is required to drive changes.

## **MATERIALS AND METHODS**

#### **PLANT MATERIAL AND CONSTRUCTS**

*Nicotiana tabacum* plants were grown according to Sparkes et al. (2005). Fluorescent protein fusion constructs including the ER marker GFP-HDEL (Batoko et al., 2000), and mRFP-myosin XI-K, XI-I, XI-1, XI-2, XI-E, XI-C, XI-A tail domains (Sparkes et al., 2008; Avisar et al., 2009) were all infiltrated according to Sparkes et al. (2006) with an optical density of 0.1 except for GFP-HDEL, which required 0.04 optical density. Expression was analyzed 3 days following inoculation.

#### **PROTEIN EXTRACTION AND WESTERN BLOTTING**

Total proteins were extracted according to Gao et al. (2013). 0.5 g of tobacco leaf material 3 days post infiltration were ground in PEB (50 mM Tris–HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% TritonX-100 plus protease inhibitors) and then centrifuged. Equal volumes of extract were separated by 10% SDS-PAGE and blotted onto PVDF membrane (Pall). mRFP fusions were detected using anti-mRFP primary antibody (Abcam) and HRP-conjugated goat anti-rabbit secondary antibody (Abcam). Chemiluminescence reaction was performed using ECL substrate (Pierce) followed by film exposure.

#### **SAMPLE PREPARATION AND IMAGE ACQUISITION**

The ER in the outermost cortical region of adaxial leaf epidermal pavement cells was imaged. Dual imaging of mRFP and GFP was done using multi-tracking in line switching mode on a Zeiss LSM510 Meta confocal microscope. GFP was excited with a 488 nm argon laser and mRFP with a 543 nm laser and their emissions detected using a 488/543 dichroic mirror and 505–530 and 560–615 nm band pass filters, respectively. All imaging was carried out using a 63 × 1*.*4 numerical aperture oil immersion objective.

## **PERSISTENCY MAPPING THE CORTICAL ER IN TOBACCO EPIDERMAL CELLS**

For persistency mapping, time-lapse images of GFP-HDEL were captured using a 2–3µm pinhole, 512 × 512 pixel resolution and 2.3× digital zoom. To reduce noise, 4× line averaging was used. The scan rate was increased by imaging a 955–960µm2 region of interest (ROI), so that 50 frames per 80 s were captured (0.63 frames/s). For samples where cells were coexpressing a fluorescent myosin tail domain and GFP-HDEL, coexpression was verified before time lapse imaging of the GFP-HDEL alone was performed.

Persistency maps were generated in Image J (version 1.45s, Wayne Rasband, National Institute of Health, Bethesda, MD) as described in Sparkes et al. (2009a) with the following modifications. As shown in the corrected tubule persistency maps in **Figures 2**, **3**, the persistent tubule subset was corrected by subtracting regions containing cisternae prior to making persistent tubule counts. This was done by directly subtracting the morphologically opened binary sum images from the morphologically closed skeletonized binary sum images. In the resulting image sets, only those with a projected area *>* 0.2µm2 were counted (excludes tubules less than 1µm long, assuming a 200 nm projection of an individual tubule). This disconnected some of the tubules because punctae or small cisternae often occur at tubule junctions, producing shorter tubules of smaller percentage area than previously described. As shown in the corrected cisternal persistency maps of **Figures 5**, **6**, the persistent cisternal subset was corrected by subtracting regions that contained continuous flow (i.e., subtracting the sum of the 5-frame differences from the cisternal persistency map). Quantitation of the most persistent tubules, and cisternae was done, normalizing each movie to the total membrane area imaged in the percentage area and number/100µm2 values. Analysis of mesh size and mesh number per cytoplasmic volume was done with manual selection of the region containing in-focus signal for each movie, then measuring the number and area of selected mesh regions for each frame of every movie. The cytoplasmic volume was estimated as the area of the region containing in-focus signal times the z-dimension optical section acquired by the confocal microscope. Analysis of the size classes of persistent cisternae was done by making a binned histogram of the different sizes of persistent cisternae for the entire collection of movies acquired for each treatment. Note, highly persistent regions are defined as being present (persisting) for more than 23 s within the movie. Statistical comparison of the data was done using the 64 bit version of R, 3.0.2 (http:// www*.*r-project*.*org/) using ANOVA and the Tukey HSD two-way comparison of means at 95% confidence interval. Each analysis is based on data generated from 15 movies in total taken from separate cells from three independent experiments.

Displaced frame difference images were made by subtracting every fifth frame, summing the differences between all frames, and dividing that value by an estimated cytoplasmic volume (area containing signal in a 3µm optical slice) for each movie.

## **RESULTS**

## **MYOSIN TAIL DOMAINS AFFECT ER MORPHOLOGY**

As previously reported, transient expression of mRFP-myosin tail fusions in tobacco leaf epidermal cells are present in puncta or diffusely throughout the cytoplasm, with XI-I collating to the nucleus and motile puncta (**Figure 1**; Avisar et al., 2009). A similar perinuclear localization for both full length and tail fusions of XI-I in Arabidopsis and tobacco respectively were reported by Tamura et al. (2013). Here, we show how these fusions relate to, and affect the morphology and remodeling of the ER. **Figure 1** highlights that whilst these fusions do not solely collocate to the surface of the ER, they appear to differentially affect the level of ER cisternalization with XI-I tail resulting in the largest observable regions of cisternal ER (**Figures 1P–R**). This phenomenon is clearly visible when comparing the morphology of the ER in two adjacent cells where one has no detectable levels of mRFP-XI-I tail fusion (**Figures 1P–R**, white line defines cell boundary). These still images do not convey whether myosin tail expression affects the dynamic remodeling of the ER and so time lapse movies were taken and dynamics quantified.

## **MYOSIN TAIL DOMAINS AFFECT GLOBAL ER REMODELING**

Movies were taken using consistent settings allowing comparisons between the effects of the myosin tail domains on ER dynamics to be drawn. In addition, the same settings were used to detect mRFP myosin tail fusions to determine whether any effects on ER dynamics were dependent on myosin tail expression level. Western blotting confirmed that the majority of the myosin tail fusions were stable under transient expression in tobacco leaf epidermal cells (**Figure S1**). Results indicated that whilst XI-A (**Movie S2**) tail domain didn't appear to affect global ER remodeling compared with control cells only expressing the ER marker (GFP-HDEL, **Movie S1**), the other tail domains perturbed remodeling to differing extents in a concentration independent manner; XI-2 slowed down movement (**Movie S3**), XI-I increased the level of cisternal ER which was still relatively mobile (**Movie S4**), whereas XI-C, XI-E, XI-K, and XI-1 all reduced active remodeling (**Movies S5**–**S9**).

## **QUANTIFICATION OF THE EFFECTS OF MYOSIN TAIL DOMAINS ON ER DYNAMICS**

To quantify the effects of myosin tail domains on ER dynamics we implemented persistency mapping to tease out regions which had become more or less persistent compared with control cells. Persistency mapping can be used to measure the more static elements within the ER network over the given time frame. Morphological elements of the ER under study include tubules, cisternae (areas larger than 0.3µm2) and punctae (areas smaller than 0.3µm2). If the different myosins act as change agents, then their dominant-negative selective inhibition should decrease change, thereby increasing the number or size of the more persistent morphological elements. Therefore, we have chosen here and

**FIGURE 1 | Coexpression of myosin tail domains with an ER marker.** Representative images of cells coexpressing GFP-HDEL (green) and mRFP myosin tail domain fusions [magenta; XI-A, **(A–C)**; XI-1, **(D–F)**; XI-2, **(G–I)**; XI-C, **(J–L)**; XI-E, **(M–O)**; XI-I, **(P–R)**; XI-K, **(S–U)**]. White line in panels **(P–R)** highlights the cell boundary between neighboring cells. Scale bar 5µm.

in prior work (Sparkes et al., 2009a) to measure the more persistent features (green to black in **Figures 2**, **3**, **5**, **6**) of the persistency map rather than less persistent features (yellow in **Figures 2**, **3**, **5**, **6**). The analysis here is further refined over prior work by

#### **FIGURE 2 | Effects of XI-K, XI-C, XI-E, and XI-1 myosin tail domains on persistent ER tubules identified with co-expressed GFP-HDEL.**

Left-hand panel is the first frame of a 50 frame movie. Middle two panels are tubule persistency expressed as the sum of skeletonized tubules that persist for five frames (7.9 s).The corrected tubule persistency subtracts out contributing skeletons from cisternal regions. The yellow-green-blue-black persistency scale represents persistency from 7.9 to 80 s. Highly persistent tubules are those that are counted for **Figure 4**, being both greater than 0.2µm2 and persisting for at least a third of total movie time (black to green on the persistency scale with black representing persistency through the whole 80 s and green representing persistency for approximately 23 s). Note, control refers to cells only expressing the ER marker GFPHDEL (green, left hand panel).

excluding cisternal regions from tubular persistency analysis and by excluding large moving cisternal fields occurring throughout the analysis period (80 s) from the cisternal persistency analysis.

In general, persistency mapping shows an increase in persistent ER tubules (**Figures 2**–**4**), cisternae (**Figures 5**–**7**), and puncta (**Figures 5**, **6**, **8**) in cells expressing certain myosin tail fusions compared to control cells. In all of the persistency map figures (**Figures 2**, **3**, **5**, **6**) the image in the first column is the first frame from a representative movie and the additional columns highlight the process of isolating the highly persistent regions in the movie shown. These highly persistent regions (i.e., present for more than 23 s) are then quantified in terms of average number, size, and the percentage of imaged membrane represented by these morphological structures. Statistical analysis (HSD Tukeys test)

of pair wise combinations of average values are shown in **Table 1** (*n* = 15). **Movies S1**–**S8** highlight the persistent ER elements in **Figures 2**, **3**, **5**, **6**.

approximately 23 s). Note, control refers to cells only expressing the ER

marker GFPHDEL (green, left hand panel).

Whilst there is a large variation within the sample population (*n* = 15), statistical analysis highlighted that average persistent tubule area and number were significantly higher in cells coexpressing the tails of XI-C, XI-E, XI-K, or XI-1 compared to the control (**Figures 2**–**4**, **Table 1**). This corresponded with an increase in mesh size of the tubular network (**Figure S2**). In addition, there appeared to be significant pair wise differences between several of these myosins except for XI-C, XI-E, and XI-1 which are presumably performing similar functions relating to ER tubule dynamics/formation.

There was also a significant increase in the percentage of total membrane imaged present in persistent cisternal regions for several myosin tails (XI-1, XI-C, and XI-K) compared with the control (**Figures 5**–**7**, **Table 1**, a histogram of persistent cisternal size classes for each combination is provided **Figure S3**).

These significant differences could relate to changes in either average size and/or number of persistent cisternae per given unit area. Both XI-1 and XI-K's effects were due to a combination of both increased number and average size of cisternal regions (**Figure 7**). XI-E and XI-I also resulted in significantly more cisternae. In addition the average area and percent area increased with XI-E and XI-I tail domain expression over control, but the data are so variable (note large standard deviations) that a firm statement of difference cannot be made. This high variability quite likely arises from the observations based on the movies, as described above, that cisternalization increases but a fairly large proportion remain motile. While the number of relatively immobile cisternae increases, there is enough general membrane cisternalization (hence larger amounts of membrane per field) that the area fraction of immobile cisternae does not reliably increase.

Pair wise statistical tests for significant differences between XI-1, XI-C, and XI-K persistent cisternal criteria indicate that XI-1 and XI-K are not different to one another. However, XI-C compared with either XI-1 or XI-K display differences in persistent cisternal number, but not persistent cisternal area. This could indicate that XI-1 and XI-K have conserved functions in generating the number and size of persistent ER cisternae, whereas XIC is more involved in maintaining cisternal number, but not size relative to the roles of XIK and XI-1.

