<figure><img src="image_25.png" /><figcaption>**Figure 6.** Immunolocalisation of extensins in Brachypodium embryos. (a–a") coleoptile with first leaf; (b–b") second leaf and (c–c") mesocotyl. All with JIM12. Abbreviations: CF—calcofluor, Col—coleoptile, FL—first leaf, Mes—mesocotyl, and SL—second leaf. Bars: 10 µm.</figcaption></figure>

Pectins are high-molecular carbohydrates with D-galacturonic acid as their main structural component of that are found in the tissues of terrestrial plants and in some algae. In some plant tissues, for example in the white part of a citrus peel, the pectin content may reach up to 30% of dry weight, while, in others, it does not exceed a fraction of a percent [31,32]. Grasses, which contain a type II cell wall, are generally pectin poor [33]. The immunolocalisation of the homogalacturonan epitopes that recognised de-methyl-esterified pectin LM19 in Brachypodium were found on the outer surface of the seed coat (Figure 7a–a"). Moreover, LM19 was localised in different embryo parts, for example, in the cell wall of the mesocotyl and radicula, respectively (Figure 7b–b",c–c"). The LM13 epitope could be a negative marker for the epidermal cells of the first leaf of a Brachypodium embryo. These antibodies gave signals in the cell walls in almost all parts of the embryo but were absent in the epidermal cells of the first leaf (Figure 7d–d"; red arrows). A different location of LM13 was observed in the coleorhiza and radicula cells. In the coleorhiza, this epitope was found in the cell walls, while in the radicula, it had an internal cell localisation (Figure 7e–e"). In Arabidopsis, the LM13 epitope is detectable in the epidermal cell walls of the younger, more flexible regions of the inflorescence stems and is almost absent in the base of the stem [34,35]. These authors demonstrated that the contribution of the arabinan structures to the cell wall mechanical properties influences the responsiveness to the mechanical stress. Therefore, the specific lack of the LM13 signals in the epidermal cells of the first leaf of Brachypodium embryo may be related to the absence of any mechanical stress in this part of an embryo.

The fluorescence signals of LM6 against arabinans were found in both the cell wall and the internal cell compartment of the mesocotyl and radicula (Figure 8a–a",b–b"). Furthermore, LM6 was localised on the external surface of the root cap but was absent in all other cells (Figure 8c–c"). As was demonstrated in Arabidopsis, the arabinans accumulated in developing and mature embryos but disappeared during germination and seedling establishment [36]. The early stages of Arabidopsis seed development showed a punctate distribution of this epitope compared to mature seeds, where LM6 labelling was very intense and distributed in the cell walls in the entire embryo. It was suggested that changes in the number and localisation of the arabinans might be connected with seedling germination.
In barley, at the beginning of this process, the cell walls in the coleoptile have comparably high levels of arabinose-rich pectic polysaccharides, which decrease strongly during germination [37]. The abundance of the arabinans in the cell walls of seeds is very often a characteristic feature in a wide range of species such as almond (*Prunus dulcis*) [38], rapeseed (*Brassica napus*) [39], and honey locust (*Gleditsia triacanthos*) [40].

<figure><img src="image_26.png" /><figcaption>**Figure 7.** Immunolocalisation of pectins in Brachypodium embryos. (a—a",b—b",c—c") Coleoptile with the seed coat, mesocotyl, and radicula are with LM19, respectively; Both (d—d") coleoptile with first lead and (e—e") coelorhiza with radicula with LM13. Arrows point to *FL* epidermal cells (d—d"). Abbreviations: CF—calcofluor, Col—coleoptile, Cr—coelorhiza, FL—first leaf, Mes—mesocotyl, Rad—radicula, and TSC—the seed coat. Bars: (a—a",b—b",c—c") 10 µm; (d—d",e—e") 20 µm.</figcaption></figure>
<figure><img src="image_27.png" /><figcaption>**Figure 8.** Immunolocalisation of pectins in Brachypodium embryos. (a–a") Mesocotyl; (b–b") radicula and (c–c") root cap. All with LM6. Abbreviations: CF—calcofluor, Rad—radicula, RC—root cap, and Mes—mesocotyl. Bars: (a–a", b–b") 10 µm; (c–c") 20 µm.</figcaption></figure>

Hemicelluloses differ from cellulose in the composition of their monomers and their branched
arrangement in molecules [7,41,42]. They are one of the components of the plastic matrix and impart
additional strength to the cell wall, but do not hinder its growth [5]. Further, due to the ease of
hydrolysis, they can also serve as reserve substances. In the Brachypodium embryo, the signals
of the hemicellulose that recognise antibodies were localised in a different pattern. The signals of
the LM21 antibodies were visualised in the internal cell compartments of all of the embryo parts,
for example in the coleoptile and radicula (Figure 9a–a”,b–b”), while the LM25 antibodies were
detected only in the radicula and root cap (Figure 9c–c”,d–d”,e–e”). Since it is difficult to determine
whether the immunostaining is limited to only the cell wall, or whether it is also present outside of it
using light microscopy, a TEM analysis was performed. Since this analysis was performed without
immunostaining, it only showed the exact thickness of the wall (Figure S1). Thus, a calculation
of the wall thickness (Figure 9c”) revealed that it was about 2 µm and that the thickness of the
fluorescence “line” that was visible on Figure 9c’–c” was about 14.2 µm. This observation indicates
that the hemicellulose epitopes were also outside the radicula protodermis, which can be explained by
taking into account the development of the embryo, in which, during the final stages of development,
detachment of the protective tissues such as the coleorhiza and coleoptile occur [43]. The localisation
of some cell wall epitopes described in this work in the cellular compartments, indicate their extensive
synthesis in the organelles, such as endoplasmic reticulum and Golgi apparatus dictosomes, which are
involved in this process [28,44,45]. It is worth noting that the same antibodies (LM21 and LM25) were
localised in a different way in the embryogenic callus of Brachypodium [46]. As was demonstrated
there, the LM21 antibodies only gave signals in the cell junctions of the embryogenic masses and in the
parenchymatous cells. The signals of LM25 were found in every cell of both the embryogenic masses
and parenchymatous cells.
<figure><img src="image_28.png" /><figcaption>**Figure 9.** Immunolocalisation of hemicelluloses in Brachypodium embryos. (a–a") coleoptile; (b–b") radicula with LM21, respectively; (c–c",d–d",e–e") the entire embryo, root cap with coleorhiza and radicula with coleorhiza with LM25, respectively. Arrows demonstrate insets: (c¹,c²) histological sections of embryo, scale bars are 50 and 2 µm, respectively. Abbreviations: CF—calcofluor, Col—coleoptile, Cr—coleorhiza, Rad—radicula, and RC—root cap. Bars: (a–a",d–d",e–e") 10 µm; (b–b") 20 µm; (c–c") 100 µm.</figcaption></figure>

In our current study, such a high deposition of xyloglucans seems to be important as a stock of elements for embryo development. Xyloglucans are known to differentially express throughout the embryo in Arabidopsis [47]. An analysis of Arabidopsis *xyl1* mutant phenotypes, which are associated with modifications of the composition of the endosperm cell wall demonstrated altered germination characteristics, such as shorter and thicker siliques, a reduced dormancy, and an increased tolerance to germination inhibitors and thermoinhibition. This study highlighted the role of xyloglucans during seed development and germination. Xyloglucans are likely to be very important components for cell adhesion and cell separation in fruit seed coat parenchyma [48]. Research on the tomato mutant *Cnr* (colourless non-ripening), which has a pleiotropic dominant mutation with reduced cell-to-cell adhesion, demonstrated that xyloglucan polymers (galactan and mannan) that are distributed more
widely can be targeted to effect cell release from a parenchyma system. It was demonstrated that a monocotyledon palm *Euterpe oleraceae* contains a highly methoxylated homogalacturonan together with small amounts of a mannoglucan. Furthermore, a type II arabinogalactan was structurally characterised [49]. The experiments in non-lignified cell walls in different monocotyledon species, using the monoclonal antibody CCRC-M1 against fucosylated xyloglucans, demonstrated the presence of this epitope in non-lignified walls. A similar pattern was also found in the palm *Phoenix canariensis*. In Zingiberales, Commelinales, and Poales this epitope was found in the phloem walls, stomatal guard, subsidiary cells, and raphide idioblasts. However, it was not found in *Tradescantia virginiana* (Commelinaceae, Commelinales), or *Zea mays* (Poaceae, Poales). On the other hand, this epitope was observed in the phloem walls of two other Poaceae species, *Lolium multiflorum* and *L. perenne*. The authors highlighted the lack of knowledge about the functions of xyloglucans in monocotyledonous plants [50]. Moreover, work on Brachypodium revealed different proteome profiles that related to the different phases of grain development, and provided information of the reorganization of cell wall constituents that occurs during these stages [9,51]. Such results indicate that changes in cell wall chemical composition are important factor involved in its assembly and remodelling on different stages of development, plant tissues, and organs, both during embryonic and post-embryonic stages of development.

**3. Materials and Methods**

**3.1. Plant Material and Histological Procedures**

Approximately 15 mature embryos of Brachypodium (29 days after fertilization), reference genotype Bd21, were isolated from seeds that were collected from plants growing in pots with soil mixed with vermiculite (3:1) in a greenhouse. Plants were grown at 20 ± 1 °C, under a 16/8 h light/dark photoperiod. To ensure synchronised flowering, approximately two-week-old plants were subjected to vernalisation for two weeks at 4 °C. The embryos were fixed in a mixture of 4% (*w*/*v*) paraformaldehyde (PFA) and 1% (*v*/*v*) glutaraldehyde (GA) in phosphate-buffered saline (PBS, pH 7.0) overnight at 4 °C. Then, they were rinsed with PBS (3 × 15 min), dehydrated in an ascending ethanol series (10%, 30%, 50%, 70%, 90%, and 100%; 2 × 30 min in each) and gradually embedded in LR White resin [52]. The material was cut into 1.5-µm thick sections using an EMUC6 ultramicrotome (Leica Microsystems, Wetzlar, Germany). Sections were collected on microscopic slides covered with poly-L-lysine. For general histology, they were stained with 0.05% (aqueous solution) Toluidine Blue O (Sigma-Aldrich, St. Louis, MO, USA) for 5 min.

**3.2. Immunocytochemistry**

Sections were incubated in a blocking buffer containing 2% (*v*/*v*) foetal calf serum (FCS) and 2% (*w*/*v*) bovine serum albumin (BSA) in PBS (pH 7.2) for 30 min at room temperature (RT). Next, they were incubated with specific primary monoclonal antibodies (Table 1), diluted at a 1:20 ratio in a blocking buffer (RT, minimum 1.5 h), rinsed with the blocking buffer 3 × 10 min and then incubated at RT for at least 1.5 h with the secondary antibody (Alexa Fluor 488 goat anti-rat IgG, Jackson Immuno Research Laboratories, West Grove, PA, USA) diluted 1:100 in the blocking buffer as above. After washing with the blocking buffer and PBS (3 × 10 min each), the sections were stained with 0.01% calcofluor (Sigma-Aldrich, St. Louis, MO, USA) in PBS for 5 min; then slides were thoroughly rinsed with PBS and sterile distilled water (3 × 10 min each). The drained slides were mounted in a Fluoromount (Sigma-Aldrich) antifade medium. Negative controls were performed for each antibody that was used by omitting the primary antibodies. All of the images were taken using an Axio Imager Z2 (Zeiss, Oberkochen, Germany) epifluorescent microscope equipped with an AxioCam Mrm monochromatic camera (Zeiss) with the narrow-band filters for AlexaFluor 488 and DAPI. The negative control for all of the antibodies revealed the absence of any signals that were specific for these antibodies and was representative for all of the other antibodies that were used in this study (Figure S2).
<table><caption>**Table 1.** The antibodies used for immunocytochemistry, the epitopes they recognise, and relevant references.</caption><thead><tr><th>Antibody</th><th>Epitope</th><th>References</th></tr></thead><tbody><tr><td>MAC207</td><td><i>Arabinogalactan proteins (AGPs)</i><br>β-GlcA1-&gt;3αGalA1-&gt;2Rha</td><td>[53–56]</td></tr><tr><td>JIM8</td><td>Arabinogalactan</td><td>[57]</td></tr><tr><td>JIM13</td><td>βGlcA1-&gt;3αGalA1-&gt;2Rha</td><td>[55,56,58]</td></tr><tr><td>JIM16</td><td>AGP glycan</td><td>[55,56,58]</td></tr><tr><td>LM2</td><td>β-linked GlcA</td><td>[56,59]</td></tr><tr><td>LM1</td><td><i>Extensins</i><br>Extensin</td><td>[60]</td></tr><tr><td>JIM11</td><td>Extensin</td><td>[55,61]</td></tr><tr><td>JIM12</td><td>Extensin</td><td>[61]</td></tr><tr><td>LM19</td><td><i>Pectins</i><br>α-GalA(1-4)α-GalA(1-4)α-GalA</td><td>[35]</td></tr><tr><td>LM13</td><td>α-1,5-arabinan</td><td>[62]</td></tr><tr><td>LM16</td><td>Processed arabinan—rhamnogalacturonan (RG)-I domain</td><td>[35]</td></tr><tr><td>LM6</td><td>αAra1-5αAra1-5αAra1-5αAra1-5Ara</td><td>[63]</td></tr><tr><td>LM21</td><td><i>Hemicelluloses</i><br>β-linked mannans polysaccharides of plant cell walls</td><td>[64]</td></tr><tr><td>LM25</td><td>Xyloglucan</td><td>[65]</td></tr></tbody></table>

### 3.3. TEM

The samples for TEM were fixed in 2.5% glutaraldehyde in a 0.1 M sodium phosphate buffer (pH 7.4) at 4 °C for 24 h. The embryos were postfixed in 1% OsO₄ (osmium tetroxide) in a 0.1 M sodium phosphate buffer for two hours at 4 °C, rinsed in the same buffer, dehydrated in an ascending series of acetone, and gradually embedded in Epon 812 (Fullam, Latham, NY, USA) [66]. Semithin sections were stained with toluidine blue and examined with a bright-field microscope. Ultrathin (70 nm) sections were cut on a Leica ultracut UCT ultramicrotome and collected on copper grids (300 mesh, Electron Microscopy Science, Hatfield, PA, USA). Sections were stained with uranyl acetate and lead citrate and examined with a Hitachi H500 TEM (Hitachi, Tokyo, Japan) at 75 kV.

### 4. Conclusions

In this work we traced the distribution of various cell wall epitopes in the Brachypodium zygotic embryo in mature stages of development. We have demonstrated that:

1. Among analysed AGP epitopes JIM13, JIM8, and LM2 were localised in cell walls of the embryo surface tissues, as well as some of them in the seed coat; and MAC207 and JIM16 were detected in cytoplasmic compartments.

2. Extensins are localised in the outer periclinal walls of embryo surface tissues. Our results suggest that in Brachypodium, the antibody LM13 can be used as a negative marker of epidermal cells of the embryonic leaf and that the LM25 antibody can be used as the positive marker of embryonic root epidermis cells, as well as the root cap in Brachypodium. These results demonstrate that the distribution of analysed cell wall epitopes in mature zygotic embryos of Brachypodium is tissue- and organ specific.

Without a doubt, the precise dissection of all functions of the cell wall in Brachypodium embryos requires the use of specific mutants, whose availability is limited. However, this problem should soon be overcome by the rapidly-developing techniques of site-directed mutagenesis, such as CRISPR/Cas9 that are already available for Brachypodium [67].

**Supplementary Materials:** The following are available online at www.mdpi.com/1422-0067/19/3/725/s1.

**Acknowledgments:** This work was supported by the National Science Centre, Poland (grant no. DEC-2014/14/M/NZ2/00519).
**Author Contributions:** Alexander Betekhtin, Anna Milewska-Hendel, Joanna Lusinska, and Lukasz Chajec performed the experiments; Alexander Betekhtin, Ewa Kurczynska, and Robert Hasterok analysed the data; the paper was written under the direction and supervision of Ewa Kurczynska and Robert Hasterok; and Alexander Betekhtin was responsible for writing this work. All authors read and approved the final manuscript.

