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

*Copyright © 2019 Buet, Costa, Martínez and Guiamet. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Clathrin-Mediated Endocytosis Delivers Proteolytically Active Phytaspases Into Plant Cells

*Svetlana V. Trusova1 , Anastasia D. Teplova2 , Sergei A. Golyshev1 , Raisa A. Galiullina1 , Ekaterina A. Morozova2 , Nina V. Chichkova1 and Andrey B. Vartapetian1 \**

*1 Department of Chemistry and Biochemistry of Nucleoproteins, Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow, Russia, 2 Faculty of Bioengineering and Bioinformatics, Moscow State University, Moscow, Russia*

Phytaspases belong to the family of plant subtilisin-like proteases and are distinct from other family members, as they have strict and rarely occurring aspartate cleavage specificity and unusual localization dynamics. After being secreted into the apoplast of healthy plant tissues, phytaspases are able to return back into cells that have been committed to cell death due to a variety of biotic and abiotic stresses. It was recently discovered that retrograde transport of phytaspases involves clathrin-mediated endocytosis. Here, consequences of phytaspase internalization were studied. Proteolytic activity of phytaspases in the apoplast and intracellular protein fractions obtained from *Nicotiana benthamiana* leaves containing either endogenous phytaspase only or transiently producing *Nicotiana tabacum* phytaspase-EGFP protein (*Nt*Phyt-EGFP) was determined. We demonstrated that triggering phytaspase internalization by antimycin A-induced oxidative stress is accompanied by re-distribution of phytaspase activity from the apoplast to the cell interior. Inhibition of clathrin-mediated endocytosis by co-production of the Hub protein prevented phytaspase internalization and phytaspase activity re-localization. Specificity of endocytic uptake of phytaspases was demonstrated by the co-production of an apoplast-targeted mRFP protein marker, which retained its apoplastic localization when phytaspase internalization was essentially complete. Overproduction of *Nt*Phyt-EGFP, but not of the proteolytically inactive phytaspase mutant, *per se* caused moderate damage in young *Nicotiana benthamiana* seedlings, whereas antimycin A treatment induced a pronounced loss of cell viability independent of the *Nt*Phyt-EGFP overproduction. Interestingly, inhibition of clathrin-mediated endocytosis abrogated cell death symptoms in both cases. In contrast to stress-induced internalization of tobacco phytaspase, *Arabidopsis thaliana* phytaspase-EGFP protein (*At*Phyt-EGFP) was spontaneously internalized when transiently produced in *N. benthamiana* leaves. The *At*Phyt-EGFP uptake was dependent on clathrin-mediated endocytosis as well, the internalized protein being initially visualized within the membranous vesicles. At later time points, the EGFP tag was cleaved off from *At*Phyt, though the elevated level of intracellular *At*Phyt proteolytic activity persisted. Our data, therefore, point to clathrinmediated endocytosis as a means to deliver proteolytically active phytaspases into plant cells. It would be interesting to learn whether or not phytaspases are unique among the large family of plant subtilisin-like proteases in their ability to utilize retrograde trafficking.

Keywords: plant cell death, clathrin-mediated endocytosis, protein localization, proteolytic activity, subtilisin-like protease, phytaspase

### *Edited by:*

*Mercedes Diaz-Mendoza, Instituto Nacional de Investigación y Tecnología Agraria y Alimentaria, Spain*

### *Reviewed by:*

*Renier A. L. Van Der Hoorn, University of Oxford, United Kingdom Juan Guiamet, National University of La Plata, Argentina Johana Catherine Misas Villamil, University of Cologne, Germany*

*\*Correspondence:*

*Andrey B. Vartapetian<EMAIL_ADDRESS>
### *Specialty section:*

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

*Received: 09 February 2019 Accepted: 19 June 2019 Published: 18 July 2019*

### *Citation:*

*Trusova SV, Teplova AD, Golyshev SA, Galiullina RA, Morozova EA, Chichkova NV and Vartapetian AB (2019) Clathrin-Mediated Endocytosis Delivers Proteolytically Active Phytaspases Into Plant Cells. Front. Plant Sci. 10:873. doi: 10.3389/fpls.2019.00873*