Persistent regions of ER less than 0.3µm2, termed persistent punctae, are present in control cells and in all cells coexpressing the myosin tail domains. XIC tail domain is the only myosin to have a significant increase in the number of persistent punctae over control conditions, and when compared with other myosins (**Figures 5**, **6**, **8**, **Table 1**).

Finally we tried to monitor the active regions of the ER network using displaced frame differences (**Figure S4**). Here, the sum of the displaced frame differences summed over the imaged volume were plotted for all myosin combinations. As expected with increased persistency for XI-K, XI-C, XI-1, and XI-E we have a slower rate of movement in these samples compared to the control, XI-2 and XI-A. This provides an initial approximation to the dynamic elements of the network.

## **DISCUSSION**

ER network structure (tubules/cisternae and three way junctions) is not completely dependent on cytoskeletal elements; depolymerization of actin results in an interconnected ER network which is more cisternal (Sparkes et al., 2009a, 2011). At present, additional molecular factors required to alter ER morphology in plants include reticulons and RHD3 (see references in reviews Sparkes et al., 2011, and Zhang and Hu, 2013). Here, we have begun to assess the relative role(s) of class XI myosins on ER geometry and remodeling. Several class XI myosins which appear to have a global effect on perturbing the movement of several organelle classes including peroxisomes, Golgi and mitochondria were investigated (XI-C, XI-E, XI-I, XI-K, XI-1, and XI-2; (Avisar et al., 2008, 2009; Peremyslov et al., 2008, 2010; Prokhnevsky et al., 2008; Sparkes et al., 2008, 2009a). XIA was also tested as the tail domain fusion did not affect global organelle dynamics in previous studies, and so was used here as a potential control for overexpressing a class XI myosin.

The main findings from this study indicate that whilst XI-A did not affect ER dynamics, XI-C, XI-E, XI-K, and XI-1 affected tubule persistency and the persistent nature of cisternae, and XI-C increased the number of persistent punctae. Whilst XI-I and XI-2 didn't affect the persistent regions within the network, visual inspection indicates that XI-I increases the level of cisternalization, while XI-2 reduces overall remodeling (**Movies S3**, **S4**). These differential affects could indicate specific roles for myosins in controlling tubulation (growth/shrinkage/lateral sliding to form polygons), cisternalization, and potential anchoring of the ER to the plasma membrane. For example, based on the differences these myosin tail domains exert on static regions within the ER network, it is interesting to speculate that XI-C, XI-E, XI-K, and XI-1 are required for most forms of tubule growth and shrinkage as overexpression of the tail domains increased the number of static tubules. This is assuming that the tail domains are acting in a dominant negative manner and the observed effect is due to a direct rather than indirect effect. It is likely that tubule growth draws upon, in part, a reservoir of membrane in the cisternal regions of the ER rather than being due to extension of completely newly synthesized membrane (Sparkes et al., 2009a). Mesh size of the tubular network increases in those treatments

that increase tubule persistency (**Figures 2**, **3** compared with **Figure S2**), indicating a lower amount of total tubulation. This explains why the myosin tail domains which increase the number of persistent tubules also have a concomitant increase in persistent cisternal size or number. The less obvious, but nonethe-less significant effect, (**Figure 7**, **Table 1**) of XI-I increasing the number of persistent cisternae relative to control may relate to a shaping role in the ER of known cisternal structures, such as the nuclear envelope, where XI-I tends to accumulate (**Figure 1**). Hence, XI-I may have more shaping-related and less movementrelated functions, while XI-C, XI-E, XI-K, and XI-1 may have more movement-related and less shaping-related functions. XI-2 tail domain expression doesn't affect the level of persistency in the ER, but appears to reduce overall modeling.

subtracts out regions of continuous flow (sum of five frame differences). The

It is important that we comment upon the large variability seen within each sample population presented herein. Organelle dynamics are affected by growth and developmental cues, with larger, older cells showing faster ER and other organelle movement than smaller, younger cells (Stefano et al., 2014). The cell population analyzed here, the pavement epidermal cells between the minor veins of fully-expanded tobacco leaves, was chosen because it is amenable to rapid transient expression assays and provides easily-imaged, robust cells. However, this population varies somewhat in cell size and developmental stage. This could be one source of the variability. Other approaches, such as using myosin mutants, may provide analysis of a single developmental stage of a uniformly-sized cell population, but would suffer from the developmental defects caused by this mutation. Another source of variability in this work could arise from subcellular variations of movement within these puzzle-piece-shaped cells, which may have tip growth dynamics in the interdigitating lobes and non-tip growth dynamics along the plane of the outer wall. The pattern of cytoplasmic movement in cells undergoing polar tip vs. radial growth are radically different (Hepler et al., 2001). Unfortunately, technical limitations prevent acquiring data sets of adequate resolution

refers to cells only expressing the ER marker GFPHDEL (green, left hand panel).

to resolve the ER network in the entire cell within a fraction of a second. However, were that possible, different patterns of movement might be sorted and the ER dynamics within each pattern analyzed. Advances in imaging will hopefully overcome these limitations.

Previous studies using RNAi, expression of myosin truncations and observations in null mutant backgrounds have shown several myosins appear to have a global affect on the motility of all organelles tested (Avisar et al., 2008, 2009, 2012; Peremyslov et al., 2008, 2010; Prokhnevsky et al., 2008; Sparkes et al., 2008, 2009a). Interestingly, as detailed above, the same myosin truncations that globally arrest spheroid organelle movement have differential effects on ER dynamics and persistent regions. The effects of XI-I on spheroid dynamics could have been due to increased ER cisternalization reducing the effective void volume (cytoplasm) in which spheroid organelles could move thus reducing their motility. XI-2 reduces the movement of all organelles tested, including the ER to a certain extent, but doesn't affect the level of persistency. This could indicate that XI-2 affects a more global common element in organelle dynamics such as the actin cytoskeleton or perhaps cyclosis. Further examination of the nature of the change in the dynamics under control conditions, and in comparison with XI-2 tail domain expression (see **Movie S3**), awaits other forms of analysis. None-the-less, these results indicate that the myosin tail domains that globally affect the movement of spheroid organelles have differential effects on ER network dynamics; increased persistency (XI-1, XI-C, XI-K, XI-E) vs. reduced remodeling (XI-2) vs. drastic changes in ER cisternalization (XI-I).

The expression profiles of the myosins under study here indicate that XI-1, XI-2, XI-I, and XI-K are ubiquitously expressed throughout the plant, whereas XI-C and XI-E are mainly expressed in the stamen and pollen (Peremyslov et al., 2011; Sparkes et al., 2011). Unlike leaf epidermal cell, pollen tubes undergo polarized tip growth where organelle movement occurs via reverse fountain streaming (Hepler et al., 2001; Lovy-Wheeler et al., 2007). This type of growth and pattern of organelle movement could place additional demands on ER morphology; persistent punctae could possibly represent tethering/interaction with the plasma membrane to maintain geometric distribution throughout growth. Ridge et al. (1999) documented changes in ER morphology during developmental transitions in polarized root hair growth; upon completion of growth the ER is tubular whereas during the elongation phase a more cisternal form prevails some way behind the tubular ER network in the expansion zone at the tip.

**FIGURE 7 | Quantification of the effects of myosin tail domains on persistent ER cisternae. (A)** Counts of the cumulative percentage area of highly persistent cisternae (see **Figures 5**, **6** for examples). **(B)** Counts of the number of persistent cisternae per hundred square micrometers of ER membrane analyzed for all of the myosin subclasses. **(C)** Average area of persistent cisternae in square micrometers. Error bars represent standard deviation. The level of significance of the difference between the samples is described in **Table 1**.

Interestingly, based on phylogenetic analysis of the myosin motor domains it appears that XI-C, XI-E, XI-K, and XI-1 from *Arabidopsis* are from closely related subfamilies XI(J) and XI(K). This could therefore indicate that they might perform similar/conserved functions in controlling ER dynamics (Peremyslov et al., 2011). Whereas XI-A, XI-I, and XI-2 are from more distantly related subfamilies XI(G), and XI(I).

Myosins have been implicated in ER formation and/or dynamics in several plant systems. Immunoblotting of subcellular fractions from Arabidopsis leaf material indicated that XI-K

highly persistent punctae (see **Figures 5**, **6** for examples). **(B)** Counts of the number of persistent punctae per hundred square micrometers of ER membrane analyzed for all of the myosin subclasses. Error bars represent standard deviation. The level of significance of the difference between the samples is described in **Table 1**.

cosedimented with an ER fraction, and that the streaming of the ER is drastically reduced in *xi-k* mutants and to a lesser extent in *xi-1* and *xi-2* (Ueda et al., 2010). Independent studies place XI-K on motile vesicles, and appears to interact and collocate with members from a novel myosin receptor family (Peremyslov et al., 2012, 2013). Moreover, immunolocalization studies indicated that XI-2 partially localized to peroxisomes in *Arabidopsis* (Hashimoto et al., 2005), whereas a 175 kDa tobacco homolog with 75% identity to XI-2 was proposed to control ER movement in tobacco BY-2 cells (Yokota et al., 2009). *In vitro* reconstitution experiments from tobacco BY-2 cells further supported a role for the 175 kDa myosin in ER tubule formation (Yokota et al., 2011). More recently, expression of the tail domain of a maize XI-I homolog Opaque1 in *N. benthamiana* disrupted ER streaming, and subcellular fractionation indicated it is also associated with the ER in maize (Wang et al., 2012). Interestingly, Ueda et al. (2010) also highlighted that actin organization is perturbed in *xik xi-2* double mutants, but not single mutants, indicating that both myosins act in a cooperative/synergistic manner to control actin organization in elongating cells. Defects in actin organization were also seen in the midvein cells of triple (*xi-k/1/2*) and quadruple (*xi-k/1/2/i*) Arabidopsis mutants (Peremyslov et al., 2010). However, actin dynamics rather than organization were

*Tukeys HSD comparison of means test to compare persistent regions in the ER generated upon expression of myosin tail domains in tobacco leaf epidermal cells. Significant differences between pair wise combinations under test are shown (\*p < 0.05; \*\*p < 0.005; \*\*\*p < 0.0005).*

affected in root hairs of an *xi-k* null line (Park and Nebenführ, 2013). In light of all of these studies, it is interesting to note that we found XI-2 may have a global affect on organelle dynamics which could be an indirect effect through perturbing actin organization. Therefore, organelle-myosin specificity, and the specific affect of myosins on actin dynamics, is still a matter of debate, and whether these differences relate to species or tissue specific effects is unclear. Determining the mechanism of myosin association and disassociation from targets is complex (Li and Nebenführ, 2007, 2008a,b). A recent publication from Peremyslov et al. (2013) highlighted a potential myosin receptor family in plants which will hopefully help resolve organelle-myosin specificity.