**Conflicts of Interest:** The authors declare no conflict of interest.

**Abbreviations**

<table><tr><td>AGP</td><td>Arabinogalactan protein</td></tr><tr><td>BSA</td><td>Bovine serum albumin</td></tr><tr><td>CRISPR</td><td>Clustered regularly-interspaced short palindromic repeats</td></tr><tr><td>FCS</td><td>Foetal calf serum</td></tr><tr><td>GA</td><td>Glutaraldehyde</td></tr><tr><td>PBS</td><td>Phosphate-buffered saline</td></tr><tr><td>PFA</td><td>Paraformaldehyde</td></tr><tr><td>RT</td><td>Room temperature</td></tr><tr><td>TEM</td><td>Transmission electron microscope</td></tr></table>

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Review

# Plasmodesmata-Related Structural and Functional Proteins: The Long Sought-After Secrets of a Cytoplasmic Channel in Plant Cell Walls

Xiao Han <fnref n="1" />,†, Li-Jun Huang <fnref n="2" />,†, Dan Feng <fnref n="3" />,†, Wenhan Jiang <fnref n="2" />, Wenzhuo Miu <fnref n="2" /> and Ning Li <fnref n="4" />,*

Received: 27 May 2019; Accepted: 13 June 2019; Published: 17 June 2019

**Abstract:** Plant cells are separated by cellulose cell walls that impede direct cell-to-cell contact. In order to facilitate intercellular communication, plant cells develop unique cell-wall-spanning structures termed plasmodesmata (PD). PD are membranous channels that link the cytoplasm, plasma membranes, and endoplasmic reticulum of adjacent cells to provide cytoplasmic and membrane continuity for molecular trafficking. PD play important roles for the development and physiology of all plants. The structure and function of PD in the plant cell walls are highly dynamic and tightly regulated. Despite their importance, plasmodesmata are among the few plant cell organelles that remain poorly understood. The molecular properties of PD seem largely elusive or speculative. In this review, we firstly describe the general PD structure and its protein composition. We then discuss the recent progress in identification and characterization of PD-associated plant cell-wall proteins that regulate PD function, with particular emphasis on callose metabolizing and binding proteins, and protein kinases targeted to and around PD.

**Keywords:** plant cell walls; plasmodesmata; callose; callose synthase

## 1. Introduction

Plasmodesmata (PD) span the plant cell walls and connect protoplasts of adjacent cells [1,2]. The presence of PD allows molecular communication conveniently between plant cells [3–5]. Several thousand PD form a network between fully differentiated plant cells [6], transforming individual cells into interconnected symplasts, providing channels for the intercellular exchange of electrical signaling, diffusion of small molecules, and transportation of macromolecules such as proteins, nucleic acids, and viruses [7–10]. The exchange channel provides a direct cell-to-cell cytoplasmic pathway for plant molecule transport and information transmission [11]. The PD also regulates plant metabolic activity and gene expression through the transfer of signaling chemicals, enabling plant cells to exchange information with each other to coordinate physiological processes [12–14].

Although being a functional important structure, the molecular composition of PD remains largely elusive [15,16]. PD are embedded in rigid cell walls; it is arduous to purify and study such an organelle via classical biochemical methods. High-resolution electron microscopy showed that proteinaceous particles are associated with PD structures [17]. Studies such as immunolocalization revealed that

<footnote n="1">¹ College of Biological Science and Engineering, Fuzhou University, Fuzhou 350116, China; hanxiao@caas.cn</footnote>

<footnote n="2">² College of Forestry, Central South University of Forestry and Technology, Changsha 410004, China; nghua@126.com (L.-J.H.); jiangwenhan1999@126.com (W.J.); alic5e@163.com (W.M.)</footnote>

<footnote n="3">³ Biotechnology Research Institute, Chinese Academy of Agricultural Science, Beijing 100081, China; gygzgzyx@126.com</footnote>

<footnote n="4">⁴ State Key Laboratory of Cultivation and Protection for Non-Wood Forest Trees, Ministry of Education, Central South University of Forestry and Technology, Changsha 410004, China</footnote>

<footnote>* Correspondence: nli@csuft.edu.cn; Tel./Fax: +86-0731-8562-3406</footnote>

<footnote>† These authors contributed equally.</footnote>
various proteins are present in PD, including kinase-like proteins [18], actin, and myosin [19], which may serve as permanent structural components. In addition, genetic and biochemical approaches applied to dissect PD-associated functional components led to the discovery of callose binding proteins and receptor-like kinase proteins in the model plants [20]. They regulate the permeability of PD and control intercellular substance transport and signal communication. PD-mediated symplasmic transport is involved in a variety of plant physiological processes that are pivotal for the development of plants of economic interest, such as shoot apex dormancy, flowering, fruit ripening, fiber elongation, and plant-microbe symbiosis [21]. Understanding PD function through characterization of PD-related proteins may have the potential for improving agronomic traits in the future.

It is generally believed that callose deposited at the PD neck region plays a regulatory role in the neck switch [22]. This is the most well-known mechanism for regulating PD permeability [23]. Callose is a polysaccharide in the form of β-1,3-glucan [24]. Callose is synthesized at the plasma membrane at the neck region of PD, squeezing the plasma membrane inwardly, thereby narrowing down the neck region, which reduces the free space that PD can use for molecular passage [25]. A large amount of evidence indicates that the change in PD permeability is due to the synthesis and hydrolysis of callose in the cell wall [26–28]. Callose synthase, β-1,3-glucanase, and PD-associated callose binding protein are involved in the regulation of callose homeostasis in PD, which in turn affects the permeability of PD channels [29,30].

## 2. The Formation and Structure of PD

From the perspective of origin, PD is generally formed during the mitotic cell division. In the primary formation of PD, although the protoplasm is split by the newly formed cell plate, the endoplasmic reticulum (ER) is still connected through the cell plate [31]. The presence of the ER in the channels prevents the deposition of wall-forming substances, thus maintaining the connection between the two separated cells [32]. Furthermore, the ER transforms into the PD under the pressure of the cell plate or membrane. There are also PD between non-sister cells, that is, PD can be formed between non-mitotic cells [6,33]. However, the molecular origin and process of secondary PD formation are not well understood.

Ding et al. [34] obtained a series of new PD structural details using rapid freezing and freezing replacement sample preparation techniques, high-resolution electron microscopy, and computer-aided image processing techniques. A structural model of PD was proposed based on their observations. At both ends of PD is an enlarged hole [35,36], and the center of the PD is a cylindrical body formed by the appressed ER, called a desmotubule [37]. The desmotubule is tightly constricted with little space [38]. Cytoskeletal proteins are often found in PD, such as actin and myosin [39,40]. Protein particles of about 3 nm are embedded between the appressed ER and the plasma membrane. Electron-dense radial fibrils connect the protein particles [34,41]. The protein particles on the ER membrane present a spiral or a series of circularly arranged rotations. In the cross-section, 7–9 particles can be seen [34]. The channel of the ER is considered to be the transport pathway of PD [42,43]. The two ends of the channel become smaller, called the neck region [37].

Actin filaments and callosum sphincters are two modes of controlling PD permeability. According to experimental observations, actin filaments are located near and inside the PD channel, and the polymerization state of actin changes the permeability of PD [44,45]. On the other hand, callose aggregates at the neck region narrow the space between cell wall and PD desmotubule.

## 3. PD-Associated Structural Proteins

As mentioned above, studies such as immunolocalization revealed that various proteins such as actin, myosin, tubulin, and calreticulin are present in PD as structural components [39,46]. Actin and myosin are components of the cell dynamic network, cytoskeleton, involved in intracellular transport [47]. Microtubule arrays consist of tubulin heterodimers which play critical roles in cell
division and intracellular organization [48]. These proteins are not cell-wall proteins, but are associated with the plant cell-wall structure of PD in the context of this review (Table 1).

<table><caption>**Table 1.** Proteins associated with plasmodesmata (PD).</caption><thead><tr><th>Gene</th><th>Protein Function</th><th>Method</th><th>Biological Function</th><th>References</th></tr></thead><tbody><tr><td>Actin</td><td>Cell cytoskeleton</td><td>Immunolocalization</td><td>PD structure component</td><td>[49–51]</td></tr><tr><td>Myosin</td><td>Cell cytoskeleton</td><td>Immunolocalization</td><td>PD structure component</td><td>[46,51,52]</td></tr><tr><td>Tubulin</td><td>Cell cytoskeleton</td><td>Immunolocalization</td><td>PD structure component</td><td>[51]</td></tr><tr><td>GSL8 (CALS10)</td><td>Callose synthase</td><td>Genetics and cell biology</td><td>Callose deposition and auxin response</td><td>[53]</td></tr><tr><td>GSL12 (CALS3)</td><td>Callose synthase</td><td>Genetics and cell biology</td><td>Callose deposition and phloem development</td><td>[36]</td></tr><tr><td>PDBGs (AtBG_ppap)</td><td>β-1,3-glucanase</td><td>Proteomics and cell biology</td><td>Callose degradation and lateral root development</td><td>[28,54,55]</td></tr><tr><td>PDCBs</td><td>Callose binding protein</td><td>Proteomics and cell biology</td><td>PD localization and callose stabilization at neck region</td><td>[29]</td></tr><tr><td>PDLPs</td><td>Transmembrane receptor-like protein kinase</td><td>Proteomics and cell biology</td><td>SAR- and SA-induced callose accumulation</td><td>[55–59]</td></tr></tbody></table>

### 3.1. Actin

Actin exists in the entire PD [60]. The systematic observation of the nature and structure of PD in the epidermal cells of *Allium sativum* L. was performed using a fluorescence probe and confocal microscope combined with transmission electron microscopy and immunogold labeling [49]. White et al. [49] indirectly demonstrated the presence of actin in PD in young and mature cell walls of *Hordeum vulgare* and *Nicotiana plumbaginifolia* by colloidal gold labeling. The results effectively confirmed the presence of F-actin in PD. It is unclear how the filaments are organized within PD. They may exist in the lumen between the desmotubule and the plasma membrane, connecting the cytoskeleton between adjacent cells [45,61]. The actin filaments may also function as routes along which vesicular trafficking is possible. Treatments with actin disrupter, cytochalasin D, and actin-stabilizing fungal toxin, phalloidin, showed opposite effects on the cell-to-cell transport of a fluorescent reporter dextran; the latter chemical severely restrained the dextran movement [49,50]. These results indicate that the PD structural component may also be involved in regulating the PD permeability, with larger molecules being able to move through once actin filaments are disrupted.

### 3.2. Myosin

Immunochemical localization studies indicated that myosin was also an integral component of PD [49,51,52]. Radford and White's experiments indirectly demonstrated a myosin distribution on PD [52]. They used an animal myosin antibody with colloidal gold to prepare an immuno-electron microscopic sample of onions, maize, and *Arabidopsis* seedlings. They found colloidal gold-labeled particles on the cytoplasm and PD, suggesting that myosin might be present in PD. To date, myosin is confirmed to belong to 15 families, of which only some are found in plants, where the myosin found in
PD belongs to the eighth family. The effects of myosin inhibitor treatments somehow led to inconsistent results as to the role of myosin in PD [39]. How myosin regulates PD function is still under debate.

### 3.3. Tubulin

Blackman and Overall systemically characterized whether the cytoskeletal-related proteins were targeted to PD and found tubulin in the extracts of the PD-containing internode cell walls of corals, whereas it was not detected in the PD-independent internode cell walls, suggesting that PD may contain tubulin [51]. The role of tubulin in PD is less well characterized as compared to actin and myosin [62]. Blackman and Overall [51] speculated that tubulin may be indirectly involved in long-distance transport.

## 4. PD-Associated Regulatory Proteins

Callose is deposited at the cell wall near the neck of the PD and regulates the PD size exclusion limit (SEL) [22]. The PD function in plants is closely linked to callose deposition in the PD neck region. PD-associated regulatory proteins are more or less callose-related (Table 1). As mentioned above, the level of callose in PD is controlled by two antagonistic callose metabolic enzymes, callose synthase (CALS), also well known as glucan synthase-like (GSL), and β-1,3-glucanase (BG), and requires other enzymes that regulate callose stability [24,35]. In addition, the PD permeability changes involved in plant development and defense, processes involving callose, also require other proteins associated with callose [63,64].

### 4.1. Callose Synthases Modify Cell Wall PD to Regulate Plant Development

In the callose-dependent PD permeability, the PD channel is turned off by a high level of callose and turned on by a low level of callose, indicating that the callose level of the PD neck region is particularly crucial to PD function. The callose deposition determines PD SEL, which in turn determines their permeability and the transportation of macromolecules. Biochemical and genetic studies in barley and tobacco pollen tubes first demonstrated that GSLs produced callose [65]. This is also evidenced by studies in *Arabidopsis* [36,66]. GSLs control developmental signals by modulating the amount of callose in PD.

Plant hormones play a role in many aspects of plant physiology and development. Auxin has a gradient distribution in plants and plays an important role in plant growth and development [67]. As a small molecule, auxin can diffuse freely through the PD [68]. A recent study revealed that the auxin gradients can be maintained by controlling the PD permeability. Han et al. [53] screened all 12 *GSL* gene mutants in *Arabidopsis*, and found that the PD callose content decreased significantly only in the *gsl8/cals10* mutant. They used hypocotyls that responded to the auxin gradient distribution as experimental systems and found that the inducible *gsl8* RNA interference (RNAi) system showed lower phototropism and agravitropism in hypocotyl, which mimics auxin defects. Further studies found that a decrease in the messenger RNA (mRNA) level of *GSL8* resulted in the decrease of callose at PD, the increase of permeability of PD, and the increase of the diffusion of auxin transported by the symplast; finally, the gradient distribution of auxin was destroyed. Moreover, they proposed that the auxin gradient in plant tissues was established by the auxin-GSL8 feedback circuit associated with auxin response factor 7 (ARF7).

PD-mediated signaling plays a role in plant morphogenesis. Callose biosynthesis in the regulation of symplasmic transport by PD plays a central role in plant cell-to-cell signaling, but it is not clear how this regulation is arranged [69]. Vatén et al. [36] screened for the *gsl12/cals3* mutant in an ethyl methanesulfonate (EMS) mutagenesis library of *Arabidopsis*. Observation of the GFP::GSL12 fusion protein expressed in the leaves indicated that it was localized in PD. The gain-of-function mutation of *GSL12* leads to a transient accumulation of callose in PD in the early stage of root development, a decrease in PD SEL, a decrease in intercellular transport, and a defect in root development. This suggests that the regulatory activity of *GSL12* influences the biosynthesis of callose in the cell wall surrounding the PD,
which determines the pore size of the PD and regulates the transport of substances. The experiment with the *GSL12* allele (*cals3m*) showed that the PD pore size between plant tissues is controlled spatially and temporally. Using this system, they assessed the movement of the short-root (SHR) protein, microRNA165/6, and cytokinin in *Arabidopsis* roots and revealed the role of symplasmic communication in lateral root development. The above results indicate that *GSL12* is a callose synthase, and that the biosynthesis of callose affects the plant developmental signals transmitted by PD. Interestingly, Yadav et al. [66] also identified *GSL12* in *Arabidopsis*, and its semi-dominant mutants overproduce callose in PD.

The new cells required for plant growth and development are derived from the differentiation of stem cells. There is a group of stem cells called stem-cell niche (SCN) at the root apical meristem, and at its center are slowly dividing cells called the quiescent center (QC). It was believed that the location information provided by intercellular communication enabled QC and SCN to be maintained, but it was not verified for a long time. Liu et al. [70] developed a system that can effectively block PD-mediated communication in QC. They controllably and specifically blocked the intercellular communication between QC and neighboring cells by expressing *icals3m* (inducible form of *cals3m*) in QC. Using this system, they explored the function of intercellular communication in the SCN. They found that changes in symplastic communication between QC and adjacent cells resulted in elevated local auxin concentrations and the establishment of AP2-domain transcription factor gradients. Changes in symplasmic communication not only affect the polar transport of auxin, but also affect the biosynthesis of local auxin, which together provide local information for stem cell maintenance and play a key role in the maintenance of SCN. These studies revealed a direct link between symplasmic translocation and auxin-dependent regulation of stem-cell identity.

Slewinski et al. [71] identified a callose synthase in maize. They obtained tie-dyed (TDY) mutants of maize exhibiting green and yellow variegated leaves by EMS mutagenesis, and the yellow leaf tissue accumulated too much soluble sugar and starch. After constructing the population, they cloned the *TDY2* gene using map-based cloning, and found that *TDY2* encodes a callose synthase. Characterization of expression patterns by RT-PCR analysis, RNA in situ hybridization, and colocalization with the vascular marker, PINFORMED1a (ZmPIN1a)-YFP, revealed that *TDY2* was preferentially expressed in vascular tissues of developing young leaves. The movement experiments of ¹⁴C-labeled sucrose and carboxyfluorescein diacetate (CFDA) showed that the *tdy2* leaves defected in the entry of sucrose into phloem. Solutes could not enter phloem in yellow leaf region of *tdy2* mutants, but its long-distance transport capacity was not impaired. The ultrastructure study of the *tdy2* mutant vein by transmission electron microscopy showed that the phloem and xylem cells in the yellow leaf region of *tdy2* were changed, and that the vein differentiation was incomplete, which hindered the symplasmic transport from phloem companion cells to sieve elements mediated by PD, resulting in functional defects in phloem. As to how the callose synthase TDY2 targets to PD in the cell walls, Baker et al. [72] hypothesized that TDY2 was synthesized in the ER and traveled through secretory vesicles from the Golgi apparatus to localize to PD, or that TDY2 proteins translocated through the secretory system to the plasma membrane and PD.

Song et al. [73] identified a plasmodesmal callose synthase gene *cannot reach the roof 1 (CRR1)* that affects ovary development in rice plants. The rice *CRR1* gene is closely related to *Arabidopsis* *GSL8* and *GSL10*. By aniline blue staining and immunocolloidal gold analysis, the *crr1* mutation had no effect on the callose deposition at root cell plates, pericarp, and pedicel sieve plates. However, iodine staining of ovarian development showed that the starch granules were significantly less numerous than wild type (WT) in the pericarp of the *crr1* ovary, and that the distribution was uneven, indicating that the unloading from the phloem to the pericarp could be defective. RT-PCR analysis and *pCRR1::GUS* reporter transgenic plants showed that *CRR1* was specifically expressed in vascular tissues. The movement analysis of CFDA revealed that sugars were inefficiently unloaded from the lateral vascular system to the developing caryopsis. Transmission electron microscopy showed that the connection between sieve tube cells and peripheral parenchyma cells of vascular bundles was
reduced, and some cells had defective cell walls, where the deposition of callose at PD was reduced. These results suggest that CRR1 may alter the transport of cell fate determinants by controlling the permeability of PD, resulting in the change of the vascular cell differentiation pattern, thereby playing a key role in determining the plant reproductive development.

## 4.2. Callose Hydrolases Degrade Callose to Enhance Intercellular Communication

Class I β-1,3-glucanase (GLU I) plays an important role in the defense against virus infection [74]. Iglesias et al. [75] studied the role of GLU I in PD and PD-dependent communication. They obtained a homozygous class I β-1,3-glucanase (GLU I)-deficient mutant tobacco (TAG 4.4) by antisense transformation. This mutant become insensitive to viral infections. Observation was carried out of the size of the necrotic area and the GFP expressed by a recombinant form of potato virus X (PVX). GFP showed that the spreading of the virus in this mutant was slowed down. Observation of the fluorescence of CMV's 3a movement protein in fusion with GFP (3aMP::GFP) revealed that the movement of viral movement proteins between cells was limited in TAG 4.4. Observations of fluorescein isothiocyanate (FITC)-dextrans and FITC-peptides indicated that plasmodesmal SEL was reduced in this mutant. Aniline blue staining showed that, after the loss of GLU I, the deposition of callose increased. The above results indicate that β-1,3-glucanase plays an important role in the metabolism of callose. It may promote the degradation of callose, change the permeability of PD, and regulate the movement of macromolecules through PD. Bucher et al. [76] further explored the relationship between GLU I and plant sensitivity to viruses. They studied the role of GLU I in viral migration between cells from both gain- and loss-of-function mutants. They inserted the different target fragments into the TMV-based expression vector to obtain products of different enzymatic activities. The positive insertion fragments expressed the active GLU I protein, and the necrotic area of leaves increased continuously after infection with the virus. The plants infected with viruses expressing antisense GLU I showed reciprocal effects. These experiments indicated that GLU I promoted the movement of virus between cells by hydrolyzing callose to increase PD SEL.

Levy et al. [28] identified a putative plasmodesmal associated protein β-1,3-glucanase (AtBG_ppap) in *Arabidopsis* and demonstrated that the permeability of PD is regulated by β-1,3-glucanase. The proteins in the PD-enriched fraction of *Arabidopsis* were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and subjected to mass spectrometry analyses. The β-1,3-glucanase identified was named AtBG_ppap (beta-1,3-glucanase_putative PD-associated protein). Observation of transient and stable expression of GFP fusion proteins and aniline blue staining revealed that AtBG_ppap was related to the callose around PD in the cell walls. In the *Atbg_ppap* mutant, the callose level around the PD was higher, and the mobility of the GFP protein between cells was decreased, that is, the permeability of PD was lowered. The above experimental results demonstrated that AtBG_ppap is a PD-associated protein involved in plasmodesmal callose removal.

Intercellular communication regulates the function of individual cells to establish organ patterning and formation. It is long known that mobile signals are important in lateral root (LR) development. Paradoxically, the role of the intercellular channel PD in this process seems to have inadvertently not been elucidated. Recently, Benitez-Alfonso et al. [54] identified two plasmodesmal localized β-1,3-glucanase (PDBG) proteins that particularly accumulated in the LR founder tissue. Callose deposition changed correlating with changes in symplasmic transport during the course of LR development. During the formation of lateral meristems, callose amount was detected at low levels in PD. At early stages of LR development, callose accumulated to a moderate level between cells. Callose formation increased progressively when LRs developed. Such callose accumulation during LR organogenesis was regulated by two PDBGs, PDBG1 and PDBG2, which were localized in PD at early stages of lateral root primordia. In *pdbg1,2* double mutants, excessive callose deposition was detected around LR founder cells. In contrast, callose levels were significantly reduced in *PDBG1*-overexpressing plants. Consistent with callose accumulation levels, *pdbg1,2* double mutants showed a significant
increase in LR formation. These results demonstrated that PD-dependent cell-wall glucanases play an
important role in plant development via modulating intercellular communication.

Furthermore, Rinne et al. [77] analyzed 10 presumed cell-wall β-1,3-glucanase genes belonging to the GH17 family (glucan hydrolase family 17) in poplar, and found that different treatments, such as photoperiod, chilling, and gibberellin, regulated these genes differently. Based on this, a model of breaking bud dormancy related to PD regulated by β-1,3-glucanase was proposed. Previous studies found that flowering locus T (FT) is the primary long-distance signal that is generated at the vascular bundle by activation of Constans (CO) and transported to the apex [78]. Centroradialis-like 1 (CENL1) in poplar is an ortholog of *Arabidopsis* TFL1 (terminal flower 1)—a hypothetical signal peptide, expressed in a small region under the shoot apical meristem (SAM) [79,80]. Both FT and CENL1 move through the phloem. They enter phloem tissue via PD which connect the cytoplasm of companion cells with the sieve tube cells. When the bud is dormant, SAM closes PD with callose, and cells in SAM become disconnected from each other [80]. Chilling induces FT and affects gibberellic acid 3 (GA3) biosynthesis by upregulating the expression level of *GA3-oxidase*; then, FT and β-1,3-glucanase proteins induced by GA3 reopen the PD in dormant buds. When the climate warms up, the reopened PD enables FT and CENL1 to move to their targets to cause buds to germinate. Following the study of the mechanism of seasonal dormant axillary buds (AXBs) releasing dormancy, Rinne et al. [81] continued to study the mechanism of semi-dormant axillary buds releasing dormancy during tree branch formation. AXB dormancy breaking also involved the GA pathway, and the expression of β-1,3-glucanase gene was induced by GA. They founded that the GA synthesis gene *gibberellin 3-oxidase 2* (*GA3ox2*) expression level in AXBs was very low; thus, GA was absent in AXBs. The expression of GA receptor gene *gibberellin insensitive dwarf 1-like* (*GID1-like*) gradually increased during the maturation of semi-dormant AXBs; thus, AXBs were sensitive to GA. They found that the expression of *GA3ox2* was significantly upregulated after decapitation of the hybrid poplar; thus, the biosynthesis of GA in AXBs increased. The GH17s have CBM43 modules that position them to callose around the PD. Upregulated GH17s responded to increased GA, degraded callose in the cell wall near PD, reducing callose deposition at PD, and promoted communication between stems and buds, thereby abolishing dormancy of semi-dormant AXBs. They studied the expression patterns of GA pathway and the *GH17s* gene at different stages of bud development, summarized their expression characteristics, and demonstrated the role of PD-related β-1,3-glucanases in bud development. Conversely, photoperiod regulates seasonal growth; prior to the advent of winter, short days (SDs) during late autumn promote bud dormancy to induce growth cessation. It was found that, in hybrid aspen trees, short photoperiods act through activation of the abscisic acid (ABA) pathway [82]. Later, gene expression analysis revealed that SVL (SVP-like), an ortholog of the *Arabidopsis* floral repressor short vegetative phase (SVP), was induced by SDs in an ABA-dependent manner. SVL negatively regulated the growth promotive GA pathways and positively mediated the expression of *callose synthase 1* (*CALS1*) in the shoot apex to induce callosic plugs in the neck region of PD [83,84]. Thus, PD-associated callose synthases and glucanases in the cell walls play important roles in plant development by regulating bud dormancy and floral transition.

Previous work showed that PD is momentarily closed during the elongation of cotton fibers
in the allotetraploid species *Gossypium hirsutum* [85]. Ruan et al. [86] applied fluorescence probe
carboxyfluorescein (CF), aniline blue staining, and immunoelectron microscopy to cotton genotypes
with different cotton fiber lengths. It was found that the closure time of cotton fiber PD varied in
different cotton genotypes, which was related to the deposition and degradation of callose at the base
of cotton fiber. They cloned the *GhGluc1* gene from cotton fibers and found that it specifically encoded
a β-1,3-glucanase localized to the fiber cells. Moreover, they also found that callose was degraded
when GhGluc1 was expressed, and GhGluc1 expression was higher in plants with shorter cotton fibers.
These results indicate that GhGluc1 is a callose degrading enzyme that allows the PD to be in an open
state. Callose formation at the neck region closes PD. PD closure is positively correlated with fiber
length; therefore, the callose catabolic enzyme GhGluc1 plays a negative role in fiber elongation.
### 4.3. PD-Associated Callose Binding Proteins (PDCBs) Regulate Callose Stability

Simpson et al. [29] identified a novel PD-related protein gene, *AT5G61130*, which encodes PD callose binding protein 1 (PDCB1) with callose binding activity in *Arabidopsis*. PDCB1 fused to the YFP coding sequence driven by the CaMV 35S or its native promoter was stably transferred into *Arabidopsis* and transiently expressed in tobacco protoplasts. The fluorescent spots were found in the epidermal cell wall of *Arabidopsis* leaf tissue. Moreover, in spongy mesophyll cells, the fluorescent points were limited to the wall-to-wall junction between adjacent cells and still retained this pattern after plasmolysis. The results indicate that PDCB1 is associated with cell walls. Immunolocalization with anti-PDCB1 antiserum indicated that PDCB1 is located at the outer neck region of the PD. Combined with aniline blue staining and confocal microscopy, they proved that PDCB1 was colocalized with callose located in PD. They also studied PDCB2 (*AT5G08000*) and PDCB3 (*AT1G18650*) belonging to this family. A homology search of the *Arabidopsis* genome revealed that PDCB2 and PDCB3 have more than 50% amino-acid sequence similarity to PDCB1. Phylogenetic analysis revealed in fact that PDCB1, -2, and -3 lack a GPI-anchor signal sequence, but contain an X8 domain adjacent to the β-1,3-glucanase catalytic domain. This is in contrast to the previously identified PD-related β-1,3-glucanase [28]. By comparing with the structure of OLE-E9, the X8 domain-containing protein with callose binding activity in olives, it was found that the PDCB1 X8 domain is very similar to the OLE-E9 domain. Gel retardation assays showed that OLE-E10, PDCB1, and PDCB2 have callose binding activity in vitro. GFP diffusion assay showed that the migration of GFP decreased with the increase of PDCB1 expression, suggesting that the deposition of callose mediated by PDCB affected the symplastic communication through PD between cells.

PDCBs contain a carbohydrate binding module family 43 (CBM43) domain (also known as an X8 domain), as well as a callose binding domain, which can stabilize callose and regulate the dynamics of callose of PD [29]. PDCB1-overexpressing plants exhibited increased callose deposition in PD and decreased intercellular movement of reporter proteins. PDCBs are also predicted to be glycosylphosphatidylinositol anchored proteins (GPI-APS). Zavaliev et al. [87] studied the effects of GPI modification on PD proteins. They selected two GPI-APs associated with the callose of PD, BG_ppap and PDCB1 in *Arabidopsis*. They also chose two GPI-APs that were not associated with PD as control, arabinogalactan protein 4 (AGP4) and lipid transfer protein 1 (LTPG1). They first verified the subcellular localization of these proteins with monomeric citrine (mCitrine) labeling and then verified the modification of GPI with mannosamine. Subsequently, they subcellular localized the full-length sequence and deleted GPI signal sequence, respectively. It was found that PD targeting of BG_ppap and PDCB1 was mediated by GPI, and that the GPI modification played a major role in protein targeting to PD; furthermore, this sorting happened before positioning.

### 4.4. PD-Located Receptor-Like Proteins (PDLPs) Regulate Plant Defense Responses

PDLPs are PD-localized transmembrane proteins. PDLPs consist of a short cytosolic tail and two extracellular cysteine-rich receptor-like kinase domains of unknown function (DUF26) [88,89]. In *Arabidopsis*, eight PDLPs were identified by homologous comparison [56]; PDLP1, PDLP5, and PDLP6, were later found in independent proteomic screens of PD-enriched membrane [55,57]. PDLP1 (*AT5G43980*) is the first member of this protein family identified by cell-wall proteomics in *Arabidopsis* [55]. PDLP1 belongs to the DUF26 domain family. Thomas et al. [58] developed *Arabidopsis* transgenic plants overexpressing *PDLP1* and performed GFP diffusion experiments, confirming that PDLP1 can regulate intercellular transport by regulating PD. Moreover, they found through deletion analysis that their single transmembrane domain (TMD) is required for PDLP1a targeting to PD.

When fungi and oomycetes enter host plants with a haustoria structure, they cause plant defense responses, and the haustoria structure is gradually entrapped by the callose produced by the host. Caillaud et al. [59] studied the molecular mechanism of accumulation of callose around haustoria. They infected *pPDLP1::GUS* and *pPDLP5::GUS* reporter transgenic plants with pathogens, and found that
PDLP1 was specifically expressed in cells containing haustoria and responded to pathogen infection. Observation of PDLP1::GFP showed that PDLP1 localized at PD in both uninfected and infected tissue, and PDLP1 was associated with haustoria at the early stages of pathogen invasion in infected tissue. Moreover, the signaling peptide that targets PDLP to haustoria is located at its C-terminus, allowing it to be localized to the haustorial membrane. Significantly more callose was detected in transgenic plants overexpressing PDLP1 than in non-transgenic plants following pathogen infection. The experimental results of Caillaud et al. [59] directly demonstrate that PDLP1 contributes to the immune defense against pathogens by affecting the deposition of callose in the plant cell walls.