Persistency mapping is an excellent tool for quantifying the more static geometric elements within the dynamic ER network, which is continually undergoing rapid geometric rearrangements. It does not, however, monitor ER luminal or surface flow. Prior work (Sparkes et al., 2009a) has shown that non-directional surface flow as measured with FRAP and photoactivation still occurs when actin is depolymerized, a condition that drastically changes ER geometry. Optical flow analysis of fluorescent ER constituent proteins also shows that actin depolymerization does not completely inhibit movement (Stefano et al., 2014). In single mutant *xi-1*, there is only a small change in ER movement as analyzed with optical flow (Ueda et al., 2010), but XI-1 tail domain expression has a large effect on membrane geometry (**Figures 1**, **2**, **4**, **5**, **7**). Persistency mapping therefore provides the only tool currently available to quantify differences in ER network geometry. Further studies are required to determine whether the same myosins which affect ER network dynamics also affect surface flow. Better analytical tools are required to quantify the complex dynamic regions which consist of tubule growth, shrinkage, lateral sliding, changes from tubular to cisternal form, formation and movement of three way junctions, polygon formation, size, movement and potential filling to form cisternae. Whilst these dynamic elements can be broken down into various aforementioned categories, ER remodeling is a complex combination where each morphological element is interdependent requiring a more sophisticated cross correlative approach. Our initial displaced frame difference provides an initial observation of such dynamic elements. Furthermore, high persistency described here does not distinguish between elements which are static over sequential frames lasting more than 23 s vs. temporally episodic events over a similar discontinuous time frame. However, observation of the movies clearly shows that persistent elements are consistently static rather than being due to events which reoccur. In addition, it is worthwhile noting that whilst the studies presented here relate to real time events, they are limited by the spatial resolution of the microscope. In essence, the ER tubules studied are likely to be around the 50–70 nm diameter making the 200 nm resolution limit of the microscope prone to potentially imaging parallel tubules rather than reflecting single tubules. Only through the development of super resolution imaging systems, which allow imaging in real time, will these types of issues be resolved.

### **AUTHOR CONTRIBUTIONS**

Imogen Sparkes designed the experiments, Lawrence R. Griffing carried out the persistency mapping and statistical analysis, Imogen Sparkes, Lawrence R. Griffing, and Hongbo T. Gao acquired the data. All authors were involved in writing and approving the manuscript.

## **ACKNOWLEDGMENTS**

This work was supported by BBSRC grant BB/1006184/2 for Hongbo T. Gao and Imogen Sparkes and the Biology department at Texas A and M University for Lawrence R. Griffing.

### **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at: http://www*.*frontiersin*.*org/journal/10*.*3389/fpls*.*2014*.*00218/ abstract

**Figure S1 | Western blot analysis of transiently expressed mRFP-myosin tail fusions in tobacco leaf epidermal cells.** Arrow heads denote expected sizes of full length mRFP myosin tail fusions (∼100 kDa apart from XI-A which is ∼124 kDa). Asterisk shows detection of full length St-mRFP used as a control. Differences in relative myosin levels are due to differences in the level of transient expression. A large proportion of each myosin fusion is stable.

**Figure S2 | Mesh number per estimated cytoplasmic volume and ER mesh size.** The ER tubules generate open polygons that can be considered as elements of a mesh. Blue bars: The polygonal regions subtended by tubules were counted for each frame in each movie, averaged over the entire movie, divided by the estimated cytoplasmic volume, and shown as the average number of meshes per 300µm3 of cytoplasm for all of the movies taken for each treatment. XI-1, XI-C, XI-E, XI-I, and XI-K are all significantly different to the control and XI-A (*p <* 0*.*005; Tukeys HSD comparison of means). Red bars: The average area, in square micrometers, of each polygonal mesh structure for each frame of each movie was also calculated. The average mesh area over all of the movies of each treatment is shown. XI-C, XI-E, and XI-K are all significantly different to the control (*p <* 0*.*005; Tukeys HSD comparison of means). The mesh areas of XI-K and XI-E were also significantly different (*p <* 0*.*05) from XI-2, XI-A, and XI-I. Error bars are standard deviation.

## **Figure S3 | Histogram of the percent of each persistent cisternae in different size classes for each treatment.** The distribution of the percentage that each different size class makes to the population of persistent cisternae produced by each treatment is shown. The value of each size class was chosen so that comparisons could be made between those that had predominantly small cisternae (e.g., control) and those that had very large cisternae (e.g., XI-1 and XI-K).

**Figure S4 | Relative movement or flow as shown by average displaced frame differences summed over imaged volumes.** The value of the intensity difference between every fifth frame, the displaced frame difference, is summed for each movie and divided by the approximate volume of the region imaged (area of signal in a 3µm optical section). The average of these values is shown for all of the movies acquired for each treatment. Error bars are standard deviation. XI-1, XI-C, XI-E, XI-K, and XI-I are significantly different to the control (*p <* 0*.*005; Tukeys HSD comparison of means), and to XI-A (*p <* 0*.*05; Tukeys HSD comparison of means), and XI-1, XI-C, XI-E, and XI-K are significantly different to XI-2 (*p <* 0*.*05; Tukeys HSD comparison of means).

**Movie S1 | Persistency mapping of ER dynamics in control tobacco leaf epidermal cells.** Time lapse movie showing the movement of the ER, along with overlays of the persistent tubules, cisternae, and puncta (magenta). Scale bar 10µm.

**Movie S2 | Persistency mapping of ER dynamics in a tobacco leaf epidermal cell expressing the myosin tail domain XI-A.** Time lapse movie of the ER in a cell coexpressing the tail domain of XI-A. Overlays of the persistent tubules, cisternae and puncta (magenta) are also shown for comparison. Scale bar 10µm.

**Movie S3 | Persistency mapping of ER dynamics in a tobacco leaf epidermal cell expressing the myosin tail domain XI-2.** Time lapse movie of the ER in a cell coexpressing the tail domain of XI-2. Overlays of the persistent tubules, cisternae and puncta (magenta) are also shown for comparison. Scale bar 10µm.

**Movie S4 | Persistency mapping of ER dynamics in a tobacco leaf epidermal cell expressing the myosin tail domain XI-I.** Time lapse movie of the ER in a cell coexpressing the tail domain of XI-I. Overlays of the persistent tubules, cisternae and puncta (magenta) are also shown for comparison. Scale bar 10µm.

**Movie S5 | Persistency mapping of ER dynamics in a tobacco leaf epidermal cell expressing the myosin tail domain XI-C.** Time lapse movie of the ER in a cell coexpressing the tail domain of XI-C. Overlays of the persistent tubules, cisternae and puncta (magenta) are also shown for comparison. Scale bar 10µm.

**Movie S6 | Persistency mapping of ER dynamics in a tobacco leaf epidermal cell expressing the myosin tail domain XI-E.** Time lapse movie of the ER in a cell coexpressing the tail domain of XI-E. Overlays of the persistent tubules, cisternae and puncta (magenta) are also shown for comparison. Scale bar 10µm.

**Movie S7 | Persistency mapping of ER dynamics in a tobacco leaf epidermal cell expressing the myosin tail domain XI-K.** Time lapse movie of the ER in a cell coexpressing the tail domain of XI-K. Overlays of the persistent tubules, cisternae and puncta (magenta) are also shown for comparison. Scale bar 10µm.

**Movie S8 | Persistency mapping of ER dynamics in a tobacco leaf epidermal cell expressing the myosin tail domain XI-1.** Time lapse movie of the ER in a cell coexpressing the tail domain of XI-1. Overlays of the persistent tubules, cisternae and puncta (magenta) are also shown for comparison. Scale bar 10µm.

#### **Movie S9 | Comparison of ER dynamics in two tobacco leaf epidermal cells expressing relatively different levels of mRFP-XI-1 tail domain fusion.**

Fluorescent image highlighting mRFP-XI-1 tail fusion (magenta) and the effects on ER dynamics in the same cell are shown (green). Two representative movies from two cells show similar dynamics in cells with relatively different levels of myosin tail expression.

## **REFERENCES**

cells mimics responses to fungal and oomycete attack. *BMC Plant Biol.* 8:63. doi: 10.1186/1471-2229-8-63

Yokota, E., Ueda, S., Tamura, K., Orii, H., Uchi, S., Sonobe, S., et al. (2009). An isoform of myosin XI is responsible for the translocation of endoplasmic reticulum in tobacco cultured BY-2 cells. *J. Exp. Bot.* 60, 197–212. doi: 10.1093/jxb/ern280

Zhang, M., and Hu, J. (2013). Homotypic fusion of endoplasmic reticulum membranes in plant cells. *Front. Plant Sci.* 4:514. doi: 10.3389/fpls.2013.00514

Zheng, M., Beck, M., Muller, J., Chen, T., Wang, X., Wang, F., et al. (2009). Actin turnover is required for myosin-dependent mitochondrial movements in *Arabidopsis* root hairs. *PLoS ONE* 4:e5961. doi: 10.1371/journal.pone.0005961

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 23 February 2014; accepted: 01 May 2014; published online: 21 May 2014. Citation: Griffing LR, Gao HT and Sparkes I (2014) ER network dynamics are differentially controlled by myosins XI-K, XI-C, XI-E, XI-I, XI-1, and XI-2. Front. Plant Sci. 5:218. doi: 10.3389/fpls.2014.00218*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Griffing, Gao and Sparkes. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## ER and vacuoles: never been closer

## *Corrado Viotti\**

*Umeå Plant Science Centre, Department of Plant Physiology, Umeå University, Umeå, Sweden*

#### *Edited by:*

*Lorenzo Frigerio, University of Warwick, UK*

#### *Reviewed by:*

*Alessandro Vitale, National Research Council of Italy, Italy Byung-Ho Kang, University of Florida, USA Enrique Rojo, Centro Nacional de Biotecnología – Consejo Superior de Investigaciones Científicas, Spain*

#### *\*Correspondence:*

*Corrado Viotti, Umeå Plant Science Centre, Department of Plant Physiology, Umeå University, Linnéusväg 6, 90187 Umeå, Sweden e-mail<EMAIL_ADDRESS>
The endoplasmic reticulum (ER) represents the gateway for intracellular trafficking of membrane proteins, soluble cargoes and lipids. In all eukaryotes, the best described mechanism of exiting the ER is via COPII-coated vesicles, which transport both membrane proteins and soluble cargoes to the *cis*-Golgi. The vacuole, together with the plasma membrane, is the most distal point of the secretory pathway, and many vacuolar proteins are transported from the ER through intermediate compartments. However, past results and recent findings demonstrate the presence of alternative transport routes from the ER towards the tonoplast, which are independent of Golgi- and post-Golgi trafficking. Moreover, the transport mechanism of the vacuolar proton pumps VHA-a3 and AVP1 challenges the current model of vacuole biogenesis, pointing to the endoplasmic reticulum for being the main membrane source for the biogenesis of the plant lytic compartment. This review gives an overview of the current knowledge on the transport routes towards the vacuole and discusses the possible mechanism of vacuole biogenesis in plants.

**Keywords: endoplasmic reticulum, COPII vesicles, Golgi apparatus, trans-Golgi network, multivesicular body, vacuole**

## **ENDOPLASMIC RETICULUM: ENTRANCE TO THE SECRETORY PATHWAY**

The endoplasmic reticulum (ER) consists on a network of interconnected membrane tubules and cisternae ("reticulum") stretching across the entire cytoplasm ("endoplasmic"). First discovered in culture cells from chicken embryos (Porter et al., 1945), the ER is present in all eukaryotic cells, and is the intracellular compartment where membrane proteins, soluble cargoes and lipids are synthesized. From the ER, correctly folded membrane and soluble proteins are transported to other endomembrane compartments or to the extracellular space along the secretory pathway (Vitale and Denecke, 1999). For all eukaryotes, the best characterized mechanism of exiting the ER is the COPII-mediated transport. The coat protein complex II (COPII) assembles on specific locations of the ER membrane, called ER-exit sites (ERES), from which COPII-coated vesicles bud off. The assembly of COPII begins with the activation of the small guanosine triphosphatase (GTPase) SAR1 provided by the ER membrane-bound guanine nucleotide exchange factor (GEF) SEC12, which leads to the coordinated recruitment of the cytosolic heterodimers SEC23/SEC24 and SEC13/SEC31 to the ERES (Nakano et al., 1988; Barlowe and Schekman, 1993; Barlowe et al., 1994). Cargo recognition is provided by SEC24 and SAR1, whereas multiple adjacent SEC13/SEC31 subcomplexes drive the bending of the ER membrane using the energy of GTP hydrolysis (Brandizzi and Barlowe, 2013). Passive incorporation of soluble cargoes into COPII vesicles can occur (Wieland et al., 1987; Denecke et al., 1990; Matsuoka and Nakamura, 1991; Phillipson et al., 2001; Thor et al., 2009), instead membrane proteins and receptors require diacidic or di-hydrophobic motifs in their cytosolic domains for efficient transport (Kappeler et al., 1997; Nishimura and Balch, 1997; Contreras et al., 2004; Hanton et al., 2005). In mammals, most COPII subunits have one or more paralogs, which generate a robust repertoire of COPII-coated vesicles with tissue specificities and selectivity for different cargo molecules (reviewed in Zanetti et al.,2011). In plants much less is known about specificities among different COPII-coated carriers, even though it has been recently shown that the concomitant function of all three SEC24 members of *Arabidopsis* is necessary for the development of the gametophytes (Conger et al., 2011; Tanaka et al., 2013). After a long debate whether COPII vesicles versus COPII-coated tubules existed in plant cells, ultrastructural analysis of high-pressure frozen samples and 3D tomography reconstructions have shown that COPII vesicles are present also in plants (Ritzenthaler et al., 2002; Donohoe et al., 2007; Robinson et al., 2007; Kang and Staehelin, 2008).