Pathogen invasion first leads to local infection, and then local infection produces signals that are transported to the uninfected parts of the plant, making the uninfected part resistant to subsequent enlargement of the infection. This is called systemic acquired resistance (SAR) [90–92]. Signals generated by local infection include salicylic acid (SA), azelaic acid (AzA), glycerol-3-phosphate (G3P), and reactive nitrogen species (RNS) [93,94]. Lim et al. [95] studied the transport mechanism of SAR signals and found that PDLPs were involved. They treated plants with mock solution or avirulent strains, then detected SA, AzA, and G3P in apoplastic fluids and petiole exudates representing apoplastic and symplasmic communication signals, respectively. It was found that SA was transported through the ectoplasmic pathway, while G3P and AZA were transported through the symplastic pathway mediated through cell-wall PD conduits. Evaluation of SAR in *pdlp1/pdlp5* double mutant or *PDLP5-overexpressing* plants showed that PDLP1 and PDLP5 were involved in SAR. Moreover, PDLP1 can not only affect the transmission of SAR signals by regulating the permeability of PD, but also affect the stability of SAR signal transporter azelaic acid induced 1 (AZI1) through direct protein–protein interactions [96]. Ye et al. [97] also reported that a PDLP1 homolog, PDLP8, was able to interact with the acyl-CoA-binding protein 6 (ACBP6) to stabilize ACBP6 accumulation in PD.

Various hormone signaling pathways are associated with basal immune responses. The pathways activated by SA, a defense hormone, contribute to the responses [90]. When a pathogen infects plants, SA concentration is elevated. The expression of the PD regulator *PDLP5* was upregulated resulting in the restriction of cell-to-cell movement via PD [57]. Accumulation of PDLP5 protein in PD restricts PD permeability by stimulating callose deposition at the PD neck. Most notably, Cui and Lee [98] revealed that *GSL6* (CALS1) is a primary partner for PDLP5. They produced a transcriptional reporter line *pGSL6::GUS*. GUS staining experiments confirmed that the *GSL6* expression profile is most highly correlated with that of *PDLP5* in SA-treatment experiments. In *gsl6-1* mutant plants, callose deposition did not change after treatment with SA. This indicates that SA-mediated callose production requires *GSL6*. The results of their experiments with carboxyfluorescein, a fluorescent dye that emits fluorescence and will not retrograde once it enters the cell, also support this conclusion. Neither changes in plasmodesmal callose deposition nor carboxyfluorescein movement were detected in *gsl6-1* mutant after SA treatment. Collectively, these sets of experimental evidence establish that *GSL6* is a bona fide callose synthase gene regulating SA-dependent plasmodesmal responses. Therefore, PDLP5 requires *GSL6* for SA-dependent plasmodesmal regulation. Evidence of subcellular localization of *GSL6* to cell-wall PD is missing. Since the *gsl10* mutant plants are gametophytic lethal [99], the *gsl10* loss-of-function plants are unavailable. The roles of *GSL10* in regulating PD permeability and plant defense responses are still not investigated. Wang et al. [100] cloned and overexpressed the extracellular domain of *Arabidopsis* PDLP5 in an insect expression system, followed by purification, crystallization, and diffraction analysis. The extracellular domain of PDLP5 includes two DUF26 domains. They collected X-ray diffraction data to 1.90 Å resolution, which indicated that PDLP5 may function as a dimer. This research provides novel understanding of the molecular mechanisms behind PD permeability regulation and plant system immunity [100]. It is still far from clear, however, how PDLPs regulate callose synthase gene expression and callose deposition in PD [101].
### 4.5. Other PD-Related Cell Wall Proteins and Even More to Be Identified

The proteins and lipids of eukaryotes are sequentially organized in the plane of the membrane, forming microdomains that distinguish and coordinate biological processes, ensuring specific kinetics and functions to specific regions within the cell [102–104]. Grison et al. [105] used *Arabidopsis* cultured cells and primary roots to strictly isolate plasma membrane (PM) domains of PD. Immunogold labeling and Western blotting showed that many false-positive membrane proteins, such as PMA2, CesA3, CesA6, and PIP2, did not target to PD, but previously known PDLP1 and PDCB1 were specific to PD PM [105].

A hypothetical sterol carrier protein gene, *GHSCP2D*, was isolated from elongated cotton fibers by Zhang et al. [106]. They inhibited the expression of *GHSCP2D* and observed the deposition of callose on PD, confirming that *GHSCP2D* is involved in the permeability of PD. They demonstrated that the decrease of PD permeability was due to the inhibition of the expression of the PD-targeted beta-1,3-glucanase GhPDBG3 in cotton fibers. Both downregulation of *GHSCP2D* and the application of a sterol biosynthesis inhibitor can reduce the permeability of PD, indicating that sterol also has an effect on PD.

Viruses move through PD between plant cells. Park et al. [107] studied the role of PD in viral infection and found that NbEXPA1 is a PD-specific expansin, specifically involved in cell wall relaxation. They performed a label-free quantitative proteomic analysis of PD-enriched fractions of leaves infected with TuMV and healthy controls in *Nicotiana benthamiana*. They collected in total 1070 PD protein candidates. After data processing, 100 upregulated proteins and 48 downregulated proteins were identified in the TuMV-infected samples. The observation of YFP or GFP fused with NbEXPA1 showed that the fusion protein had a dotted pattern along the cell-wall boundary, suggesting that NbEXPA1 might be located in PD. Co-localization between NbEXPA1::YFP and PDLP1::CFP and between NbEXPA1::GFP and CI::RFP indicated that NbEXPA1 was indeed a PD-localized protein. The results of proteomics and RT-PCR analysis showed that TuMV infection downregulates the gene expression and protein accumulation of NbExpA1. Colocalization of NbEXPA1::YFP and TuMV-6K2::mCherry suggests that NbEXPA1 may be involved in the movement of the virus between cells. Infection of leaves with recombinant virus expressing CFP ligated with a viral RNA polymerase NIB (CFP::NIB), combined with a yeast two-hybrid assay and BiFC analysis, suggested that NbEXPA1 acts in the viral replication complex via interaction with NIB. Silencing and overexpression of *NbEXPA1* inhibits and promotes the movement of the virus between cells, respectively. The above results indicate that NbEXPA1 is a cell-wall protein associated with PD, which affects viral gene replication and its movement through the PD between cells through interaction with NIB.

## 5. Further Perspectives

As the only channels for cytoplasmic communication between neighboring cells, PD are vital for the developmental process of plants. By combination of genetic, biochemical, and cell biology studies, many PD-associated proteins were identified and characterized (Figure 1). However, we still lack a fuller understanding of how the PD functions, which is largely due to the methodological difficulties in identification and characterization of PD components [108].

Increasing lines of evidence support that callose levels at the PD neck region play a central role in modulating the PD permeability and the symplasmic pathway. So far, the PD-related proteins, such as GSLs, PDGBs, PDCBs, and PDLPs, are all involved in maintaining callose homeostasis at the PD zone [28,29,36,53,56]. Further experiments are required to elucidate how transcription factors regulate the expression pattern of those genes in response to specific developmental cues and environmental stimuli. Since callose deposition and degradation in PD respond readily and rapidly to a wide range of signals, we speculate that PD-associated callose balancing proteins are mainly regulated at a post-transcriptional level. Another common feature is that those PD proteins contain transmembrane domains. Subcellular localization analysis showed that PD proteins are specifically targeted to PD
embedded in the cell wall. It is also of interest to address the routes via which those proteins reach PD and more specifically the membrane of PD [109].

<figure><img src="image_31.png" /><figcaption>**Figure 1.** A simplified schematic representation of plasmodesmata (PD) structural and functional proteins. Permeability of PD in the plant cell wall is controlled by dynamic callose deposition (closed, right side) and degradation (open, left side) at the neck region of PD. ER, endoplasmic reticulum; GSL, glucan synthase-like; PM, plasma membrane; PDCB, PD-associated callose binding protein; PDGB, plasmodesmal associated β-1,3-glucanase; PDLP, PD-localized protein.</figcaption></figure>

Previously, high-resolution electron microscopy and image processing methods greatly advanced our understanding of PD structure [110]. Immunochemical localization studies facilitated the detection of PD components. Indeed, our knowledge on PD is much dependent on technological progress. Despite their structural complexity, PD are important for plants and require further studies [21].

**Acknowledgments:** This work was supported by the Startup Foundation of Central South University of Forestry and Technology to N.L. (Numbers: 2018YJ049). We appreciate the Managing Editors and two anonymous reviewers for their corrections, suggestions, and in-depth comments, which greatly improved the manuscript.

**Conflicts of Interest:** The authors declare no conflicts of interest.

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© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).
<figure><img src="image_33.png" /></figure>

Article

*Arabidopsis* Trichome Contains Two Plasma
Membrane Domains with Different Lipid
Compositions Which Attract Distinct EXO70 Subunits

Zdeňka Kubátová¹, Přemysl Pejchar 1,2, Martin Potocký 1,2, Juraj Sekereš 2, Viktor Žárský 1,2 and Ivan Kulich 1,*

¹ Department of Experimental Plant Biology, Faculty of Science, Charles University, 12800 Prague, Czech Republic

² Institute of Experimental Botany, Czech Academy of Sciences, 165 02 Prague, Czech Republic

* Correspondence: kulich@natur.cuni.cz

Received: 21 June 2019; Accepted: 1 August 2019; Published: 3 August 2019

**Abstract:** Plasma membrane (PM) lipid composition and domain organization are modulated by polarized exocytosis. Conversely, targeting of secretory vesicles at specific domains in the PM is carried out by exocyst complexes, which contain EXO70 subunits that play a significant role in the final recognition of the target membrane. As we have shown previously, a mature *Arabidopsis* trichome contains a basal domain with a thin cell wall and an apical domain with a thick secondary cell wall, which is developed in an EXO70H4-dependent manner. These domains are separated by a cell wall structure named the Ortmannian ring. Using phospholipid markers, we demonstrate that there are two distinct PM domains corresponding to these cell wall domains. The apical domain is enriched in phosphatidic acid (PA) and phosphatidylserine, with an undetectable amount of phosphatidylinositol 4,5-bisphosphate (PIP₂), whereas the basal domain is PIP₂-rich. While the apical domain recruits EXO70H4, the basal domain recruits EXO70A1, which corresponds to the lipid-binding capacities of these two paralogs. Loss of EXO70H4 results in a loss of the Ortmannian ring border and decreased apical PA accumulation, which causes the PA and PIP₂ domains to merge together. Using transmission electron microscopy, we describe these accumulations as a unique anatomical feature of the apical cell wall—radially distributed rod-shaped membranous pockets, where both EXO70H4 and lipid markers are immobilized.

**Keywords:** cell wall; EXO70; exocyst complex; phosphatidic acid; phosphatidylinositol 4,5-bisphosphate; phospholipids; plasma membrane domains; polar exocytosis; trichome

**1. Introduction**

Despite their unicellularity, *Arabidopsis* trichomes grow into extraordinarily shaped and precisely polarized structures, which makes them a potent model of cell polarization and morphogenesis [1]. According to [2], trichome development can be divided into six stages. During the first stage, several rounds of endoreduplication lead to an increased DNA content and, later, to a remarkably large trichome cell. The second and third stages are crucial for trichome shaping because they involve oriented growth above the surface and two or three branching events. During the fourth and fifth stages, the branches elongate significantly and the trichome reaches its final size and shape. When the growth is complete, the trichome enters the last stage of its development, which is maturation of the cell wall. In this stage, the trichome is divided into two markedly different domains—the basal domain, with a thin cell wall, and the apical domain, with an extremely thick cell wall. This latter secondary cell wall (SCW) consists of at least two distinct layers—the outer, cellulose-rich layer and the inner, callose-rich layer [3]. The inner layer is autofluorescent and silicified in a callose-dependent manner [4].
The apical cell wall domain also contains surface papillae, which accumulate cuticular waxes that may be different from those in other epidermal cells [5]. Basal and apical trichome domains are separated by a ring-shaped, callose-rich structure named the Ortmannian ring [3].

The development of this cell wall is dependent on the EXO70H4 exocyst subunit, which is necessary for callose synthase delivery to the plasma membrane (PM) [4]. EXO70H4 is a subunit of the exocyst complex, which is a eukaryotic protein complex involved in the tethering of post-Golgi secretory vesicles, which carry membrane and cell wall components to the PM [6]. The exocyst is composed of eight different subunits forming a functional complex, including Sec3, Sec5, Sec6, Sec8, Sec10, Sec15, Exo70, and Exo84 (for review, see [7,8]). Exo70 and Sec3 have a special position in the complex, as both of these subunits are able to directly bind to the target membrane lipids [9–11], thus regulating where the secretion will occur [12]. In yeast and mammals, the Exo70 subunit recruits the rest of the exocyst complex to the PM via a specific interaction of the EXO70 C-terminus with phosphatidylinositol 4,5-bisphosphate (PIP₂) [9,13]. Sec3 is then responsible for initiating the binary SNARE (Soluble NSF Attachment Protein Receptor) complex [14]. The crosstalk of phospholipids with their protein partners and its importance in plant cell polarity determination and membrane traffic regulation was summarized in [15].

The *Arabidopsis* genome contains 23 EXO70 paralogs [16–18]. This multiplicity of EXO70 subunits in plants led to the hypothesis that EXO70s may have divergent lipid-binding properties and thus regulate exocytosis in distinct PM domains within a single cell [19]. So far, there is one documented example of such domain separation observed for *NtEXO70A1* and *NtEXO70B1* paralogs in *Nicotiana tabacum* pollen tubes, where EXO70A1 was more apically localized than EXO70B1 [20]. There is also apparent functional specialization among the EXO70 paralogs, as neither EXO70A1, B1, nor any other of 18 tested paralogs can complement the *exo70H4-1* loss-of-function mutant phenotype [4]. In this study, we document the differential localization of EXO70H4 and EXO70A1 in the mature trichome and show that EXO70H4 is required for the development and separation of two PM domains with different compositions of signaling phospholipids and distinct abilities to attract other EXO70 members.

## 2. Results

### 2.1. Mature *Arabidopsis* Trichomes Contain Two Distinct Lipid Domains and EXO70H4 is Involved in Their Formation

To investigate the distribution of the PM lipids in the *Arabidopsis* trichome, we observed localization of several fluorescent phospholipid markers. We focused on phosphatidylinositol-4-phosphate (PI4P) [21], phosphatidylinositol 4,5-bisphosphate (PIP₂) [21], phosphatidic acid (PA) [22,23], and phosphatidylserine (PS) membrane lipid markers [23]. While the PIP₂ marker was localized almost exclusively to the trichome base beneath the Ortmannian ring (Figure 1), the PI4P and PA markers were distributed evenly around the whole PM, with their signals enhanced at the Ortmannian ring and in intramural pockets throughout the apical domain (Figure 1). However, the PI4P marker signal inside these intramural pockets was weaker than the PA marker signal, which is highly accumulated there (Figure 1). The identities of these pockets will be described in more detail in Section 2.3. The PS marker was mostly visible at the apical membrane domain (Figure 1 and Supplemental Video S1). No signal was observed at the base, but a weak signal may have remained undetected due to a strong cytoplasmic background of this marker line. Thus, we concluded that a mature *Arabidopsis* trichome contains two distinct membrane domains with different lipid compositions.
<figure><img src="image_34.png" /><figcaption>**Figure 1.** Representative images of different phospholipid markers in wild type mature trichome. PI4P—phosphatidylinositol-4-phosphate; PIP₂—phosphatidylinositol 4,5-bisphosphate; PA—phosphatidic acid; PS—phosphatidylserine. Blue—cell wall autofluorescence; magenta—chlorophyll autofluorescence; green—mCitrine or YFP (yellow fluorescent protein). White arrows point at the Ortmannian ring. Scale bars = 20 µm.</figcaption></figure>

Next, we wanted to investigate when these two domains differentiated during trichome development. For this, we used a double marker line expressing a PA marker tagged with mCitrine and a PIP₂ marker tagged with mCherry (mCH), provided by the authors of [23]. In the wild type (WT) background, the distribution of the lipid markers was consistent with previous observations (Figure 2a). Distinct domains only appeared at stage 6 of trichome development, along with the establishment of the Ortmannian ring. In the WT trichomes of stage 4 (elongation), where the Ortmannian ring was not formed yet, PM domains were not visible (Figure 2b). The PIP₂ marker was evenly distributed all around the PM and the PA marker was not bound to the membrane at all (Figure 2b), suggesting that PA is not present in young trichome PM. Later, we introduced both lipid markers into the *exo70H4-1* mutant background, which was unable to finish the cell wall maturation [3]. Here, the membrane domains lacked a sharp border and were poorly visible (Figure 2c). We concluded that the establishment of the apical and basal membrane domains occurred while the EXO70H4-dependent SCW layer in the apical domain was formed, which depended on the SCW formation.
<figure><img src="image_35.png" /><figcaption>**Figure 2.** Lipid markers in WT and *exo70H4-1* trichomes. (a) Colocalization of PA and PIP~2~ markers in WT trichome; (b) colocalization of PA and PIP~2~ markers in WT trichome in the elongation stage; (c) colocalization of PA and PIP~2~ markers in an *exo70H4-1* mutant background. Blue—cell wall autofluorescence; magenta—chlorophyll autofluorescence; green—YFP; red—mCherry. White arrows point at the Ortmannian ring. Scale bars = 20 µm.</figcaption></figure>
## 2.2. Trichome Apical and Basal Plasma Membrane Domains Recruit Different EXO70 Proteins

To address the biological relevance of the trichome lipid domain distribution, we observed multiple EXO70 proteins under the control of the EXO70H4 promoter. EXO70A1 was previously found to bind to PIP₂ [24] and to localize to the PIP₂-rich region of the pollen tube [20]. Corresponding with its lipid affinity, EXO70A1 localized preferentially to the basal trichome domain and, in many cases, the apical trichome domain was completely devoid of EXO70A1, although there was a certain degree of variability and it was often seen also within the Ortmannian ring area (Figure 3a, Supplemental Figure S1). As reported previously [4], a lack of EXO70H4 resulted in EXO70A1 localizing all around the trichome PM (Figure 3a). To verify that EXO70A1 is natively present in *Arabidopsis* trichomes, we also generated a reporter construct of the EXO70A1 promoter fused to 2xGFP (green fluorescent protein). This revealed that EXO70A1 was indeed expressed in the mature trichome (Figure 3b).

EXO70H4 is known to localize to the apical trichome domain and to the Ortmannian ring [4]. To correlate its localization together with EXO70A1, we co-transformed EXO70H4p:mCHERRY-EXO70H4 (mCH-EXO70H4) with EXO70H4p:GFP-EXO70A1 (GFP-EXO70A1). In this case, the EXO70 isoforms localized to distinct and mostly non-overlapping PM domains that corresponded to the apical and basal trichome domains (Figure 3c).

We also investigated the localization of the remaining EXO70s, which we observed previously in the *exo70H4-1* mutant background [4]. This time, we expressed the constructs in a WT background to see their localization when it was unaffected by the *exo70H4-1* mutation. Eight paralogs were tested—EXO70A1, A2, B1, C1, D2, F1, H7, and H8. The only paralogs found to localize to the PM in the WT background in addition to EXO70A1 were EXO70A2 (localized similarly to EXO70A1) and EXO70H8. EXO70H8 strongly accumulated at the PA-rich domain in the apical part of the trichome, including Ortmannian ring and cell wall ingrowths. Although EXO70H8 mimicked EXO70H4 localization in the WT trichome, it was not capable of functionally complementing the *exo70H4-1* mutant phenotype in our previous cross-complementation study [4]. The other EXO70 paralogs that were tested remained in the cytoplasm or nucleus, as in the previous experiments in the *exo70H4-1* background.

Since the lipid-binding capacities of EXO70H4 have not yet been described, we performed a protein-lipid overlay assay with in vitro translated HA-tagged EXO70H4. This revealed a clear affinity to PS and conceivably PA, but no apparent binding to PIP₂ (Figure 3d). This corresponded well with the colocalization of EXO70H4 with PA and PS markers in the apical domain of the mature trichome. Based on these and previously published data, we concluded that different EXO70 proteins exhibit a specific capacity to bind membrane lipids and thus are recruited to distinct PM domains, contributing to the biogenesis of different cell wall domains within a single plant cell.
<figure><img src="image_36.png" /><figcaption>**Figure 3.** Trichome plasma membrane domains recruit different EXO70 proteins. (a) EXO70H4p::GFP-EXO70A1 (GFP-EXO70A1) preferentially localizes to the basal trichome domain in WT. This preference is lost in the *exo70H4-1* mutant. (b) EXO70A1p::GFP;GFP (A1p::2xGFP) expression marker in a WT trichome. (c) Colocalization of GFP-EXO70A1 with EXO70H4p::mCherry-EXO70H4 (mCh-EXO70H4). (d) Protein–lipid overlay assay of EXO70H4. Blue—cell wall autofluorescence; magenta—chlorophyll autofluorescence; green—GFP; red—mCherry. Scale bars = 20 µm.</figcaption></figure>
## 2.3. *The Apical Cell Wall Contains Entrapped Membranous Pockets*

While observing the apical trichome domain, we noticed that a large portion of the signal came from within the cell wall. This was true for all of the constructs which localized to the apical membrane domain, including markers of PA, PS, PI4P, mCh-EXO70H4, GFP-EXO70H8, and occasionally also EXO70A1 and EXO70A2 with a weak signal. This strange intramural localization directed our further investigation. The signal within the SCW was distributed in a pattern remarkably similar to the callose deposits shown before [3,4]. The radially distributed rays of signal were visibly embedded within the cell wall and were well-apparent on fluorescence microscopy optical cuts of matured branches together with a wrinkled PM-SCW interface (Figure 4a,b). To be sure that this was indeed the fluorophore signal and not cell wall autofluorescence, we performed lambda scans in plants expressing GFP-EXO70H4, mCh-EXO70H4, YFP-PA marker, and untransformed control. In all of these cases, the expected emission peak was observed (Supplemental Figure S2), with no such peaks in the negative control. To check if these intramural signals were still connected to the rest of the cytoplasm, we performed a fluorescence recovery after photobleaching (FRAP) experiment on the PI4P marker, which revealed that there was no detectable recovery of the signal, unlike in the case of the signal bleached within the apparent continuum of the PM (Supplemental Figure S3). We therefore hypothesized that the intramural signal may come from the PM and cell interior, being physically entrapped within the cell wall pockets.

To investigate the SCW structure in more detail, we used transmission electron microscopy (TEM). This revealed internal cell wall structures, supporting our observations of oblong traversing pockets with light microscopy. TEM cross-sections of mature WT trichome branches displayed SCW-transpassing structures, probably aggregates of entrapped secretory lipid membranes, proteins, and the cell interior, often arranged in a radial pattern of concentric transpassing channels (Figure 4c, left). These intramural pockets were obviously separated from the continuum of cytoplasm (Figure 4c, middle), but their PM origin was apparent from images where pieces of electron-dense, non-cell wall materials were embedded into the cell wall (Figure 4c, right).
<figure><img src="image_37.png" /><figcaption>**Figure 4.** Plasma membrane proteins reside within the trichome apical cell wall. (a) GFP-EXO70H4 (EXO70H4) signal, cell wall autofluorescence (autofluor.), and transmission channel (trans.) at the branch of a mature WT trichome. The dotted square represents area enlarged in (b). Scale bar = 20 µm. (b) Detail of (a). Scale bar = 10 µm. (c) Left—TEM image of a cross-section of a trichome branch. Entrapped cell interior (EC) is visible as concentric rays. Scale bar = 3 µm. Middle—detail of entrapped cell interior obviously separated from the cytoplasm. Scale bar = 1 µm. Right—a detail of one concentric ray. Scale bar = 200 nm.</figcaption></figure>

**3. Discussion**

We showed that the mature *Arabidopsis* trichome contains, along with two cell wall domains, two distinct PM domains that differ in their phospholipid composition and also in their ability to recruit different EXO70 proteins. EXO70A1 is recruited to the basal trichome domain, which is PIP₂-rich and contains a thin, pectin-rich cell wall [25]. EXO70H4 is recruited to the apical domain, which is PS- and PA-rich and has a thick, autofluorescent cell wall [3].

There are a number of different examples of cell types having different phospholipid domains and many of them can be linked with differential cell wall deposition. Pollen grains also accumulate PIP₂ at the site of the future aperture, which is an area marked with a thin cell wall [26]. Similarly, the pollen tube accumulates PIP₂ at its tip, while PS and PA are in the shank [22,23]. This corresponds
with the apical cell wall of the pollen tube being almost solely made of secreted pectins, while callose and cellulose are deposited in the shank [27]. This domain separation in the pollen tube is further marked by differential membrane localization of EXO70A1 and EXO70B1 in tobacco [20]. In contrast to the pollen tube, trichomes are extremely large and static structures. The border of the two membrane domains appears to be very sharp and defined by the Ortmannian ring. A lack of a sharp border between these two domains and a partial loss of polarity in the *exo70H4-1* mutant suggests that there is a positive feedback loop and that EXO70s have a role in domain development.

The observed lack of PIP~2~ and abundance of PA in the apical domain could also imply that, in the apical domain, PIP~2~ is metabolized to PA. In general, PA is formed through two pathways: First, by the direct hydrolysis of structural phospholipids by phospholipase D and second, through the consecutive actions of phospholipase C and diacylglycerol kinase, where the hydrolysis of PIP~2~ produces inositol-1,4,5-trisphosphate (released into the cytosol) and diacylglycerol within the PM, which is quickly phosphorylated at the membrane into PA by diacylglycerol kinase. PA produced by phospholipase Dα1 is a crucial signaling lipid, mediating the abscisic acid response in guard cells, where its role in NADPH oxidase-mediated reactive oxygen species (ROS) production was clearly demonstrated [28]. Both PA and phospholipase Dα1 have previously been linked with ROS production [29]. NADPH oxidase-mediated ROS production acts in the plant defense response [30,31]. In plants, PA is also produced in response to several stress factors, including pathogen attack [32]. EXO70H4 is also induced by *flagellin* 22 in epidermal pavement cells [4]. The apical cell wall domain of the trichome is also remarkably similar to the defense papillae, as they both accumulate callose deposits, phenolic compounds [33], and an extracellular signal of membrane proteins (such as SYP121—SYNTAXIN OF PLANTS 121 [34]). We therefore suggest that the mechanism and domain organization in the trichome may be a manifestation of a general response to pathogen attack, which runs constitutively in the trichome.

It was shown that binding of PA to its protein interactors is enhanced by a negative curvature stress and that a complex membrane lipid composition strongly influences lipid-protein interactions [35]. Surprisingly, EXO70 by itself is able to induce negative membrane curvature, specifically through the homodimerization mechanism, as was demonstrated in mammals [36]. Together, these biophysical properties could induce PM deformation sufficient to form extreme membrane curvatures, leading to the formation of membranous pockets within the SCW. Another factor contributing to the pocket formation may be the cuticular wax migration across the cell wall. The apical trichome domain displays several unique features, including surface papillae formation. Papillae start to form at stage 5 of trichome development (expansion) and continue developing during stage 6 (maturation). At these stages, the apical cell wall is already quite thick (>2 µm) [37]. Papillae are little bumps filled with lipophilic cuticular substances. These waxes differ from those of other epidermal cells by a high content of C35+ alkanes [5]. How these compounds get through the thick cell wall to the surface is not understood. Membranous pockets may drift along the migration routes of the cuticular waxes through the inner layer of the SCW. This feature may be very useful for the study of cuticular wax migration across the cell wall. Another intriguing possibility is that formation of these membranous pockets within the SCW is directly linked to the mechanism of callose deposition during SCW biogenesis—a possibility we aim to study.

# 4. Materials and Methods

## 4.1. Plant Material

Plants were grown in Jiffy soil pellets in standard growth chamber conditions (long day 16 h:8 h, 100–120 µM photosynthetically active radiation m⁻² s⁻¹). The *exo70H4-1* mutant line was described previously [3]. As WT control, outcrossed WT plants were used. Seeds of PI4P and PIP~2~ markers were obtained from Yvon Jaillais and are described in detail in [21]. From this set, we used line P21Y (mCitrine-2xPH FAPP PI4P binding domain), line P24Y (mCitrine-2xPHPLC PIP~2~ binding domain),
and line P24R (mCherry-2xPH<sup>PLC</sup> PIP<sub>2</sub> binding domain). Two different PA markers were used in this study. In Figure 1, it was YFP:NES-2xSpo20p (cloning described below) and in Figure 2 and Supplemental Figure S2, it was a double marker line expressing a PA marker tagged with mCitrine (PAY) and a PIP<sub>2</sub> marker tagged with mCherry (P24R, mentioned above), provided by [23]. PS marker cloning is described below.

## 4.2. Confocal Microscopy

Confocal microscopy images were taken on Zeiss LSM880 with C-Apochromat 403/1.2 W Korr FCS M27 objective [GFP (488): 508–540 nm, chlorophyll (488) 650–721 nm, cell wall autofluorescence (405) 426–502, mCherry (561) 597–641]. Images were processed using the Fiji platform [38].

## 4.3. Trichome Isolation and TEM

Leaves of 4-week-old plants were collected and incubated for 3 h in falcon tubes in a solution of acetic acid:ethanol (1:3) and then washed three times with deionized water. Washed leaves were transferred to a solution of 150 mM KH<sub>2</sub>PO<sub>4</sub> pH 9.5 and incubated overnight with shaking at 150–180 rpm. Released trichomes were collected by centrifugation (1 min, 1000 G, no break). For TEM, isolated trichomes were fixed for 24 h in 2.5% (w/v) glutaraldehyde in 0.1 M cacodylate buffer (pH 7.2) at 4 °C and postfixed in 2% (w/v) OsO<sub>4</sub> in the same buffer. Fixed samples were dehydrated through an ascending ethanol and acetone series and embedded in Epon-Araldite.

## 4.4. Cloning and Stable Transgenic Line Preparation

For cloning of the expression reporter construct EXO70A1p::GFP:GFP, the multisite gateway approach was used. The EXO70A1 promoter (1 kb upstream, EXO70A1 prom for and EXO70A1 prom rev listed in Supplemental Table S1 were used) was subcloned into pDONORP4-P1r and sequenced using M13 primers. GFP constructs—pEN-L1-F-L2 and pEN-R2-GFP-L3,0—and destination vector pB7m34GW were obtained from [39]. An expression clone was assembled from these by a multisite reaction using Gateway LR clonase (Thermo Fischer, Waltham, MA, USA). All EXO70H4p::GFP-EXO70XY constructs used in this study were previously described in [4].

To prepare the construct for the PA marker (pUBQ::YFP:NES-2xSpo20p PABD), first NES-2xSpo20p was amplified from YFP:Spo20p-PABD [22] and cloned together with the ubiquitin (UBQ) promoter and YFP into the binary vector pHD71 (kindly provided by Dr. Benedikt Kost).

To prepare the construct for the PS marker (pUBQ::YFP:C2<sup>LACT</sup>), YFP-C2<sup>LACT</sup> from Lat52:YFP-C2<sup>LACT</sup> [22] was cloned together with the UBQ promoter into the binary vector pHD71; the primers used are listed in Supplemental Table S1.

Final constructs were electroporated into competent cells of *Agrobacterium tumefaciens*, strain GV3101. Col-0 wild type plants were transformed by the floral dip method [40] and transformants were selected on kanamycin plates or soil by BASTA spraying (150 mg/L of glufosinate-ammonium). No fewer than five individual transformants were observed in each experiment and at least two biological replicates were made.