## **LYTIC VACUOLES**

The plant lytic vacuole can occupy up to 90% of the total volume in mature vegetative cells. Its remarkable size allowed Antonie van Leeuwenhoek to notice the vacuole already in the 1670s, at the dawn of microscopy. The name "vacuole" was coined from "vacuum," because Felix Dujardin, in 1872, thought he was facing an empty space (Leigh and Sanders, 1997; De, 2000). On the contrary, the vacuolar content can generate a stationary turgor pressure of up to five bars (Zimmermann et al., 1980), which provides the driving force for plants' growth by pushing the cells to expand in oriented directions. Moreover, the lytic vacuole plays a crucial role in pH homeostasis, storage of ions, degradation of cellular waste, defense against pathogens, and in buffering abiotic stresses. The rapid release from or uptake to the vacuolar lumen of ions and water allow plants to efficiently cope with diversified environmental challenges. The multiple roles of plant lytic vacuoles are regulated by the activity of transporters that use the energy of the electrochemical gradient generated across the tonoplast by the vacuolar H+-ATPase (V-ATPase) and vacuolar H+-PPase (V-PPase). Despite good knowledge of the biochemistry and function of the vacuolar proton pumps (Maeshima, 2001; Schumacher and Krebs, 2010), little is known about the mechanisms of their sorting and the intracellular routes they follow to reach the tonoplast. However, recent data has shown that both the V-ATPase and V-PPase of *Arabidopsis* are incorporated to the tonoplast via a novel mechanism that also challenges the current model for vacuole biogenesis (Viotti et al., 2013).

## **GOLGI- AND POST-GOLGI-MEDIATED TRANSPORTS TO THE LYTIC VACUOLE**

Tonoplast-resident proteins and vacuolar soluble cargoes are synthesized in the ER, many of them are delivered to the *cis*-side of the Golgi apparatus via COPII vesicles, and from the Golgi they proceed further through the secretory pathway (**Figure 1**; Pedrazzini et al., 2013; Xiang et al., 2013). Vesicle transport between endomembrane compartments is mediated by different effector molecules, among which are the Rab GTPases, that are members of the ras superfamily of regulatory GTPases (Rutherford and Moore, 2002). The dissection of distinct steps of vacuolar transport using nucleotide-deficient mutants of different Rab GTPases in tobacco leaf epidermis cells has shown that tonoplast-resident proteins might follow at least three different routes (Bottanelli et al., 2011). In agreement with this finding, it has been shown that the sucrose transporter SUC4 and the *myo*-inositol transporter INT1 of *Arabidopsis* are delivered to the tonoplast in an adaptor protein complex 3 (AP3)-dependent and -independent manner respectively (Wolfenstetter et al., 2012). AP complexes sort cargo proteins into coated vesicles, and AP3 is involved in vacuolar trafficking. The exact localization of AP3 in plants is uncertain, because this adaptor seems to interact with clathrin (Lee et al., 2007; Zwiewka et al., 2011), which is present at the *trans*-Golgi network (TGN; Kang et al., 2011), whereas SUC4 accumulates at the Golgi apparatus in protoplasts isolated from *ap3* mutant seedlings, suggesting a Golgi-derived vesicle transport (Wolfenstetter et al., 2012).

From the TGN other two clathrin-mediated vacuolar transport carriers have been proposed to exist. Each of them has an EPSIN N-TERMINAL HOMOLOGY (ENTH) protein, EPSIN1 or MTV1 respectively, that acts as a monomeric adaptor for clathrin recruitment. Both EPSIN1 and MTV1 localize to the TGN, and both the respective knock-out mutants show defects in vacuolar transport (Song et al., 2006; Sauer et al., 2013).

The last and best known carrier in the vacuolar branch of the secretory pathway is the multivesicular body (MVB), which is an independent organelle (Tse et al., 2004) that arises from the TGN through a maturation process that involves the function of the TGN-located vacuolar proton pump VHA-a1, the calcium dependent phospholipid binding protein ANNEXIN 3, and the ESCRT-machinery (Scheuring et al., 2011). As in yeast and mammals, plant membrane proteins destined for degradation due to physiological turnover are incorporated to the MVB's intraluminal vesicles through the function of the ESCRT complexes, and then released inside the vacuole via MVB-to-vacuole fusion (Reichardt et al., 2007; Spitzer et al., 2009; Viotti et al., 2010; Scheuring et al., 2011). Soluble vacuolar cargoes dissociate from vacuolar sorting receptors (VSRs) in an acidic environment (Kirsch et al., 1996), therefore this event might occur in the lumen of the TGN, which is the most acidic organelle among the intermediate compartments

**FIGURE 1 | Model for lytic vacuole biogenesis in** *Arabidopsis***.** The precursors of vacuoles, the provacuoles, arise from the endoplasmic reticulum (ER) via maturation. The vacuolar proton pumps VHA-a3 and AVP1 (and perhaps other tonoplast proteins) are incorporated to the nascent provacuole directly from the ER-membrane through an uncharacterized mechanism (lower right corner). Golgi- and post-Golgi trafficking (solid arrows) continuously contribute, during the different steps of vacuole formation (dotted lines), to the development of the vacuolar lumen via the

delivery of soluble cargoes (e.g., acid proteases) and tonoplast-resident proteins that exit the ER with COPII-coated vesicles (dashed arrow). Provacuoles can also fuse with already developed vacuoles (*fusion*). Impairment of Golgi- and post-Golgi trafficking (crossed arrows) leads to aberrant multilayered provacuoles.This observation could be explained if impairment of Golgi- and post-Golgi trafficking would interfere (⊥) with the process of provacuoles-release from the ER, or with the fusion between provacuoles and vacuoles.

(Martinière et al., 2013; Shen et al., 2013). Soluble cargoes are then incorporated into the lumen of nascent MVBs, that arise from the TGN (Scheuring et al., 2011), for vacuolar transport. However, VSRs are localized both to the TGN and MVBs (Niemes et al., 2010; Stierhof and El Kasmi, 2010; Viotti et al., 2010), and the location from where they recycle is a matter of controversy (De Marcos Lousa et al., 2012; Robinson et al., 2012). An example of soluble cargo transported via MVBs is the cysteine protease aleurain (Miao et al., 2008), while regarding tonoplast resident proteins it has been recently shown that the auxin transporter WAT1 additionally colocalizes with the late endosomal marker RabG3f (Ranocha et al., 2013).

## **UNCONVENTIONAL ER-EXPORT OF PROTEINS TO THE VACUOLE**

The conventional transport of proteins to the vacuole involves COPII-mediated ER-exit and the passage through several intermediate steps and compartments. Hence, the vacuole, together with the plasma membrane, may be seen as the most distal point of the secretory pathway.

However, it was shown byGomez and Chrispeels (1993)that the tonoplast intrinsic protein α-TIP, unlike the soluble vacuolar protein phytohemagglutinin (PHA), can reach the lytic vacuole even after brefeldin A (BFA) or monensin treatment when transiently expressed in tobacco leaves, suggesting the presence of different vacuolar transport routes. Few years later, Jiang and Rogers (1998) showed that a chimera composed by the C-terminal domain of α-TIP fused to the transmembrane domain of the VSR BP80 reaches the protein storage vacuole (PSV) through a direct route from the ER. Evidence for an alternative mechanism of vacuolar trafficking was provided by the analysis of the calcineurin B-like (CBL) proteins, which are calcium sensors functioning in different locations within a cell (Batistiˇc and Kudla, 2009). Among the ten members of *Arabidopsis*, CBL2, CBL3, CBL6 and CBL10 are targeted to the tonoplast in a COPII-independent manner, since overexpression of a dominant-negative mutant of SAR1did not interfere with their localization (Batistiˇc et al., 2010). Moreover, it was shown that CBL6 is transported to the vacuole bypassing both the Golgi and post-Golgi compartments (Bottanelli et al., 2011). CBL proteins, however, seem not to enter the secretory pathway, but are rather synthesized in the cytosol and delivered to the tonoplast due to the presence of a tonoplast targeting signal (TTS) in their N-terminal domain (Bottanelli et al., 2011; Batistiˇc et al., 2012; Tang et al., 2012). An example of soluble cargo transported to the plant vacuole through an unconventional route is the human αmannosidase MAN2B1, which still reaches the vacuolar lumen even upon BFA treatment when transiently expressed in tobacco leaf mesophyll protoplasts (De Marchis et al., 2013).

The most abundant tonoplast resident protein, the vacuolar H+-ATPase VHA-a3, is transported to the vacuole through a novel mechanism. By blocking COPII-mediated transport via BFA treatment of GNL1 BFA-sensitive *Arabidopsis* seedlings (Richter et al., 2007), VHA-a3 was not retained in the ER and was detected as normal at the tonoplast, whereas the TGN-located H+-ATPase VHA-a1 was efficiently retained in the endoplasmic reticulum, indicating that VHA-a3 exits the ER in a COPII-independent manner (Viotti et al., 2013). Interestingly, while the N-terminal domain

of the a1 subunit carries a typical di-acidic motif (EE--D) for COPII-mediated export, in those of the a2 and a3 isoforms there is none. Moreover, in a β-AP3 knock-out mutant (Feraru et al., 2010)VHA-a3 was detected as normal at the tonoplast (Viotti et al., 2013), and its transport was not stopped at the level of intermediate compartments by using the post-Golgi-transport inhibitor concanamycin A (ConcA). Similarly, the second *Arabidopsis* vacuolar proton pump, the H+-PPase AVP1, did not accumulate to the Golgi/TGN interface upon ConcA treatment, and it did not localize to the limiting membrane of MVBs (Viotti et al., 2013). In other words, none of the known Golgi- and post-Golgi trafficking routes seemed to be involved in the delivery of the two vacuolar proton pumps.

AVP1 was not only detected to the limiting membrane of rounded vacuoles, but also uniformly present on the membranes of lytic vacuole precursors, the provacuoles. Provacuoles display a much finer (down to 30 nm thickness) tubular network in provascular cells of the root meristem, they are acidic, they carry VHA-a3 too, can fuse with already-developed vacuoles, and are distinct structures respect to autophagosomes (Viotti et al., 2013).

How is AVP1 transported from the ER to the provacuole and where does the latter originate from? A hypothesis is provided in the last section on this review.

## **MECHANISMS OF VACUOLE BIOGENESIS**

Relatively little is known about the biogenesis of vacuoles in plants. Even the donor membrane from where newly formed vacuoles originate from is unclear. The model that boasts most of the credits in text books suggests that newly formed lytic vacuoles in root-tip cells originate from post-Golgi-derived vesicles (Marty, 1999; De, 2000; Robinson and Rogers, 2000). These vesicles would homotypically fuse to form tubular structures that represent the precursors of vacuoles, the provacuoles. The tubular provacuoles are supposed to fuse with one another, forming a complex network that finally will give raise to the central vacuole (Marty, 1999). This hypothesis is based on an early electron microscopy study that revealed tubular structures at the *trans*-side of the Golgi apparatus which were strongly electrondense after incubation with sodium β-glycerophosphate or cytidine 5 -monophosphate (Marty, 1978). These two compounds serve as substrates to detect acid phosphatase and thiolacetic acid esterase activity respectively, thus they were used as biochemical markers to highlight acidic compartments. The tubular-vesicular structure at the *trans*-side of the Golgi was named Golgi-associated endoplasmic reticulum (GERL; Marty, 1978), and later it was renamed as TGN (Griffiths and Simons, 1986). Due to its acidic intraluminal pH, the TGN was proposed to represent the donor membrane for the biogenesis of the lytic vacuole (Marty, 1999).

A few years ago it was shown in *Arabidopsis* that the vacuolar H+-ATPase localizes also to the TGN, thus it is not a purely "vacuolar" enzyme. The V-ATPase (VHA) is a holoenzyme composed by two subcomplexes: the membrane-integral complex V0, and the cytosolic complex V1, both composed by multimeric subunits (Schumacher and Krebs, 2010). The subcellular localization depends on which isoform of the "a" subunit is incorporated in the V0 subcomplex. Enzymes incorporating a1 are exclusively located to the TGN, instead enzymes incorporating either a2 or a3 localize

to the tonoplast (Dettmer et al., 2006). The presence of VHA-a1 at the TGN contributes to the acidification of tubules at the *trans*-side of the Golgi, and this might explain Marty's (1978) data. However, conclusive experimental evidence to unequivocally prove that the TGN represents the donor membrane for the biogenesis of the vacuole is lacking up to date.

## **THE ER IS THE MAIN MEMBRANE SOURCE FOR VACUOLE BIOGENESIS**

Immunogold electron microscopy found AVP1-positive provacuoles directly connected to the ER, and immuno-fluorescent *in situ* visualization of sterols showed that ER-export of newly formed membranes can be COPII-independent in *Arabidopsis* roots (Viotti et al., 2013).These data point to the presence of an unknown mechanism in the endoplasmic reticulum that incorporates VHA-a3 and AVP1 to the provacuolar membrane that arises from the ER (**Figure 1**). It is important to mention that this mechanism could not occur anymore in fully mature cells where the growth of the central vacuole and the turnover of tonoplast resident proteins would involve only Golgi- and post-Golgi trafficking.

The hypothesis that the ER was the membrane source for the biogenesis of the vacuole was already proposed decades ago by Matile and Moor (1968) after ultrastructural analysis via freezeetching of *Zea mays* seedlings, and one year later Mesquita (1969) published intriguing electron-micrographs showing connections between vacuoles and the rough ER in *Lupinus albus* roots. During the 1980s, other studies reproposed the ER to be the donor compartment for vacuolar biogenesis (Amelunxen and Heinze, 1984; Hilling and Amelunxen, 1985), nevertheless the absence of immunocytochemistry in these old works did not allow an univocal determination of structures' identity.

The *Arabidopsis* gene *VACUOLESS 1* (*VCL1*) is crucial for vacuole development, since embryo and suspensor cells in the *vcl1* knock-out mutant do not develop vacuoles and the mutant is embryo-lethal (Rojo et al., 2001). Interestingly in this study the authors reported the presence of a high number of auotophagosome-like structures in the embryo cells, which could have been, at least partially, provacuoles. From an ultrastructural point of view provacuoles appear indeed similar to autophagosomes. A recent study nicely depicted autophagosomes at the ultrastructural level for the first time in plants (Zhuang et al., 2013). Provacuoles and autophagosomes seem to be distinct entities, since the former were normally found in *atg2*, *atg5*, and *atg7* knock-out mutants, which lack these key players for the formation of the phagophore (Viotti et al., 2013). Since immunocytochemistry of AVP1 did not label several ring-like structures either in wild-type or in *pat2* seedlings (**Figure 2A**), this data point to the presence of distinct populations of circular doublebilayered membranes in the root meristem of *Arabidopsis*. It is likely that provacuoles and autophagosomes might at some point fuse, both contributing to the development of the vacuole (**Figure 2B**).

Several Golgi- and post-Golgi-trafficking mutants, such as *pat2*, *vps45*, and *amsh3*, display alterations in vacuole morphology (Zouhar et al., 2009; Feraru et al., 2010; Isono et al., 2010), and in all these mutants provacuoles were still present albeit often with aberrant profiles (Viotti et al., 2013). The Golgi and the TGN do

**FIGURE 2 | (A)** Immunogold labeling on ultra-thin sections of highpressure frozen, freeze-substituted, and HM20-embedded *Arabidopsis pat2* root-tip cells shows a provacuole (dashed red line) carrying the V-PPase AVP1. Adjacent to the provacuole, a ring-like structure which is not labeled by the antibody (αAVP1) could be an autophagosome. This

picture shows that at least two distinct populations of circular/semicircular double-bilayered structures existin *Arabidopsis* root-tip cells. Scale bar, 200 nm. **(B)** The development of mature vacuoles is probably the result of fusion events between provacuoles, autophagosomes and vacuolar transport carriers.

not seem to be the donor compartments for provacuole formation, but seem to be required for a rapid and efficient development of the vacuolar lumen, and for the delivery of some, although not all, tonoplast resident proteins. The presence of multilayered provacuoles in the *pat2* and *vps45* mutants suggests that an impaired Golgi- and post-Golgi-trafficking could affect the release of provacuoles from the ER, or the fusion between provacuoles and vacuoles, with the result of a proliferation of membranes that at some point start to curl concentrically forming multilayered compartments.

The incorporation of VHA-a3 and AVP1 already in nascent provacuoles at the ER could be explained through the immediate necessity of acidification of the lumen, which is a key feature of vacuolar activities. The molecular players involved in this putative mechanism of sorting and biogenesis are unknown. The functions of VHA-a2, VHA-a3 and AVP1 seem not to be required, since provacuoles and vacuoles are normally present in *vha-a2/vha-a3* and *avp1* mutants (Viotti and Schumacher, unpublished data). Since α-TIP transport in tobacco leaf cells was shown to be BFA-insensitive (Gomez and Chrispeels, 1993) and can be blocked by SEC12 overexpression (Bottanelli et al., 2011), it cannot be excluded that α-TIP follows the same route of VHA-a3 and AVP1, with SEC12 playing an additional role in this process. While we propose that newly formed lytic vacuoles arise from the ER, a subpopulation might originate from the conversion of protein storage vacuoles (PSVs) when seeds start to germinate (Zheng and Staehelin, 2011). Interestingly, direct transport from the ER to PSVs was reported in pumpkin cotyledons and seeds, where precursors-accumulating vesicles (PAC) arise from the endoplasmic reticulum and are delivered to PSVs (Hara-Nishimura et al., 1998). It is tempting to imagine PACs (diameter of 200–400 nm) being the precursors of PSVs, that slowly acquire their final size and identity via Golgi- and post-Golgi-mediated transport, as we have proposed to happen between provacuoles and lytic vacuoles. The idea that membraneous sheets (as provacuoles look like) arise from the ER (that also has similar structures, the cisternae) appears more reasonable in terms of geometry. One of the elements that could contribute to the formation of provacuoles is the different lipid composition of the nascent membrane compared to that of the ER. Theoretically, the clustering in discrete domains of one or more specific kind of lipids could drive the maturation of an organelle from another one. This could also be the case for the maturation of MVBs, which are enriched in phosphatidylinositol-3-phosphate (PI3P), while the Golgi and TGN mainly have PI4P (Vermeer et al., 2006; Vermeer et al., 2009).

More and more evidence is accumulating for direct ER-tovacuole transport, and those that were supposed to be the "farest" intracellular compartments in plant cells could be, although briefly in time and space, even physically attached.

### **ACKNOWLEDGMENTS**

I apologize with colleagues whose work was not cited or reviewed in depth for space matters. I am grateful to the Department of Plant Physiology at Umeå University, to VIN-NOVA and to the Knut and Alice Wallenberg Foundation for support.

#### **REFERENCES**

**Conflict of Interest Statement:** The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 16 December 2013; accepted: 17 January 2014; published online: 04 February 2014.*

*Citation: Viotti C (2014) ER and vacuoles: never been closer. Front. Plant Sci. 5:20. doi: 10.3389/fpls.2014.00020*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Viotti. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Endoplasmic reticulum-mediated protein quality control in Arabidopsis

## *Yidan Liu and Jianming Li\**

*Department of Molecular, Cellular, and Developmental Biology, University of Michigan, Ann Arbor, MI, USA*

#### *Edited by:*

*Stephen H. Howell, Iowa State University, USA*

#### *Reviewed by:*

*Richard Strasser, University of Natural Resources and Life Sciences, Austria Stephen H. Howell, Iowa State University, USA*

#### *\*Correspondence:*

*Jianming Li, Department of Molecular, Cellular, and Developmental Biology, University of Michigan, 4085 Natural Science Building, 830 North University, Ann Arbor, MI 48109-1048, USA e-mail<EMAIL_ADDRESS>
A correct three-dimensional structure is crucial for the physiological functions of a protein, yet the folding of proteins to acquire native conformation is a fundamentally error-prone process. Eukaryotic organisms have evolved a highly conserved endoplasmic reticulummediated protein quality control (ERQC) mechanism to monitor folding processes of secretory and membrane proteins, allowing export of only correctly folded proteins to their physiological destinations, retaining incompletely/mis-folded ones in the ER for additional folding attempts, marking and removing terminally misfolded ones via a unique multiplestep degradation process known as ER-associated degradation (ERAD). Most of our current knowledge on ERQC and ERAD came from genetic and biochemical investigations in yeast and mammalian cells. Recent studies in the reference plant *Arabidopsis thaliana* uncovered homologous components and similar mechanisms in plants for monitoring protein folding and for retaining, repairing, and removing misfolded proteins. These studies also revealed critical roles of the plant ERQC/ERAD systems in regulating important biochemical/physiological processes, such as abiotic stress tolerance and plant defense. In this review, we discuss our current understanding about the molecular components and biochemical mechanisms of the plant ERQC/ERAD system in comparison to yeast and mammalian systems.

**Keywords: endoplsamic reticulum-associated degradation,** *Arabidopsis***, endoplasmic reticulum-mediated quality control, misfolded glycoproteins, receptor-like kinases**

## **INTRODUCTION**

It is well known that the proper function of a protein strictly depends on its native conformation, but protein folding is a fundamentally error-prone process. The endoplasmic reticulum (ER) is the cellular port of entry for secretory and membrane proteins to enter the secretory pathway and is a folding compartment for proteins to attain their native conformations through interactions with molecular chaperones, sugar-binding lectins, and folding enzymes (Gidalevitz et al., 2013). Misfolded proteins not only lead to functional deficiency but also induce dominant-negative and cellular toxicity effects, and it is thus essential that the ER should possess several highly stringent protein quality control mechanisms to closely monitor the folding process, allowing export of only correctly folded proteins to their final destinations but retaining incompletely/mis-folded proteins for additional rounds of chaperone-assisted folding. A highefficient ER-mediated protein quality control (ERQC) system can also differentiate terminally misfolded proteins from folding intermediates and/or reparable misfolded proteins, stopping the futile folding cycles of the former proteins and eliminating them via a multistep degradation process widely known as ER-associated degradation (ERAD) that involves ubiquitination, retrotranslocation, and cytosolic proteasome (Smith et al., 2011). Our current understanding of the eukaryotic ERQC/ERAD system derived largely from studies in yeast and mammalian cells. However, recent genetic, biochemical, and cell biological studies in the reference plant *Arabidopsis thaliana* and other model

plant species not only identified homologous ERQC/ERAD components but also revealed evolutionarily conserved features as well as unique aspects of the plant ERQC/ERAD mechanisms (Hong and Li, 2012; Huttner and Strasser, 2012; Howell, 2013), especially their connections with the stress tolerance and plant defense pathways.