## 4.5. Protein-Lipid Overlay Assay

The protein-lipid overlay assay was performed using N-terminally HA-tagged EXO70H4 and echelon lipid strips. First, the EXO70H4 coding sequence was cloned into a pTNT vector (Promega, Fitchburg, WI, USA), optimized for in vitro coupled transcription and translation, using primers listed in Supplemental Table S1. The cloning was performed in two steps. First, the EXO70H4 coding sequence was amplified with the use of the primers EXO70H4-forward and EXO70H4-reverse. Subsequently, a megaprimer containing the HA-tag sequence and Kozak consensus sequence facilitating efficient translation initiation was used together with EXO70H4-reverse primer. The product was cloned into the pTNT vector via Sall and NotI restriction sites.
The construct was used as a template for in vitro coupled transcription and translation reactions using TNT® SP6 High-Yield Wheat Germ Protein Expression System (Promega), according to the manufacturer's instructions, in a total volume of 50 µL. To verify protein expression and stability, 10 µL of the yield was used in Western blotting. Since the protein was tagged on the N-terminus, Western blot analysis would have detected products of prematurely terminated translation, but none were observed (see Supplemental Figure S4).

The remaining 40 µL of the reaction product was used for the protein-lipid overlay assay. The echelon lipid strip (Elcheon Biosciences, Salt Lake City, UT, USA) was blocked for one hour in blocking solution (50 mM Tris, 150 mM NaCl, 0.05% Tween 20, pH 7.6 + 3% bovine serum albumine). After three washes in wash solution (50 mM Tris, 150 mM NaCl, 0.05% Tween 20, pH 7.6), the strip was incubated for two hours with primary mouse anti-HA antibody (Sigma-Aldrich, St. Louis, MO, USA) diluted 1:1000 in blocking solution. After three washes with wash solution, the strip was subsequently incubated with secondary anti-mouse antibody conjugated to horseradish peroxidase (Promega), diluted 1:20,000 in blocking solution. Finally, the strip was washed three times with wash solution and incubated for three minutes with Amersham ECL Prime Western Blotting Detection Reagent (GE Healthcare, Chicago, IL, USA) and imaged with Bio-Rad Laboratories (Hercules, CA, USA) Chemidoc gel imaging system. The experiment was repeated twice with consistent results.

**Supplementary Materials:** Supplementary Materials can be found at http://www.mdpi.com/1422-0067/20/15/3803/s1.

**Author Contributions:** Conceptualization, I.K. and V.Z.; validation, Z.K. and I.K.; investigation, Z.K., J.S. and I.K.; resources, P.P. and M.P.; writing—original draft preparation, Z.K. and I.K.; writing—review and editing, P.P., M.P., J.S. and V.Z.; visualization, Z.K. and I.K.; supervision, I.K. and V.Z.; funding acquisition, I.K.

**Funding:** This project was financed by Czech Science Foundation grant 18-12579S. Microscopy was performed in the Laboratory of Confocal and Fluorescence Microscopy co-financed by the European Regional Development Fund and the state budget of the Czech Republic (project no. CZ.1.05/4.1.00/16.0347). Part of income of V.Ž. is supported by the Ministry of Education, Youth and Sports of CR from European Regional Development Fund-Project "Centre for Experimental Plant Biology": No. CZ.02.1.01/0.0/0.0/16_019/0000738.

**Acknowledgments:** We would like to thank Yvon Jaillais for providing us with the lipid markers, Patrick Moxon for cloning EXO70A1 promoter, Jachym Metlicka for help with image preparation, and our technician, Marta Cadyova, for running the laboratory.

**Conflicts of Interest:** The authors declare no conflict of interest.

**Abbreviations**

FRAP fluorescence recovery after photobleaching

mCherry mCherry

PA phosphatidic acid

PI4P phosphatidylinositol-4-phosphate

PIP₂ phosphatidylinositol 4,5-bisphosphate

PM plasma membrane

PS phosphatidylserine

SCW secondary cell wall

TEM transmission electron microscopy

WT wild type

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<figure><img src="image_38.png" /></figure>

© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).
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Article

# Hydroxyproline-Rich Glycoproteins as Markers of Temperature Stress in the Leaves of *Brachypodium distachyon*

Artur Pinski¹, Alexander Betekhtin ¹,*, Katarzyna Sala², Kamila Godel-Jedrychowska ², Ewa Kurczynska ²,*, and Robert Hasterok ¹

¹ Department of Plant Anatomy and Cytology, Faculty of Biology and Environmental Protection, University of Silesia in Katowice, 40-032 Katowice, Poland; apinski@us.edu.pl (A.P.); robert.hasterok@us.edu.pl (R.H.)

² Department of Cell Biology, Faculty of Biology and Environmental Protection, University of Silesia in Katowice, 40-032 Katowice, Poland; katarzyna.sala@us.edu.pl (K.S.); kgodel@us.edu.pl (K.G.-J.)

* Correspondence: alexander.betekhtin@us.edu.pl (A.B.); ewa.kurczynska@us.edu.pl (E.K.); Tel.: +48-32-2009-484 (A.B.); +48-32-2009-447 (E.K.)

Received: 7 May 2019; Accepted: 23 May 2019; Published: 25 May 2019

**Abstract:** Plants frequently encounter diverse abiotic stresses, one of which is environmental thermal stress. To cope with these stresses, plants have developed a range of mechanisms, including altering the cell wall architecture, which is facilitated by the arabinogalactan proteins (AGP) and extensins (EXT). In order to characterise the localisation of the epitopes of the AGP and EXT, which are induced by the stress connected with a low (4 °C) or a high (40 °C) temperature, in the leaves of *Brachypodium distachyon*, we performed immunohistochemical analyses using the antibodies that bind to selected AGP (JIM8, JIM13, JIM16, LM2 and MAC207), pectin/AGP (LM6) as well as EXT (JIM11, JIM12 and JIM20). The analyses of the epitopes of the AGP indicated their presence in the phloem and in the inner bundle sheath (JIM8, JIM13, JIM16 and LM2). The JIM16 epitope was less abundant in the leaves from the low or high temperature compared to the control leaves. The LM2 epitope was more abundant in the leaves that had been subjected to the high temperatures. In the case of JIM13 and MAC207, no changes were observed at the different temperatures. The epitopes of the EXT were primarily observed in the mesophyll and xylem cells of the major vascular bundle (JIM11, JIM12 and JIM20) and no correlation was observed between the presence of the epitopes and the temperature stress. We also analysed changes in the level of transcript accumulation of some of the genes encoding EXT, EXT-like receptor kinases and AGP in the response to the temperature stress. In both cases, although we observed the upregulation of the genes encoding AGP in stressed plants, the changes were more pronounced at the high temperature. Similar changes were observed in the expression profiles of the EXT and EXT-like receptor kinase genes. Our findings may be relevant for genetic engineering of plants with increased resistance to the temperature stress.

**Keywords:** arabinogalactan proteins; *Brachypodium distachyon*; cell wall; extensins; immunohistochemistry; leaf; RT-qPCR; temperature stress

## 1. Introduction

Plant growth and productivity are compromised by various abiotic stresses, among which are high and low temperature stress. Even short periods of temperature stress may significantly decrease the yield, especially when it occurs during the crucial stages of plant development [1]. As was predicted, heat waves and other extreme temperature events are to become more intense, frequent and long-lasting
because of global climate change [2]. Thus, thermal stresses must be better understood in the context
of the response and adaptation of plants, which may enable crops with improved thermotolerance to
be obtained and bred [3].

*Brachypodium distachyon* L. Beauv. (Brachypodium), which is a member of the Pooideae subfamily, is a wild annual grass species that is widespread in the regions of the Mediterranean basin, Western Europe, the Middle East, south-west Asia, north-east Africa, North and South America and Australia. It is closely related to many temperate zone key cereals, including wheat, barley, rye, oats and various forage grasses [4]. Due to its numerous favourable biological features such as a relatively small nuclear genome that ranges from 270 Mb to 350 Mb (depending on the methodology that is used), small stature, self-fertility, a life cycle of less than four months and undemanding growth requirements, *B. distachyon* constitutes an excellent model species [5].

Low-temperature stress results in the downregulation of many photosynthesis-related proteins and, at the same time, the upregulation of the proteins that are involved in reactive oxygen species (ROS) scavenging, redox adjustment, cytoskeletal rearrangements, cryoprotection and cell wall remodelling [6]. Similar results are observed in plants that are stressed by a high temperature [7]. Though the cell wall structure is not primarily altered under heat stress, numerous studies have indicated various changes in its architecture. In low temperature stress, changes in the cell wall rigidity may be an important factor in thermotolerance. Changes in the cell wall are more pronounced in roots because they are more sensitive to temperature stresses than the aerial parts of a plant, though the adverse effect of such stress on leaves directly affects plant productivity. Alterations in the cell wall in response to temperature stress concern cellulose and hemicelluloses biosynthesis, pectin modifications by pectin methylesterases, lignin biosynthesis and changes in the abundance of hydroxyproline-rich glycoproteins (HRGP) [8].

HRGP are usually divided into three complex multigene families: (i) arabinogalactan proteins (AGP); (ii) extensins (EXT); and (iii) proline-rich proteins [9]. AGP are further divided into four sub-families according to their polypeptide core: classical AGP, lysine-rich AGP (Lys-rich AGP), arabinogalactan peptides (AG peptides) and fasciclin-like AGP (FLA) [10]. Typically, AGP are strongly O-glycosylated and most of them have glycosylphosphatidylinositol (GPI) anchors that attach the proteins to the plasma membrane, though some of them can be released into the wall matrix *via* GPI cleavage [11]. In connection with their abundance, ubiquitous presence and localisation, AGP play a crucial role in various biological processes such as cell division, cellular communication, programmed cell death, organ abscission, plant-microbe interactions, plant defence and growth as well as in the reproductive processes [12–14]. A decrease in the amount of AGP has also been linked with the loss of embryogenic potential in callus cultures of *B. distachyon* [15]. Despite many studies on the role of AGP in plant development, our understanding of their role in the reaction of the plant to temperature stress is still quite limited. Recent studies have shown that temperature stress strongly affects the distribution and content of AGP in the stigma and ovule of *Solanum lycopersicum* as well as in banana leaves and roots, which may indicate that AGP are differentially regulated in the response to temperature stress and that their expression and distribution is tissue specific [16–18].

Based on a bioinformatic analysis, EXT were divided into seven classes: classical, short, leucine-rich repeat extensins (LRX), proline-rich extensin-like receptor kinases (PERK), formin-homolog EXT (FH EXT), chimeric and long chimeric EXT. EXT are characterised by the presence of serine, which is followed by three to five proline residues. These prolines are hydroxylated and glycosylated [19]. EXT are known to play important roles in the response to wounding and pathogen infections [20]. This family was also indicated as playing an important role in root-microbe interactions [14,21]. A study on a *B. distachyon* callus showed that one of the chimeric EXT could be considered to be a good marker for embryogenic cells [15]. A chimeric leucine-rich repeat/extensin, LRX1, was shown to be required for root hair morphogenesis in *Arabidopsis thaliana* [22]. However, information on the synthesis and location of extensins in response to temperature stress is scarce.
Thus, the aim of this work was to investigate any changes in the distribution of the epitopes of AGP and EXT in *B. distachyon* leaves through an immunostaining analysis. This approach enabled the distribution of these epitopes and the changes in their leaves that had been stressed by a high or low temperature to be determined. We also determined the level of transcript accumulation of selected genes encoding EXT, EXT-like receptor kinases, and FLA in the leaves of *B. distachyon* that had been stressed by a high or low temperature using RT-qPCR.

**2. Results**

**2.1. Distribution of the Epitopes of AGP and EXT in Leaves in Response to Temperature Stress**

The distribution of the epitopes of AGP (JIM8, JIM13, JIM16, LM2 and MAC207), pectin/AGP (LM6) and EXT (JIM11, JIM12 and JIM20) in the leaves of *B. distachyon* under normal (21 °C) and thermal stress conditions (4 and 40 °C) was determined. Considering the phenotype, we observed no visible changes induced by the thermal stress. The general anatomy of a *B. distachyon* leaf is shown in Figure 1. In order to present the results clearly, only the antibodies for which changes in their localisation or the intensity of the fluorescence signal in different temperature conditions were observed, are presented in the main text. Figures A1–A5 show the localisation of the remaining epitopes, in which no changes were identified in their response to temperature treatment.

<figure><img src="image_41.png" /><figcaption>**Figure 1.** A cross-section of a *B. distachyon* leaf through the major vascular bundle (nomenclature according to Botha [23]) that had been stained with a fluorescent brightener (FB). Scale bar: 50 µm.</figcaption></figure>

The occurrence of the epitopes of AGP was mostly associated with the vascular bundle. The JIM8 epitope was present in the walls and cellular compartments of the inner bundle sheath cells and phloem (Figure 2D–F). This epitope also occurred in the sclerenchyma fibres that were located next to the vascular bundle (Figure A1A–C) or were developing at the edge of the leaf blade (Figure A1D–F). The presence and spatial distribution of the JIM8 epitope were diverse at different temperatures. The JIM8 epitope was less represented in the leaves that were growing at a low temperature (Figure 2A–C). However, in the leaves that had been subjected to a temperature of 40 °C, an increase in the intensity of fluorescence signal was observed in the walls of phloem cells compared to the leaves that were growing at a low temperature (Figure 2G–I). JIM13 was found at the same locations as the JIM8 epitope (Figure A2A–F) and was additionally detected in the intercellular compartments (under intercellular compartments we
define the localisation of epitope within the cytoplasm endomembrane system or organelles that are associated with the biosynthesis and secretion pathway to the wall, however these are not visible on the light microscope level [24]) of the prickles (Figure A2G-I). There were no changes in the distribution of the JIM13 epitope or in the intensity of the fluorescence signal between the analysed temperatures. The JIM16 epitope in the control leaves was present in a low amount in the intercellular compartments of the inner and outer bundle sheath cells and phloem as well as in the xylem parenchyma (Figure 3D-F). This epitope was not detected in the xylem parenchyma in the leaves that were growing at 40 °C (Figure 3G-I) and in the leaves from 4 °C, the presence of this epitope was not detected (apart from single dots in the vascular bundle cells; Figure 3A-C). Another AGP epitope, LM2, occurred in the cellular compartments of bundle sheath, phloem and xylem parenchyma in the control leaves (Figure 4D-F). At a low temperature (4 °C), the occurrence of this epitope was very low (Figure 4A-C), while in the leaves that were growing at a high temperature, it was more abundant (Figure 4G-I) compared to the control plants (Figure 4D-F). Additionally, LM2, was detected in the intercellular compartments and/or walls of the epidermis and bulliform cells in the leaves from plants that had been subjected to a high temperature (Figure 5A-I). A signal in the mesophyll cells was detected only in the leaves that were growing at 40 °C (Figure 5G-I). The MAC207 epitope was present in large amounts in the intercellular compartments and/or walls of the phloem cells, mesophyll cells and bulliform cells (Figure A3A-F). At all of the temperatures, its fluorescence had a similar cellular distribution and intensity. The LM6 epitope was detected abundantly in the phloem, xylem parenchyma and, in lower amounts, in the cellular compartments and/or walls of the outer bundle sheath (Figure 6D-F). The fluorescence signal of this epitope in the leaves that were growing at 40 °C was more intense compared to the other temperature treatments (Figure 6G-I vs. Figure 6A-F). Outside the vascular bundle, LM6 was present in the cell walls and/or in the intercellular compartments of the mesophyll cells (Figure A4A-C).