## *N***-GLYCAN-BASED ER RETENTION MECHANISM**

Many secretory and membrane proteins are co-translationally glycosylated when entering the ER (Aebi, 2013). The so-called Nlinked glycosylation occurs on the asparagine (Asn or N) residues within the Asn-X-Ser/Thr sequons (X indicating any amino acid except proline while Ser/Thr denoting serine/threonine residue) of a nascent polypeptide. This reaction is catalyzed by the enzyme oligosaccharyltransferase (OST), an integral membrane protein complex that transfers a preassembled oligosaccharide precursor Glc3Man9GlcNac2 (Glc, Man and GlcNac denoting glucose, mannose and *N*-acetylglucosamine, respectively) from a membrane-anchored dolichylpyrophosphate (DolPP) carrier to the Asn residue (**Figure 1**; Mohorko et al., 2011). The assembly of Glc3Man9GlcNac2 involves a series of highly specific asparagine-linked glycosylation (ALG) proteins that sequentially add a monosaccharide onto the DolPP linker or a DolPPlinked oligosaccharide precursor (Aebi, 2013; **Figure 1A**). The structure of a N-linked glycan plays an important role in the protein folding and quality control (Aebi et al., 2010). Immediately after transferring of Glc3Man9GlcNac2 to an Asn residue,

**FIGURE 1 | (A)** Stepwise assembly of *N*-glycan precursor on the ER membrane. The assembly of the *N*-glycan precursor starts at the cytosolic face of the ER membrane by adding two GlcNAc and five Man residues to the membrane-anchored Dol-PP linker. The resulting Dol-PP-Man5GlcNAc2 flips over into the ER lumen. Four Man residues are sequentially added to the flipped Dol-PP-Man5GlcNAc2 by three mannosyltransferases, ALG3, ALG9 (known as EBS3 in *Arabidopsis*) and ALG12 (known as EBS4 in *Arabidopsis*) with the ALG9 catalyzing two reactions of adding the terminal α1,2 Man residues on the middle and

the terminal and middle Glc residues are removed sequentially by glucosidase I (GI) and glucosidase II (GII), producing a monoglucosylated *N*-glycan, GlcMan9GlcNac2, which is recognized by the ER chaperone-like lectins, a membrane-anchored calnexin (CNX) and its ER luminal homolog calreticulin (CRT; Caramelo and Parodi, 2008; **Figure 2**). The high-specificity high-affinity binding between GlcMan9GlcNac2 and CNX/CRT is crucial for folding a nascent polypeptide as CNX/CRT can recruit other ER-chaperones and folding enzymes, including protein disulfide isomerases (PDIs) essential for generating inter/intra-molecular disulfide bonds. The removal of the remaining Glc residue by GII releases the nascent glycoprotein from CNX/CRT, thus terminating its folding process (Caramelo and Parodi, 2008). If the protein folds correctly, it will be transported out of the ER to reach its final destination. However, if the protein fails to attain its native conformation, it will be recognized by UDPglucose:glycoprotein glucosyltransferase (UGGT), an ER-resident folding sensor consisting of a large non-conserved N-terminal domain presumably involved in recognizing non-native conformations and a smaller highly conserved C-terminal catalytic domain capable of catalyzing a glucosyltransferase reaction using uridyl diphosphate-glucose (UDP-Glc) as a substrate (D'Alessio et al., 2010). As a result, a single Glc is added back to deglucosylated *N*-glycans of the incompletely/mis-folded protein, permitting its reassociation with CNX/CRT and their associated proteins for another round of assisted folding. The alternate reactions of GII and UGGT drive many cycles of dissociation and reassociation of CNX/CRT with an incompletely/mis-folded glycoprotein [widely known as the CNX/CRT cycle (Hammond et al., 1994)], till the protein attains its native conformation

right branches. Three Glc residues are subsequently added to the right branch, generating the 14-sugar precursor, Dol-PP-Glc3Man9GlcNAc2. Two α1,6 Man residues are marked by brown and salmon color. The Dol-PP linker and different sugar residues are indicated. **(B)** The structure of N-linked Glc3Man9GlcNac2 glycan with three dimannose branches (branch A, B and C). Lower case letters inside sugar residues represent the order of sugar addition. The sugar linkage bonds and enzymes (GI, GII) that remove the three Glc residues are indicated. Figure adapted from Hong et al. (2009).

(**Figure 2**). It is worthy to mention that the budding yeast (*Saccharomyces* cerevisiae), which is widely used for studying the ERAD process, lacks the CNX/CRT-UGGT system due to the presence of a catalytically inactive UGGT homolog (Meaden et al., 1990).

The *Arabidopsis* genome encodes only one UGGT homolog, and its physiological function was inadvertently found in a study for identifying additional signaling proteins of the plant steroid hormones, brassinosteroids (BR; Jin et al., 2007). A genetic screening for extragenic suppressors of an *Arabidopsis* dwarf mutant *brassinosteroid-insensitive 1-9* (*bri1-9*) led to the discovery of *Arabidopsis* UGGT (also known as EBS1 for EMS mutagenized bri1 suppressor 1; Jin et al., 2007). BRI1 is a cell surface-localized leucine-rich-repeat receptor-like-kinase that function as a BR receptor and contains a single transmembrane domain and 14 putative N-glycosylation sites in its N-terminal extracellular domain (Li and Chory, 1997). The mutant bri1- 9, carrying a Ser662-Phe mutation in the BR-binding domain (Noguchi et al., 1999), was found to be retained in the ER by an EBS1/AtUGGT-dependent mechanism and subsequently degraded by a plant ERAD process (Jin et al., 2007; Hong et al., 2009). Loss-of-function mutations in EBS1/AtUGGT compromise such an ER-retention mechanism and allow some bri1-9 proteins to escape from the ER to reach the plasma membrane, resulting in phenotypic suppression of the dwarfism of the *bri1-9* mutant. The same genetic screen also identified CRT3 (Jin et al., 2009), a unique member of the *Arabidopsis* CNX/CRT family consisting of two CNXs and three CRTs, which actually retains bri1-9 via the CRT3-GlcMan9GlcNac2 binding. Both UGGT and CRT3 were also identified from two other independent genetic

CNX/CRT. A correctly folded protein can leave the ER while an incompletely/mis-folded glycoprotein is recognized by UGGT (known as *Arabidopsis*) and Htm1/EDEM. A terminally misfolded glycoprotein with α1,6 Man-exposed glycan is selected to enter the ERAD pathway.

screens aiming to identify key regulators of the plant innate immune response to a bacterial translational elongation factor EF-Tu (Li et al., 2009; Saijo et al., 2009). Interestingly, while lossof-function mutations in AtUGGT/CRT3 led to regaining partial sensitivity to BRs, *atuggt/crt3* mutants were insensitive to elf18, a biologically active epitope of EF-Tu. Further studies showed that both UGGT and CRT3 are absolutely required for the correct folding of EFR (EF-Tu Receptor; Saijo, 2010), a BRI1-like receptor-like kinase that binds elf18/EF-Tu to initiate a plant defense process (Zipfel et al., 2006). The importance of *N*-glycanmediated folding control was further supported by discoveries that loss-of-function mutations in STT3A, a OST subunit, and GII resulted in significant reduction of the EFR protein abundance, presumably caused by incomplete folding and subsequent degradation (Lu et al., 2009; Haweker et al., 2010; von Numers et al., 2010).

In addition to the glycan-dependent ER retention system, the ER is equipped with additional retention systems to prevent export of misfolded proteins, especially those non-glycosylated ones. One system uses the family of ER-localized HSP70 proteins (known as BiPs), which have a N-terminal ATP-binding domain and a C-terminal substrate-binding domain that recognizes and binds to exposed hydrophobic patches of incompletely/mis-folded proteins in an ATP-dependent manner (Buck et al., 2007). The *Arabidopsis* has three BiP homologs, AtBiP1, AtBiP2 and AtBiP3, all of which were known to exhibit higher levels of gene expression under ER stresses (Sung et al., 2001). In *Arabidopsis*, BiPs were shown to bind both bri1-9 and bri1-5, another mutant variant of BRI1 carrying a Cys69Tyr mutation that destroys a disulfide bridge crucial for the structural integrity of the BR receptor, and were thought to contribute for the ER retention of the two mutant BR receptors (Jin et al., 2007; Hong et al., 2008). BiPs and their associated factors ERdj3B (an *Arabidopsis* ER-localized DNAJ homolog) and SDF2 (the *Arabidopsis* homolog of the murine stromal cell-derived factor 2) are also involved in the biogenesis/folding control of EFR (Nekrasov et al., 2009). BiPs were also known to interact with the orphan heavy chain of a murine IgG1 antibody or an assembly defective form of the trimeric vacuolar storage protein phaseolin in transgenic tobacco plants (Pedrazzini et al., 1997; Nuttall et al., 2002). Another glycan-independent ER retention mechanism relies on mixed disulfide bridges between incompletely/mis-folded

proteins with PDIs and related ER-localized oxidoreductases (Reddy et al., 1996; Anelli et al., 2003, 2007). The *Arabidopsis* genome encodes 13 PDI-like proteins (Houston et al., 2005), none of which has been implicated in retaining misfolded proteins. However, a recent study on bri1-5 carrying an orphan cysteine residue (Cys62) suggested involvement of a thiol-mediated retention system in keeping the mutant BR receptor in the ER (Hong et al., 2008). Further biochemical studies are needed to verify this prediction and to identify one or more PDIs that form the predicted mixed disulfide bridge with the orphan Cys62 residue.

## **MARKING OF A TERMINALLY MISFOLDED GLYCOPROTEIN FOR ERAD**

A protein that fails to attain its native conformation within a given time window is eliminated by ERAD (Vembar and Brodsky, 2008). One of the key events in ERQC is to terminate a futile folding cycle and to deliver an irreparable misfolded protein into the ERAD pathway. Although little is known about the marking mechanism for irreparable nonglycosylated ERAD clients, recent studies indicated that removal of the terminal α1,2-Man residue from the C-branch of *N*glycan (**Figure 2**), catalyzed by homologous to mannosidase 1 (Htm1) in yeast and ER-degradation enhancing α-mannosidaselike proteins (EDEMs) in mammals, is required for generating the ERAD signal, an exposed α1,6 Man residue on an Nlinked glycan (Quan et al., 2008; Clerc et al., 2009; **Figure 1B**). The *Arabidopsis* genome encodes at least two homologs of the yeast Htm1/mammalian EDEMs [known as MNS4 and MNS5 (Liebminger et al., 2009)], but their involvement in a plant ERAD process awaits functional investigation. Nevertheless, recent genetic screening for *Arabidopsis* mutants defective in ERAD of bri1-5/bri1-9 and subsequent molecular cloning and biochemical studies indicated that the glycan ERAD signal is well conserved in plants (Hong et al., 2009, 2012). Loss-offunction mutations in either EBS3 and EBS4 (homologs of the yeast/mammal ALG9 and ALG12, respectively) prevent complete assembly of the *N*-glycan precursor (**Figure 1**), resulting in glycosylation of the two ER-retained mutant BR receptor with truncated *N*-glycan lacking the α1,6-Man residue that would function as the ERAD signal and consequential inhibition of ERAD of bri1-5/bri1-9. In contrast, forcing the addition of the missing α1,6 Man residue to Dol-PP-Man6GlcNAc2 by overexpression of EBS4/ALG12 in an *Arabidopsis ebs3/alg9 bri1-9* mutant promoted the ERAD of bri1-9 (**Figure 1A**; Hong et al., 2012). Similarly, the ERAD of bri1-9 was presumably accelerated when its N-linked glycans carried a different exposed α1,6 Man residue (the inner α1,6 Man; Hong et al., 2012) caused by a loss-of-function mutation in ALG3 that adds an α1,3 Man to the inner α1,6-Man (Henquet et al., 2008; Kajiura et al., 2010; **Figure 1A**). The exposed inner α1,6 Man residue was shown to function as an alternative ERAD signal in both yeast and mammalian cells (Clerc et al., 2009; Hosokawa et al., 2009).