<figure><img src="image_42.png" /><figcaption>**Figure 2.** Immunolocalisation of the JIM8 epitope (A-I) in cross-sections of the *B. distachyon* leaves, (A-C): 4 °C; (D-F): 21 °C; (G-I): 40 °C. Abbreviations: FB—fluorescent brightener, ib—inner bundle sheath, ob—outer bundle sheath, ph—phloem, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 10 µm.</figcaption></figure>
<figure><img src="image_43.png" /><figcaption>**Figure 3.** Immunolocalisation of the JIM16 epitope (A–I) in cross-sections of the *B. distachyon* leaves, (A–I): through the major vascular bundle. (A–C): 4 °C; (D–F): 21 °C; (G–I): 40 °C. Abbreviations: FB—fluorescent brightener, ib—inner bundle sheath, ob—outer bundle sheath, ph—phloem, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 10 µm.</figcaption></figure>

<figure><img src="image_44.png" /><figcaption>**Figure 4.** Immunolocalisation of the LM2 epitope (A–I) in cross-sections of the *B. distachyon* leaves, (A–I): through the major vascular bundle. (A–C): 4 °C; (D–F): 21 °C; (G–I): 40 °C. Abbreviations: FB—fluorescent brightener, ib—inner bundle sheath, ob—outer bundle sheath, ph—phloem, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 10 µm.</figcaption></figure>
<figure><img src="image_45.png" /><figcaption>**Figure 5.** Immunolocalisation of the LM2 epitope (A–I) in cross-sections of the *B. distachyon* leaves, (A–I): through the mesophyll and bulliform cells. (A–C): 4 °C; (D–F): 21 °C; (G–I): 40 °C. Abbreviations: bc—bulliform cells, ep—epidermis, FB—fluorescent brightener, me—mesophyll, pr—prickle. The green colour shows epitope occurrence. Scale bars: 20 µm.</figcaption></figure>

<figure><img src="image_46.png" /><figcaption>**Figure 6.** Immunolocalisation of the LM6 epitope (A–I) in cross-sections of the *B. distachyon* leaves, (A–I): through the major vascular bundle. (A–C): 4 °C; (D–F): 21 °C; (G–I): 40 °C. Abbreviations: FB—fluorescent brightener, ib—inner bundle sheath, ob—outer bundle sheath, ph—phloem, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 10 µm.</figcaption></figure>
All three extensin epitopes that are recognised by the JIM11, JIM12 and JIM20 antibodies were mostly associated with the mesophyll cells. The JIM11 epitope was present only in the mesophyll cell walls (Figure A5A–C). In addition to occurring in the mesophyll cell walls (Figure A5D–F), JIM12 was also found in the walls of some of the outer bundle sheath cells and vessels (Figure A5G–I). The occurrence of the JIM20 epitope was similar to JIM12 (Figure A5J–O), but had an additional location in the walls and/or cellular compartments of the phloem (Figure A5M–O). All three extensin epitopes occurred abundantly and there were no differences in their distribution or in the intensity of the fluorescence signal among the temperatures that were analysed.

## 2.2. Analysis of the Level of Transcript Accumulation of the Genes Encoding the FLA, EXT and EXT-Like Receptor Kinases

In this study, we determined the level of transcript accumulation of five different genes encoding FLA (*Bradi4g34420*, *Bradi2g00220*, *Bradi5g18950*, *Bradi3g39740* and *Bradi2g60270*). The transcript accumulation levels of *Bradi4g34420* and *Bradi2g00220* increased in both temperatures, 4 and 40 °C, compared to the control conditions (Figure 7A). The increase in expression of the *Bradi4g34420* gene at 40 °C (4.7-fold) was higher than at 4 °C (1.9-fold) (Figure 7A). In the case of the *Bradi5g18950* gene, the expression at 4 °C was approximately the same as in the control, while its expression at 40 °C was 6-fold higher (Figure 7A). A similar pattern of expression was found for the *Bradi3g39740* gene, though there was only a slight (1.7-fold) increase in its expression at 4 °C (Figure 7B). Interestingly, there was a dramatic increase (28-fold) in the expression of this gene at 40 °C. Notably, the expression of the *Bradi2g60270* gene was only detectable in the leaves at 40 °C. Generally, temperature stress resulted in a higher level of transcript accumulation of *FLA*, though the increase was more pronounced at 40 °C.

<figure><img src="image_47.png" /><figcaption>**Figure 7.** Relative level of transcript accumulation of the fasciclin-like AGP (FLA) genes: (A) *Bradi4g34420*, *Bradi2g00220* and *Bradi5g18950* and (B) *Bradi3g39740*. The relative expression levels were normalised to an internal control (*Bradi1g32860*, gene encoding ubiquitin) and calibrated to the control. Asterisks indicate significant differences from the control using the Student's t-test (*p* < 0.05; mean ± SD, *n* = 3).</figcaption></figure>

The level of transcript accumulation of nine different genes encoding EXT and EXT-like receptor kinases were also determined. Each gene was assigned to a group based on its structure: *FH EXT* (*Bradi1g22980*, *Bradi3g59780* and *Bradi4g03720*), *chimeric EXT* (*Bradi4g11250* and *Bradi3g12902*) and *PERK EXT* (*Bradi2g00900*, *Bradi2g49240*, *Bradi1g07010* and *Bradi3g31967*). The level of transcript accumulation of the *Bradi1g22980* gene in the treated plants was not statistically different from the control (Figure 8A). The level of transcript accumulation of two other *FH EXT*, *Bradi3g59780* and *Bradi4g03720* was only statistically higher at 40 °C (Figure 8A). Conversely, the level of transcript accumulation of the *chimeric*
EXT, Bradi4g11250, increased significantly at 4 °C, though there was no clear difference in its expression for the Bradi3g12902 gene (Figure 8B). When considering PERK, a higher transcript accumulation of the Bradi2g00900 gene at 4 °C and a higher level of transcript accumulation of the Bradi2g49240 gene at 4 °C and 40 °C was determined (Figure 8C). Intriguingly, the expression of the other PERK gene, Bradi3g31967, was only observed in the temperature-stressed samples (Figure 8D). The distribution of all of the epitopes together with changes in the level of transcript accumulation of the analysed genes are summarised in Figure 9.

<figure><img src="image_48.png" /><figcaption>**Figure 8.** Relative level of transcript accumulation of the extensins (EXT) genes: (A) formin-homolog (FH) EXT: Bradi1g22980, Bradi3g59780, Bradi4g03720, (B) chimeric EXT: Bradi4g11250, Bradi3g12902, (C) proline-rich extensin-like receptor kinase (PERK): Bradi2g00900, Bradi2g49240, Bradi1g07010 and (D) Bradi3g31967. The relative expression levels were normalised to an internal control (Bradi1g32860, gene encoding ubiquitin) and calibrated to the control. Asterisks indicate significant differences from the control using the Student's t-test (*p* < 0.05; mean ± SD, *n* = 3).</figcaption></figure>

<figure><img src="image_49.png" /><figcaption>**Figure 9.** Consolidated results of the distribution of the epitopes EXT and AGP in the leaves of *B. distachyon* and changes in the level of transcript accumulation of the analysed genes.</figcaption></figure>
### 3. Discussion

Although immunohistochemical analyses are widely used to study changes in the chemical components of the cell wall during different developmental processes, in vivo and in vitro information concerning the presence and distribution of the cell wall proteins in leaves that have been subjected to biotic and/or abiotic stresses are scarce and remain largely unexplored [16,18,25]. Previous studies have primarily focused on the differential expression of AGP in response to temperatures stresses in roots and seedlings; however, the involvement of AGP in the response to temperature stress has rarely been studied in leaves [16]. For example, the transient appearance of two AGP proteins in *Triticum aestivum* in response to a low temperature were observed, thus indicating their involvement in the activation of the plant cell defence [16,26]. In transgenic *A. thaliana* plants, non-classical AGP improved the freezing tolerance of seedlings [27]. Temperature stress is one of the factors that limit plant growth and productivity [2,28]. Therefore, data showing changes in the distribution of individual cell wall components, particularly AGP, in connection with temperature stress, are particularly important as the results can be used in genetic engineering for stress tolerance [29].

In the present study, while the distribution of the epitopes of the analysed AGP and EXT was primarily observed in the major vascular bundles (nomenclature according to Botha [23]) and sclerenchyma cells, in the case of the LM2 and MAC207 epitopes, they were present in the mesophyll and epidermal cells, especially in the bulliform cells. Generally, the results for *B. distachyon* presented here are similar to those that have been described for banana leaves in terms of the distribution of epitopes in leaf tissues [18]. In banana, the JIM8 epitope increased in abundance at lower temperatures, thus indicating its role in the tolerance to strong chilling stress [18]. In the pistils of *Solanum lycopersicum* cv Micro-Tom, a high temperature strongly affected the distribution of the JIM8 epitope, which decreased in the stigma and ovule [17]. These varied results with respect to this and other epitopes mean that further intensive studies are necessary. Moreover, such results may indicate that the changes in the chemical composition of the cell walls in response to temperature stress are species-specific. In banana leaves, the presence of the JIM16 epitope was higher in a tolerant cultivar at low temperature stress, thus suggesting that these epitopes may be involved in determining the tolerance of banana to temperature stress [18]. Among the analysed epitopes, the LM2 epitope was detected in most of the leaf tissues and an increase of the LM2 epitope as a response to high temperature stress was observed. A similar distribution was detected in banana leaves, in which this epitope was found in the phloem, bundle sheath, mesophyll and epidermal cells. Low temperature treatment increased the abundance of this epitope in banana [18]. In the leaves of *Tilia x euchlora*, the LM2 epitope was present in the epidermis, hypodermis and parenchyma cells, although as a response to salt stress [25]. Such results may indicate that this wall epitope may be a marker of the plant response to diverse stresses. The abundant presence of the LM6 antibody in *B. distachyon* leaves was found. As LM6 antibody exhibits a high affinity to the (1-5)-α-L-arabinosyl residues, it detects the (1-5)-α-L-arabinan (a pectin rhamnogalacturonan I side chain), however, it can also bind to some AGP (http://www.plantprobes.net/index.php). Thus, the increase in the fluorescence signal in the xylem parenchyma of the temperature-stressed plants may not necessarily indicate changes in the presence of AGP. The function of the arabinan side chains is not well understood and their roles are postulated to be an involvement in the rehydration of the cell wall and flexibility [30–33]. The more abundant presence of the wall components that are recognised by LM6 antibody, especially in the cytoplasmic compartments, that were observed in our study indicate that leaves react to temperature stress by synthesising and depositing (1-5)-α-L-arabinans into the cell walls.

As has been shown in previous studies, analyses of the distribution and changes in the signal intensity of epitopes can be compared with the expression profiles of the genes encoding the proteins that are targeted by these antibodies [15,34]. In this work, we focused on the FLA that have been implicated in modulation of signalling upstream of cell wall polymer biosynthesis, remodelling, as well as in the stress response as one of the AGP sub-families [35–37]. We found an increase in the level of transcript accumulation of the FLA in response to temperature stress, which concurs with the
immunohistochemistry observations that have been made for the LM2 antibody. It is worth noting that the increase in the level of transcript accumulation of these genes was more pronounced at the high temperature. However, in wheat, four genes encoding *FLA* had a decreased level of transcript accumulation in response to low temperature stress [38]. Similarly, two other *FLA* genes (*OsFLA1* and *OsFLA4*) were downregulated by cold stress in rice [39]. This may reflect intrinsic differences between the analysed species or might be the result of fragmentariness of the conducted experiments, which only focus on a few of the numerous *FLA* genes that are present in a genome. The *Bradi5g18950* gene, which was analysed in our work, exhibited a higher level of transcript accumulation at the high temperature and it was previously shown to be upregulated in a 30-day-old callus that was characterised by an increased embryogenic potential. Conversely, the *Bradi3g39740* gene was linked with a gradual loss of embryogenic potential in *B. distachyon* [15]. Temperature stresses that are induced by low or high temperatures inhibit water uptake, which immediately leads to a slowing down of leaf growth. This observation was correlated with a loss of turgor in leaf cells and an adjustment of the osmoticum, which enables cells to regain turgor [40]. AGP are well known for their water-holding properties [41]. For example, in the resurrection plants, side chains of pectin are highly enriched in arabinose-rich polymers, including AGP. Their presence can prevent water loss during desiccation [8]. In *Coffea arabica* plants that had been subjected to heat stress, there were extensive changes in the cell wall of the leaves. The plants accumulated a higher content of arabinose and galactose, which may suggest that the response of coffee leaves to heat stress is related to type II arabinogalactans and pectins. Moreover, during heat stress, the palisade parenchyma cells were more separated and thinner relative to the control, which resulted in a decreased thickness of the leaves [42]. It has been hypothesised that the organs that are susceptible to water loss such as leaves increase the thickness of the cell walls, thereby limiting desiccation through the production of specific molecules such as AGP [16]. Our results seem to support this hypothesis.

Similar to the temperature stress, salt stress results in a decrease of available water due to a reduction in osmotic potential of the soil solution, which leads to a water deficit [8]. AGP have also been shown to play an important part in the salt stress response and an upregulation of *AGP* in salt-adapted tobacco BY-2 cell cultures was observed. It has been proposed that AGP act as a possible sodium carrier via vesicle trafficking from the apoplast to the vacuoles in salt-adapted tobacco BY-2 cells [43]. A significant upregulation of the *FLA* genes in the salt stress response was observed in the roots of *Populus trichocarpa* [44]. Moreover, AGP were found to act as pectin plasticisers [8,45]. As was shown for an *FLA sos5* (salt-overly sensitive) mutant of *A. thaliana*, it exhibits a root-swelling phenotype under salt stress [46]. Further studies showed that the SOS5 protein mediates adherence via its interaction with the cell wall pectin [47]. *At-FLA4* is one of the *FLA* genes in *A. thaliana* that encodes the predicted lipid-anchored glycoprotein and it was shown to positively regulate cell wall biosynthesis and root growth by modulating abscisic acid signalling. Moreover, an *At-fla4* mutant was found to be sensitive to the salt stress [37]. It has been suggested that *At-fla4* might interact with the pectin network via the covalent or non-covalent interactions of its glycans. *At-fla4* may mechanically link pectin with the *AtFei1* and *AtFei2* receptor kinases and the plasma membrane, thus contributing to some biophysical properties such as swelling and interpolymer connectivity [48]. As was indicated by another inactivated mutant of *A. thaliana*, the *FLA1* gene is involved in the early events of lateral root and shoot development in tissue cultures [49]. Considering the salt stress response, it is possible that the upregulation of *FLA* and the increased signal intensity of some of the epitopes of AGP during temperature stress may link with other cell wall polymers such as pectins and thus may modulate the signalling pathways. As has been shown by a number of studies on various species, the pectin content increases during cold stress. Conversely, it decreases during heat stress [8].

An immunohistochemical analysis of the distribution of the *EXT* using the JIM11, JIM12 and JIM20 antibodies showed the presence of all of these epitopes in the mesophyll and the JIM12 and JIM20 epitopes in the vessels. Although no immunohistochemical studies targeting the EXT using the JIM11, JIM12 and JIM20 antibodies were done in the leaves, the distribution of these epitopes has been
widely studied in the callus embryos and roots [20,50–53]. For example, changes in the signal intensity of the JIM11 and JIM12 antibodies were connected with a gradual loss of embryogenic potential in *B. distachyon* callus cultures [15]. Zhang, et al. [54] showed that the EXT that are recognised by the JIM20 antibody were present in the pollen tubes and transmitting tissue of *Nicotiana tabacum* and that the application of hydroxyproline synthesis inhibitor, 3,4-dehydro-L-proline, decreased pollen tube growth. However, studies dedicated to the role of EXT in abiotic stress and especially temperature stress are still scarce. In our study, while we did not observe any changes in the selected epitopes of the EXT in the mesophyll, outer bundle sheath, phloem or vessels at the level of the immunohistochemical analyses, we found an increase of EXT and EXT-like receptor kinase level of transcript accumulation in the response to temperature stresses, especially during the high temperature stress. This may be partially explained by the fact that we used only three antibodies that bind to the EXT, but it is possible that the application of other anti-EXT antibodies could reveal some changes. Additionally, the effectiveness of immunohistochemical analyses is limited, since it does not provide sufficient resolution and does not focus on individual genes, as is the case of RT-qPCR-based analyses. Changes in the EXT gene expression greatly depended on the class of extensins that were being analysed as was shown for *A. thaliana* plants that had been subjected to low temperature stress [55]. Another transcriptomic analysis of the *A. thaliana* response to cold stress showed the downregulation of one of the PERK genes [56]. In our experiment, we found that the PERK extensin, Bradi3g31967, was only expressed in the stressed leaves. The genes that belong to this class have been found in the apical dominance, floral organ defects and root cell elongation [57]. An increased accumulation level of the PERK4 gene transcript in *A. thaliana* was observed in response to abscisic acid, which is a key regulator of abiotic stress tolerance in plants [58,59]. PERK1 mRNA from Brassica napus was shown to be dramatically and rapidly accumulated in response to wounding and moderately accumulated in response to infection by the fungal pathogen Sclerotinia sclerotiorum [60]. Interestingly, the LRX proteins were found to regulate salt tolerance in *A. thaliana*. A triple mutant in the LRX genes exhibited a severe salt hypersensitivity and these genes were determined to be an important sensor of the cell wall integrity signals [61]. Moreover, recent studies have hinted at the role of EXT in the plant defence against phytopathogens as well as in interactions with beneficial microorganisms [14,21,62]. EXT have also been implicated in aluminium resistance and its accumulation in the cell walls of pea roots was observed [63]. The changes in the plant cell wall in response to temperature stress are diverse and not only include AGP and EXT, but also alterations in cellulose, hemicellulose, pectin and lignin biosynthesis [8]. Further investigations into the changes in cell wall proteomes could unravel the involvement of other proteins in the stress response because the proteome of the *B. distachyon* cell walls is complex and consists of at least 594 proteins [64–68].