## **RECRUITMENT OF ERAD SUBSTRATES**

The *N*-glycan ERAD signal is decoded by one or two ER luminal lectins, osteosarcoma 9 (OS9, also known as Yos9 in yeast) and XTP3-B (Yoshida and Tanaka, 2010; **Figure 3**). Yos9 and its mammalian homologs contain the mannose-6-phosphate receptor homology (MRH) domain that specifically recognizes and binds *N*-glycans with an exposed α1,6 Man residue (Hosokawa et al., 2010). In addition to OS-9/Yos9, selection of an ERAD client requires another ER resident protein, Hrd3 (HMG-CoA reductase degradation 3) in yeast and Sel1L (Suppressor of lin-12-Like) in mammals (Hirsch et al., 2009), a type I transmembrane protein with a large ER luminal domain consisting of multiple copies of the tetratricopeptide repeat motif. It was believed that Hrd3/Sel1L, exhibiting high affinity binding to exposed hydrophobic amino acid residues on misfolded proteins, makes the initial selection of a potential ERAD client, which is subsequently inspected by OS-9/Yos9 for the presence of an *N*-glycan ERAD signal (Denic et al., 2006; Gauss et al., 2006; **Figure 3**). Such a bipartite ERAD signal of a misfolded domain plus an α1,6-Man-exposed *N*-glycan ensures degradation of only terminally misfolded glycoproteins but not folding

intermediates carrying *N*-glycans with no exposed α1,6 Man residue.

The *Arabidopsis* genome has two *Hrd3/Sel1L* homologous genes, *AtSel1A* (also known as *EBS5 or HRD3A* or) and *AtSel1B* (also known as *HRD3B,* an apparent pseudogene) and just one *OS9/Yos9* homolog, *AtOS9* (also known as *EBS6*; Liu et al., 2011; Su et al., 2011, 2012; Huttner et al., 2012). AtSel1A/EBS5 complemented the ERAD-defect of the yeast *hrd3* mutant assayed by ERAD of a mutant variant of carboxypeptidase Y (CPY∗; Su et al., 2011), a commonly used ERAD substrate for many ERAD studies in yeast. By contrast, AtOS9 failed to rescue the defective ERAD of CPY∗ when expressed in a *yos9* yeast strain (Huttner et al., 2012). Interestingly, a chimeric AtOS9-Yos9 protein consisting of the full-length AtOS9 and the Yos9's C-terminal region (amino acids of 277–542) promoted CPY∗ degradation in *yos9* yeast cells (Huttner et al., 2012), suggesting that the MRH domain is interchangeable but the Yos9's C-terminal domain might be crucial for interacting with other components of the yeast ERAD machinery. Loss-of-function mutations in either AtSel1A/EBS5 or AtOS9/EBS6 inhibit ERAD of bri1-5, bri1-9, misfolded EFR (in an *ebs1/uggt* mutant background), and/or the transgenically expressed MLO-1 (Liu et al., 2011; Su et al., 2011, 2012; Huttner et al., 2012), a mutant variant of barley powdery resistance o (MLO) that carries a single amino acid change in the cytoplasmic region and was previously shown to be an ERAD substrate (Muller et al., 2005). As expected, AtSel1A/EBS5 and AtOS9/EBS6 physically interacted with bri1-9 or bri1-5 in a tobacco transient expression system or an *in vitro* pull-down assay (Huttner et al., 2012; Su et al., 2012). Consistent with what was known in yeast and mammalian cells, AtSel1A/EBS5 binds AtOS9/EBS and seems to be required for maintaining the stability of AtOS9/EBS6 (Huttner et al., 2012; Su et al., 2012). These results strongly suggested that the selection mechanism for a terminally misfolded glycoprotein for ERAD is conserved in *Arabidopsis*. It is important to point out that *Arabidopsis* mutants of AtSel1A/EBS6 or AtOS9/EBS6 are hypersensitive to NaClinduced salt stress, suggesting a relationship between a cellular stress response and an environmental stress pathway (Liu et al., 2011; Huttner et al., 2012). It is quite possible that environmental stresses lead to decreased folding efficiency and increased accumulation of misfolded proteins in the ER, which require a highly efficient ERAD system for their removal to maintain ER homeostasis.

## **UBIQUITINATION OF CHOSEN ERAD CLIENTS**

Hrd3/Sel1L and Yos9/OS9 not only select irreparable misfolded glycoproteins but also bring the chosen ERAD substrates to the membrane-anchored ERAD complexes responsible for ubiquitination and retrotranslocation. The central component of these ERAD complexes is a polytopic membrane protein with a RING finger-type ubiquitin ligase (E3) activity exposed to the cytosolic surface of the ER membrane, which not only ubiquitinates ERAD substrates but also connects to various ER luminal/cytosolic adapters (Hirsch et al., 2009). Yeast contains at least two distinct E3 ligases, 6 transmembrane-spanning Hrd1 (HMG-CoA reductase degradation) and 14-transmembrane-spanning Doa10 (Degradation of alpha2), that ubiquitinate three different types of ERAD substrates differing in the location of folding lesions: ERADL (lesion in the ER luminal area), ERADM (lesion in the transmembrane segment), and ERADC (lesion in the cytosolic domain; Vashist and Ng, 2004; Carvalho et al., 2006). The Hrd1 complex ubiquitinates ERADL/M substrates while the Doa10 complex deals with ERADC clients. Mammals have at least 9 membrane-bound ERAD E3 ligases (Olzmann et al., 2013), including two Hrd1 homologs (HRD1 and gp78), one Doa10 homolog (TEB4), and several other RING-type E3 ligases such as RING membrane-anchor 1 (RMA1; Younger et al., 2006), whose founding member was initially discovered in*Arabidopsis*(Matsuda and Nakano, 1998).

The *Arabidopsis* genome encodes two Hrd1 homologs (AtHrd1A and AtHrd1B; Su et al., 2011; Huttner and Strasser, 2012), at least two Doa10 homologs (Doa10A/At4g34100 and Doa10B/At4g32670; Liu et al., 2011), and three homologs of RMA1, AtRMA1-AtRMA3 that were shown to be localized to the ER and exhibit *in vitro* E3 ubiquitin ligase activity (Son et al., 2009; **Table 1**), but it remains unclear if plants use distinct E3 ligases to removal different classes of ERAD substrates. Loss-offunction mutations in AtHrd1A or AtHrd1B had no detectable effect on bri1-5/bri1-9 degradation, but simultaneous elimination of the two Hrd1 homologs inhibited degradation of the two mutant BR receptors, indicating that AtHrd1A and AtHrd1B function redundantly in a plant ERAD pathway (Su et al., 2011). By contrast, the role of the two Doa10 homologs in the plant ERAD pathway remains unknown. Two recent genetic studies revealed important regulator roles of Doa10A (also known as SUD1 for SUPPRESSOR OF DRY2 DEFECTS1 or CER9 for ECERIFERUM9) in the cuticle lipid biosynthesis and in controlling the activity but not the protein level of an *Arabidopsis* HMG-CoA reductase (Lu et al., 2012; Doblas et al., 2013). Further studies are needed to determine if the *Arabidopsis* Doa10A is indeed involved in an ERAD pathway that regulates the protein abundance of key regulatory factors or metabolic enzymes involved in the cuticle lipid biosynthesis. Unlike yeast but similar to mammals, plants have additional membrane-anchored RING-type E3 ligases for ERAD. For example, the three *Arabidopsis* RMA1 homologs (Rma1H1) and a hot pepper (*Capsicum annuum*) Rma1H1 are involved in the degradation of a cell surface water channel to regulate its plasma membrane level (Lee et al., 2009). A recent study also suggested that a legume (*Medicago truncatula*) homolog of RMA1 seems to play a role in the regulation of biosynthesis of plant defense compounds, triterpene saponins that share the same biosynthetic precursors with sterols, through regulated degradation of HMG-CoA reductase (Pollier et al., 2013).

In a typical ubiquitination reaction, ubiquitin is attached to a substrate through a three-step process consisting of activation, conjugation, and ligation catalyzed by an ubiquitin-activating enzyme (E1), ubiquitin-conjugating enzyme (E2), and E3 (Pickart, 2004). In yeast, the Hrd1/Doa10 E3 ligases work together with a membrane-anchored E2 (Ubc6) and two cytosolic E2s (Ubc1 and Ubc7) that are recruited to the ER membrane by an ER anchor protein Cue1 (Hirsch et al., 2009), which also activates both E2 and E3 (Bagola et al., 2013; Metzger et al., 2013). *Arabidopsis* has a total of 37 E2 enzymes (Kraft et al., 2005), including

#### **Table 1 | A list of known/predicted components of the** *Arabidopsis* **ERAD system.**

one potential Ubc1 homolog (UBC27), three putative homologs of Ubc7 known to be the cognate E2 for Hrd1 (UBC7 UBC13, and UBC14), and three likely homologs of Ubc6 associated mainly with Doa10 (UBC32, UBC33, and UBC34 each having a predicted transmembrane domain at their C-termini; Liu et al., 2011), but our understanding of the roles of these potential ERADparticipating E2s in the plant ERAD process is extremely limited. One of the *Arabidopsis* Ubc6-like E2 gene, *UBC32*, was recently found to be induced by salt, drought, and ER stress (Cui et al., 2012). Interestingly, the *Arabidopsis ubc32* mutant seedlings are more tolerant whereas *UBC32*-overexpressing transgenic *Arabidopsis* lines are more sensitive to salt and ER stress (Cui et al., 2012). The observed salt tolerance of the *ubc32* mutant is in contrast to the reduced salt tolerance of other known *Arabidopsis* ERAD mutants, including *ebs5/hrd3a*, *ebs6/os9*, and *hrd1ahrd1b* (Liu et al., 2011; Huttner and Strasser, 2012; Huttner et al., 2012). This discrepancy might be explained by different ERAD substrates being degraded by different E3 ligase complexes: ERADL/M substrates by AtHrd1A/AtHrd1B in association with cytosolic E2s and ERADC substrates by Doa10 using membrane-anchored E2s. Indeed, UBC32 was found to interact with *Arabidopsis* Doa10B and to stimulate the ubiquitination and degradation of a known ERAD substrate MLO-12, another variant of MLO carrying a single amino acid change in its cytosolic domain (Muller et al., 2005), in a tobacco leaf transient expression experiment (Cui et al., 2012). However, the tobacco result was quite different from the results obtained with the yeast MLO experiment showing that the ERAD of MLO-12 plus two other mutant MLOs (all carrying a cytosolic mutation) used the Ubc7-Hrd1 pathway but was unaffected by either *ubc6* or *doa10* deletion in yeast (Muller et al., 2005). Such inconsistency might be simply caused by heterologous expression of ERADC substrates in two different eukaryotic systems. Nevertheless, UBC32 was implicated in the Hrd1-mediated degradation of bri1-9 (a presumed ERADL substrate) as the *ubc32* mutation partially inhibited the degradation of the mutant BR receptor and weakly suppressed the corresponding dwarf phenotype (Cui et al., 2012). The partial inhibition could be attributed to a redundant role of UBC32 with its two close homologs or the potential *Arabidopsis* homologs of Ubc1/Ubc7. However, blast searches failed to find a single homolog of the yeast *Cue1* gene from published sequences of plant genomes and expressed sequence tags (our unpublished results), suggesting that plant ERAD processes might exclusively rely on ER-anchored membrane E2s. Alternatively, plants could recruit cytosolic E2s to the membrane-anchored E3 complexes via yet unknown recruiting factors that share no sequence homology but are functionally similar to Cue1.