# 4. Materials and Methods

## 4.1. Plant Material

The plants of the *B. distachyon* reference genotype Bd21 that were used in this experiment were cultivated in pots that had been filled with soil mixed with vermiculite at a ratio of 3:1 in a greenhouse. The seeds of *B. distachyon* genotype Bd21 (accession number: PI 254867) were sourced from the collection held by the United States Department of Agriculture—National Plant Germplasm System. The plants were grown in the greenhouse under a 16 h/8 h light/dark photoperiod at 21 ± 1 °C and were illuminated by lamps emitting white light at an intensity of 10 000 lx. For the low temperature stress, the plants were incubated at 4 °C for 24 h and for the high temperature stress, the plants were incubated at 40 °C for 24 h in growing chambers [69]. Plants at the fourth stage of principal growth according to the Hong, et al. [70] were used in this experiment. This stage is referred to as booting and is characterised by the emergence of the head at the top of the growing shoot. The flag leaf was harvested and used to isolate the RNA and to perform the RT-qPCR analysis. For the immunohistochemistry
analysis, the middle part of the leaf was collected because this permitted clear observations of the major vascular bundle, epidermis, bulliform cells, mesophyll and sclerenchyma.

## 4.2. Sample Preparation

To determine the chemistry of the cell wall, a set of monoclonal antibodies against the specific cell wall epitopes of the AGP (antibodies JIM8, JIM13, JIM16, LM2, MAC207), pectin/AGP (LM6) and EXT (JIM11, JIM12 and JIM20) (Plant Probes, Leeds, UK) were used. The references and information on the antibodies are shown in Table 1. The leaves were excised, fixed and embedded in Steedman's wax [20,71]. Transverse sections of the leaf blade (7 µm thick) were cut using a HYRAX M40 rotary microtome (Zeiss, Oberkochen, Germany) and collected on microscopic slides coated with poly-L-lysine (Menzel Gläser, Braunscheig, Germany).

## 4.3. Immunohistochemistry

The sections were de-waxed and rehydrated in an ethanol series (three times in 100, 90 and 50% ethanol in phosphate buffered saline PBS, v/v, each for 10 min) and PBS (10 min) [71]. The detailed procedure for immunohistochemical analysis and histological section observation was as previously described [20]. The slides were stained with 0.01% (w/v) fluorescent brightener 28 (FB) (Sigma-Aldrich, St. Louis, MO, USA) in PBS, which was used to visualise cell walls due to its affinity to cellulose. Two biological replicates were performed with at least eight sections for each replicate.

## 4.4. RT-qPCR

In order to characterise the level of transcript accumulation of the selected genes, RT-qPCR was performed using a LightCycler® 480 SYBR Green I Master in a LightCycler® 480 Real-Time PCR System (Roche, Basel, Switzerland). The total RNA was isolated from the leaves of *B. distachyon*. The primers used in this research are shown in Table A1. The genes encoding extensins with their division into classes were as previously described [19]. The *FLA* genes were selected based on the annotation found in the Phytozome database (https://phytozome.jgi.doe.gov/pz/portal.html). The detailed procedure for RT-qPCR was as in Betekhtin, et al. [72]. Briefly, the isolated RNA were treated with the DNase (QIAGEN, Hilden, Germany), and subsequently used for first-strand cDNA generation. Samples were run in the LightCycler® 480 Real-Time PCR System (Roche, Basel, Switzerland). The PCR conditions were as follow: 5 min at 95 °C, 45 cycles of 10 s at 95 °C, 20 s at 60 °C and 10 s at 72 °C with signal acquisition. Ubiquitin was used as the reference gene and analysis was performed using the 2⁻ΔΔCT method. The significant differences between the samples and control were calculated using the Student's t-test.

<table><caption>**Table 1.** The antibodies that were used for the immunocytochemistry, the epitopes they recognise and the relevant references.</caption><thead><tr><th>Antibody</th><th>Epitope</th><th>References</th></tr></thead><tbody><tr><td colspan="3"><strong>AGP</strong></td></tr><tr><td>JIM8</td><td>Arabinogalactan</td><td>[73]</td></tr><tr><td>JIM13</td><td>(β)GlcA1-&gt;3(α)GalA1-&gt;2Rha</td><td>[74-76]</td></tr><tr><td>JIM16</td><td>AGP glycan</td><td>[74-76]</td></tr><tr><td>LM2</td><td>β-linked GlcA</td><td>[75,77]</td></tr><tr><td>MAC207</td><td>(β)GlcA1-&gt;3(α)GalA1-&gt;2Rha</td><td>[74,75,78,79]</td></tr><tr><td colspan="3"><strong>Pectin/AGP</strong></td></tr><tr><td>LM6</td><td>(1-5)-α-L-arabinosyl residues, can also bind to some AGP</td><td>[80,81]</td></tr><tr><td colspan="3"><strong>EXT</strong></td></tr><tr><td>JIM11</td><td>Extensin</td><td>[74,82]</td></tr><tr><td>JIM12</td><td>Extensin</td><td>[82]</td></tr><tr><td>JIM20</td><td>Extensin</td><td>[82]</td></tr></tbody></table>
## 5. Conclusions

In our work, we demonstrated changes in the abundance and diversified expression of the epitopes of the AGP genes encoding FLA, EXT and EXT-like receptor kinases in the leaves of *B. distachyon* in response to temperature stress.

The main findings are as follows:

1. An increase in the JIM8 signal in the walls of phloem cells at 40 °C and a decrease at 4 °C.

2. An increase in the abundance of the LM2 epitope in the leaves of plants that had been subjected to the high temperature.

3. A decrease in the JIM16 signal intensity at 4 and 40 °C.

4. The upregulation of some *FLA*, *EXT* and *EXT-like* receptor kinases genes in response to temperature stress (4 and 40 °C).

5. The expression of the *PERK EXT* gene *Bradi3g31967* only in the leaves under low and high temperature stress.

To summarise, our results extend the knowledge about the presence of these epitopes in connection with temperature stress in *B. distachyon*. A precise dissection of the functions of AGP and EXT in response to abiotic stresses and to temperature stress, among others, requires the use of specific mutants whose availability is still limited. However, recent developments in the site-directed mutagenesis techniques such as CRISPR/Cas9 should allow the selective targeting of these genes, which may be helpful to better understand their roles.

**Author Contributions:** Conceptualisation, A.P., A.B.; methodology, A.P., A.B., K.S., K.G.-J.; validation, A.P., K.S., A.B.; formal analysis, A.P., K.S., A.B.; investigation, A.P., K.S., A.B.; resources, R.H., E.K.; data curation, A.P., A.B., K.S.; writing—original draft preparation, A.P., A.B., K.S., E.K.; writing—review and editing, A.P., A.B., K.S., E.K., R.H.; visualisation, A.P.; supervision, A.B., R.H.; funding acquisition, R.H., E.K.

**Funding:** This research was funded by the National Science Centre Poland (grant DEC-2014/14/M/NZ2/00519).

**Conflicts of Interest:** The authors declare no conflict of interest.

## Abbreviations

<table><tr><td>AG Peptides</td><td>Arabinogalactan peptides</td></tr><tr><td>AGP</td><td>Arabinogalactan proteins</td></tr><tr><td>EXT</td><td>Extensins</td></tr><tr><td>FH EXT</td><td>Formin-homolog EXT</td></tr><tr><td>FLA</td><td>Fasciclin-like AGP</td></tr><tr><td>GPI</td><td>Glycosylphosphatidylinositol</td></tr><tr><td>HRGP</td><td>Hydroxyproline-rich glycoproteins</td></tr><tr><td>LRX</td><td>Leucine-rich repeat extensins</td></tr><tr><td>Lys-rich AGP</td><td>Lysine-rich arabinogalactan proteins</td></tr><tr><td>PBS</td><td>Phosphate-buffered saline</td></tr><tr><td>PERK</td><td>Proline-rich extensin-like receptor kinases</td></tr><tr><td>ROS</td><td>Reactive oxygen species</td></tr><tr><td>RT-qPCR</td><td>Reverse transcription-quantitative polymerase chain reaction</td></tr></table>
Appendix A

<figure><img src="image_50.png" /></figure>

**Figure A1.** Immunolocalisation of the JIM8 epitope (A–F) in cross-sections of *B. distachyon* leaves (21 °C), (A–C): through the major vascular bundle; (D–F): through the edge of the leaf blade. The white arrows point to the respective parts of the leaf. Abbreviations: ep—epidermis, FB—fluorescent brightener, ib—inner bundle sheath, me—mesophyll, ob—outer bundle sheath, ph—phloem, pr—prickle, sc—sclerenchyma, vb—vascular bundle, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 20 µm.

<figure><img src="image_51.png" /></figure>

**Figure A2.** Immunolocalisation of the JIM13 epitope (A–I) in cross-sections of *B. distachyon* leaves (21 °C), (A–C): through the major vascular bundle; (D–F): through the edge of the leaf blade; (G–I): prickle. The white arrows point to the respective parts of the leaf. Abbreviations: bc—bulliform cells, ep—epidermis, FB—fluorescent brightener, ib—inner bundle sheath, me—mesophyll, ob—outer bundle sheath, ph—phloem, pr—prickle, sc—sclerenchyma, vb—vascular bundle, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 20 µm.
<figure><img src="image_52.png" /><figcaption>**Figure A3.** Immunolocalisation of the MAC207 epitope in cross-sections of *B. distachyon* leaves (21 °C), (A–C): through the major vascular bundle; (D–F): through the mesophyll and bulliform cells. Abbreviations: ep—epidermis, b—bulliform cells, FB—fluorescent brightener, ib—inner bundle sheath, me—mesophyll, ob—outer bundle sheath, ph—phloem, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 20 µm.</figcaption></figure>

<figure><img src="image_53.png" /><figcaption>**Figure A4.** Immunolocalisation of the LM6 epitope (A–C) in cross-sections of *B. distachyon* leaves (21 °C), through the major vascular bundle. The white arrows point to the respective parts of the leaf. Abbreviations: ep—epidermis, FB—fluorescent brightener, ib—inner bundle sheath, me—mesophyll, ob—outer bundle sheath, ph—phloem, sc—sclerenchyma ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 20 µm.</figcaption></figure>
<figure><img src="image_54.png" /><figcaption>**Figure A5.** Immunolocalisation of the JIM11 (A–I), JIM12 (D–I) and JIM20 (J–O) epitopes in cross-sections of *B. distachyon* leaves (21 °C), (A–F,J–L): through the edge of the leaf blade; (G–I,M–O): through the major vascular bundle. Abbreviations: bc—bulliform cells, ep—epidermis, FB—fluorescent brightener, ib—inner bundle sheath, me—mesophyll, ob—outer bundle sheath, p—phloem, pr—prickle, sc—sclerenchyma, vb—vascular bundle, ve—vessels, xp—xylem parenchyma. The green colour shows epitope occurrence. Scale bars: 20 µm.</figcaption></figure>
<table><caption>**Table A1.** The oligonucleotide primers that were used for the RT-qPCR reaction with relevant descriptions of the genes.</caption><thead><tr><th>Genes</th><th>Description of the Genes</th><th>Primer Sequence (5'-3')</th></tr></thead><tbody><tr><td>Bradi1g32860</td><td>ubiquitin</td><td>pF-GAGGGTGGACTCCTTTGGA<br>pR-TCCACACTCCACTTGGTGCT</td></tr><tr><td colspan="3"><strong>EXT and EXT-like receptor kinase</strong></td></tr><tr><td>Bradi4g11250</td><td>extensin (chimeric EXT)</td><td>pF-GCGACTGCGACAATGATGTG<br>pR-ACCCCTTGCTAAGCCCTCTA</td></tr><tr><td>Bradi3g12902</td><td>extensin (chimeric EXT)</td><td>pF-CATCTGGACCTGCCAATGG<br>pR-TCCCAGTTTTGGAGTCTCGC</td></tr><tr><td>Bradi3g59780</td><td>formin-homolog extensin (FH EXT)</td><td>pF-GATGAATGCCGGAACAGCAC<br>pR-GTGGAGAAGAGTGGTGCCCTC</td></tr><tr><td>Bradi4g03720</td><td>formin-homolog extensin (FH EXT)</td><td>pF-GAAGCAGATTGAGGCCGAGA<br>pR-CGCGCCTCCATCTTTTGATT</td></tr><tr><td>Bradi1g22980</td><td>formin-homolog extensin (FH EXT)</td><td>pF-CAGCAGAGCCTGTTGCTTGAC<br>pR-TTCTAGGTTTCCGTGCATGAGT</td></tr><tr><td>Bradi2g49240</td><td>proline-rich extensin-like receptor kinase<br>(PERK)</td><td>pF-TTCTCAGCCGTTGGGAGATG<br>pR-GGAAGGTCCCCAAAGTCTCG</td></tr><tr><td>Bradi1g07010</td><td>proline-rich extensin-like receptor kinase<br>(PERK)</td><td>pF-CCTCCACCGTAAAGGGCTG<br>pR-GATCCGTGGATGGCAGTCTT</td></tr><tr><td>Bradi3g31967</td><td>proline-rich extensin-like receptor kinase<br>(PERK)</td><td>pF-CCGTCGCCATTAAGAATCTGC<br>pR-GATTCTTGTGCCGAACTCGC</td></tr><tr><td>Bradi2g00900</td><td>proline-rich extensin-like receptor kinase<br>(PERK)</td><td>pF-TAACTTTGAGGCACAGGTTGCT<br>pR-AGCCATGTATCCAAAAGTCCCC</td></tr><tr><td colspan="3"><strong>FLA</strong></td></tr><tr><td>Bradi2g00220</td><td>fasciclin-like arabinogalactan protein</td><td>pF-AGCTCAACAGCTCCCAGAC<br>pR-CGAAAGCGAGTTGAGCGTG</td></tr><tr><td>Bradi5g18950</td><td>fasciclin-like arabinogalactan protein</td><td>pF-AATAAGGGAAGTCACCGTCGC<br>pR-CCGTTCTTCTTGTCATGGACCT</td></tr><tr><td>Bradi4g34420</td><td>fasciclin-like arabinogalactan protein</td><td>pF-CACATCCTCCAGATGCACGTC<br>pR-CCGGACTCCTGGAACATGG</td></tr><tr><td>Bradi3g39740</td><td>fasciclin-like arabinogalactan protein</td><td>pF-GTACTATTCCCTGGCGGAGTTC<br>pR-CCATGTTGTCGGTGAGGTTGAG</td></tr><tr><td>Bradi2g60270</td><td>fasciclin-like arabinogalactan protein</td><td>pF-AGCAGAGCAATCCTCTAGTAGC<br>pR-TGGGTTCTTCTCGCCATTGTTA</td></tr></tbody></table>

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