The ubiquitination of ERAD substrates, especially those lacking *N*-glycan degradation signals, by the Hrd1 complex requires two additional adapters: U1-Snp1 associating-1 (Usa1; HERP in mammals), an ER membrane protein containing a ubiquitin-like (UBL) motif near it N-terminus and two predicted transmembrane domains in the middle, and Der1 (degradation in the ER; Derlins for Der1-like proteins in mammals), another integral ER membrane protein with four transmembrane segments (Kostova et al., 2007). Usa1 is thought to regulate the stability and/or oligomerization of Hrd1 and to recruit Der1 to the Hrd1

complex (Carvalho et al., 2006, 2010; Horn et al., 2009; Carroll and Hampton, 2010), while Der1 is believed to function either as a receptor for soluble non-glycosylated ERAD substrates or a potential retrotranslocation channel (Lilley and Ploegh, 2004; Ye et al., 2004; Kanehara et al., 2010). The *Arabidopsis* contains no homolog of Usa1/HERP1 but its genome encodes three Der1 homologs whose functional involvement in a plant ERAD pathway awaits detailed genetic and biochemical investigations (Kirst et al., 2005). An earlier study showed that at least two maize Der1 homologs could complement the yeast *der1* mutant, suggesting a potential role for a plant Der1 homolog in an ERAD pathway; however, there is no genetic evidence for proving the hypothesis (Kirst et al., 2005).

## **RETROTRANSLOCATION OF ERAD SUBSTRATES**

Because the catalytic domains of the ERAD-participating E2s and E3s are on the cytosolic surface of the ER membrane, ERAD substrates need to undergo retrotranslocation for ubiquitination and to access the cytosolic proteasome system for their degradation. However, the molecular nature of this retrotranslocon remains controversial (Hampton and Sommer, 2012). It was previously thought that the Sec61 translocon, which imports nascent polypeptides into the ER lumen during protein biosynthesis, is responsible for retrotranslocation ERAD substrates through the ER membrane (Pilon et al., 1997; Plemper et al., 1997). Other studies suggested that the yeast Der1 and its mammalian orthologs Derlins are the suspected retrotranslocon (Lilley and Ploegh, 2004; Ye et al., 2004). A recent study, however, showed that the E3 ligase Hrd1 itself could serve as the retrotranslocation channel for ERADL substrates (Carvalho et al., 2010). It is quite possible that all three proteins are capable of retrotranslocation different ERAD substrates involving different adapter proteins.

Compared to the knowledge gained from the yeast and mammalian studies, we know almost nothing about the retrotranslocation step of a plant ERAD pathway. Several earlier studies did suggest the existence of a retrotranslocon in plant cells to move ERAD substrates into the cytosol. A confocal microscopic analysis of subcellular localization of a fusion protein between green fluorescent protein (GFP) with the P-domain of a maize CRT in tobacco leaf protoplasts suggested a retrotransport route from the ER to the cytosol (Brandizzi et al., 2003). In addition, a series of studies revealed that the A chain (known as RTA) of a ribosome-inactivating toxin, ricin that is normally produced as a dimeric protein of RTA covalently linked to a galactose-binding B chain via a single intramolecular disulfide bond and stored in the central vacuole of the endosperm cells of castor bean (*Ricinus communis*), was detected to be deglycosylated and eventual degraded in the cytosol when expressed alone in tobacco leaf protoplasts (Di Cola et al., 2001, 2005; Marshall et al., 2008). It is important to mention that ricin and a few other plant toxins were known to exploit the ERAD pathway to reach their cytosolic targets after being internalized by mammalian cells and retrograde-transported from the cell surface to the ER (Lord et al., 2003). In both yeast and mammalian systems, retrotranslocation of ERAD substrates was driven by ubiquitination (Bagola et al., 2011); however, a recent RTA study using plant protoplasts showed that retrotranslocation is

independent of ubiquitination as the lysine-lacking (hence nonubiquitinated) variant of RTA could still be retrotranslocated from the ER into the cytosol (Di Cola et al., 2005), suggesting that the ubiquitination-retrotranslocation coupling might be substrate-dependent.

## **SUBSTRATE EXTRACTION, PROCESSING, AND DELIVERY TO THE PROTEASOME**

Without regard to the identity of the actual retrotranslocons, ubiquitinated ERAD clients are extracted from the ER lumen (ERADL substrates) or ER membrane (ERAD*M*/*C* substrates) by a trimeric complex consisting of a homohexameric Cdc48 (p97 or valosincontaining protein in mammals), an AAA-type ATPase and its two substrate-recruiting factors Ufd1 and Npl4 (each having a ubiquitin-binding domain; Wolf and Stolz, 2012). The (CDC48)6- Ufd-Npl4 complex itself is recruited to the Hrd1/Doa10 E3 complexes by Ubx2 (VIMP for p97/VCP-interacting membrane protein in mammals), one of the 7 ubiquitin regulatory X (UBX) domain-containing proteins in yeast (13 UBX proteins in mammals; Neuber et al., 2005; Schuberth and Buchberger, 2005, 2008). The current working model posits that extracted ERAD substrates are further processed through antagonistic interactions between an U-box-containing E4 multiubiquitination enzyme Ufd2 and a WD40 repeat-containing protein Ufd3 with unknown enzyme activity plus a deubiquitylating enzyme Otu1, and/or through deglycosylation by the cytoplasmic peptide:*N*-glycanase (PNGase) Png1 (Raasi and Wolf, 2007). The processed ERAD substrates were subsequently delivered to the cytosolic proteasome by Cdc48 in association with two ubiquitin receptors Rad23 and Dsk2, each containing a UBL domain that interacts directly with the cytosolic proteasome and a polyubiquitininteracting ubiquitin-associated (UBA) domain (Raasi and Wolf, 2007).

The *Arabidopsis* genome encodes three Cdc48 homologs, AtCDC48A, AtCDC48B, and AtCDC48C (Rancour et al., 2002). AtCDC48A was able to complement a yeast *cdc48* mutant (Feiler et al., 1995) and was shown to play a role in the ERAD of a mutant form of MLO and a mutant variant of the *Arabidopsis* vacuolar carboxypeptidase carrying the same Gly-Arg mutation as the yeast CPY∗ and in the retrotranslocation of RTA and the orphan subunit (RCA A) of another castor bean toxin agglutinin in plant cells (Muller et al., 2005; Marshall et al., 2008; Yamamoto et al., 2010). AtCDC48A is likely to be recruited to the ER membrane by UBX-containing proteins as the *Arabidopsis* genome encodes a total of 15 UBX-containing proteins (known as AtPUXs; **Table 1**), some of which were shown to interact with AtCDC48A (Rancour et al., 2004; Park et al., 2007). It remains to be determined which of the 15 AtPUX proteins are actually involved in recruiting an AtCDC48 to the ER membrane-anchored E3 ligase complexes and play a role in degrading known plant ERAD substrates. Our BLAST searches using the known ERAD components of yeast and mammals as query identified multiple homologs of the Ufd1, Ufd2, Ufd3, Npl4, Rad23, Dsk2 but only a single PNGase homolog in *Arabidopsis* (**Table 1**). The functional involvement of these potential ERAD components in an *Arabidopsis* ERAD process remains unknown except AtPNG1, which was recently shown to contain the suspected PNGase activity and could stimulate the degradation

of two mutant variants of RTA in an *N*-glycan-dependent manner in yeast cells (Diepold et al., 2007; Masahara-Negishi et al., 2012).

### **CONCLUSION AND CHALLENGES**

Despite rapid progress in recent years for identifying molecular components of plant ERQC/ERAD systems and studying their biochemical functions, our understanding of the plant ERQC/ERAD processes remains rather limited, especially about the later stages of the ERAD pathway, such as retrotranslocation, processing of polyubiquitin chains, and delivery (to cytosolic proteasome) of the known plant ERAD substrates. While forward genetic screens in*Arabidopsis* identified the GII-UGGT-mediated CNX/CRT cycle in retaining incomplete/mis-folded glycoproteins and ERAD components that function inside the ER lumen to promote the degradation of the two mutant BR receptors, reverse genetic approaches using T-DNA insertional mutants or RNAi-mediated knockdown of candidate ERAD genes listed in **Table 1** will certainly provide additional knowledge on the plant ERAD mechanisms. Transgenic *Arabidopsis* lines expressing carefully engineered substrates of glycosylated/non-glycosylated ERADC/ERADM coupled with forward genetic screens and reverse genetic studies will reveal if *Arabidopsis* has several distinct ERAD subpathways using different E3 ligases and adapter proteins that recruits distinct ERAD clients. Similarly, genetic screens for enhancers/suppressors of the *Arabidopsis* wax mutant *cer9* [defective in Doa10A (Lu et al., 2012)] or *drought hypersensitive 2* mutant [that led to independent discovery of Doa10A (Doblas et al., 2013)] could uncover additional ERAD components, reveal unique features of the plant ERAD processes, and a better understanding of the regulatory function of the plant ERAD system in biosynthetic processes. Proteomic studies with the existing *Arabidopsis* mutants of the ERAD E3 ligases could lead to the discovery of additional biochemical pathways and/or physiological processes regulated by the plant ERAD machinery. However, the biggest challenges for the plant ERQC/ERAD research is whether the forward genetic approach in *Arabidopsis* could identify novel ERQC/ERAD components that haven't been discovered in other eukaryotic systems and if the combination of the *Arabidopsis* genetics with cutting-edge biochemical studies in *Arabidopsis* and transient expression systems could reveal novel biochemical functions of known or predicted ERAD components and provide satisfactory answers to some of the outstanding questions of the general ERQC/ERAD research field.

## **REFERENCES**

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 15 December 2013; paper pending published: 14 March 2014; accepted: 07 April 2014; published online: 30 April 2014.*

*Citation: Liu Y and Li J (2014) Endoplasmic reticulum-mediated protein quality control in Arabidopsis. Front. Plant Sci. 5:162. doi: 10.3389/fpls.2014.00162*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Liu and Li. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Conserved and plant-unique strategies for overcoming endoplasmic reticulum stress

## *Cristina Ruberti 1,2 and Federica Brandizzi 1,2 \**

*1 Plant Research Laboratory, Department of Energy, Michigan State University, East Lansing, MI, USA 2 Department of Plant Biology, Michigan State University, East Lansing, MI, USA*

#### *Edited by:*

*Lorenzo Frigerio, University of Warwick, UK*

#### *Reviewed by:*

*Aldo Ceriotti, Consiglio Nazionale delle Ricerche, Italy Ben Matthew Abell, Sheffield Hallam University, UK*

#### *\*Correspondence:*

*Federica Brandizzi, Plant Research Laboratory, Department of Energy, Michigan State University, 612 Wilson Road, East Lansing, MI 48824, USA e-mail<EMAIL_ADDRESS>