# INTRODUCTION

Phytaspases belong to the vast family of plant subtilisin-like proteases (subtilases), which includes many members in each plant species, e.g., 56 in *Arabidopsis thaliana* (Rautengarten et al., 2005), 63 in rice (*Oryza sativa*, Tripathi and Sowdhamini, 2006), and 82 in grape (*Vitis vinifera*, Cao et al., 2014; Figueiredo et al., 2016) and tomato (*Solanum lycopersicum*, Reichardt et al., 2018). Subtilases are known to be involved in diverse processes, from unselective protein degradation (Yamagata et al., 1994; Hamilton et al., 2003) to precise processing of precursor proteins (Liu et al., 2007; Liu and Howell, 2010; Sénéchal et al., 2014; Ghorbani et al., 2016; Schardon et al., 2016; Beloshistov et al., 2018). However, function of the majority of these proteolytic enzymes remains unknown. Similar to other plant subtilases, phytaspases are synthesized as proteolytically inactive precursor proteins, which possess an N-terminal signal peptide, a prodomain, and a peptidase domain (Chichkova et al., 2010; Schaller et al., 2018). The precursor protein is autocatalytically and constitutively processed, and the mature proteolytically active enzyme is released into the apoplast (Chichkova et al., 2010), which is also typical for plant subtilases. However, phytaspases differ from other subtilases in two ways. First, phytaspases display strict aspartate (Asp) specificity of hydrolysis. The efficiency of hydrolysis after Asp residue strongly depends on the preceding three amino acid-long motif, which confers strong selectivity of phytaspase-mediated protein fragmentation. For native rice phytaspase, the preferred upstream recognition motif is remarkably hydrophobic (Galiullina et al., 2015). Computerbased modeling of the *Nicotiana tabacum* phytaspase with its peptide inhibitor provided an explanation for Asp specificity of phytaspases (Vartapetian et al., 2011). A direct consequence of Asp specificity is observed at the prodomain-peptidase domain junction in the phytaspase precursor protein. The C-terminal residue of prodomain is Asp, which is consistent with the self-processing mode of generation of the mature enzyme. Mutating this junction Asp residue precludes processing/ activation of the phytaspase precursor and the release of the mature enzyme into the apoplast (Chichkova et al., 2010). The presence of junction Asp residue may serve as a phytaspase signature within the plant subtilase family, and this sign has been successfully used to identify phytaspase-encoding genes in several plant species. The number of phytaspase genes appears to vary in plant genomes, from a single gene in *A. thaliana* to 12 in *S. lycopersicum* (Chichkova et al., 2018; Reichardt et al., 2018).

The second distinctive characteristic of phytaspases is their dynamic localization. The apoplast is not the end point of phytaspase trafficking. Phytaspases are known to be crucial to the implementation of programmed cell death (PCD) in plants triggered by biotic and abiotic stresses. Increased phytaspase levels were shown to enhance stress-induced and spontaneous plant cell death, whereas down-regulation of phytaspase activity suppressed cell death (Chichkova et al., 2004, 2010; Reichardt et al., 2018). Upon the induction of cell death, phytaspases become physically re-localized from the apoplast toward the cell interior (Chichkova et al., 2010, 2012). This retrograde transport is considered to be unique among plant subtilisin-like proteases. The involvement of clathrin-mediated endocytosis in phytaspase internalization has recently been documented (Trusova et al., 2019), which provokes important questions regarding possible mechanisms and consequences of retrograde phytaspase trafficking.

Here, we examined possible correlations between clathrinmediated phytaspase re-entry into plant cells and proteolytic activity of phytaspases both outside and inside plant cells. We found that clathrin-mediated endocytosis provides a gateway for delivery of proteolytically active phytaspases into plant cells. Also, results from this study suggest that a specific recognition mechanism for phytaspase internalization may exist. Finally, our study points to the importance of clathrinmediated endocytosis for the accomplishment of antimycin A-induced and phytaspase overexpression-promoted plant cell death.

# MATERIALS AND METHODS

### Plant Growth Conditions

*Nicotiana benthamiana* plants were grown at 25°C in soil in a controlled environment under a 16 h/8 h day/night cycle. Protein transient expression was performed using 6-week-old plants. For evaluation of cell death symptoms, *Nicotiana benthamiana* seedlings were grown on the half-strength Murashige and Skoog medium (pH 5.7) containing 1% glucose and 0.8% agar.

# Plasmid Construction

For construction of recombinant *Nt*Phyt-EGFP, *Nt*Phyt-S537A-EGFP, *Nt*Phyt-mRFP, *At*Phyt-EGFP, and *At*Phyt-S553A-EGFP fusion proteins, the downstream GST tag gene in the previously described Phyt-GST constructs within the pLH7000 binary vector backbone (Chichkova et al., 2010, 2018) was substituted with the PCR-amplified EGFP or mRFP gene. To create the SP-mRFP fusion protein, the signal peptide-encoding region of the *Nt*Phyt cDNA was PCR-amplified and ligated upstream of and in frame with the mRFP gene between the *Nco*I and *Sac*I sites of the pLEX7000 expression vector (Beloshistov et al., 2018). The mRFP gene alone was inserted in the same vector in an analogous fashion to serve as a control. To obtain the mRFP-Hub1 fusion protein, a pCambia1300-derived expression vector pCambia1300EX was constructed by inserting the 1,200 bp long *Sal*I-*Eco*RI DNA fragment of pLEX7000 encompassing the dual 35S promoter, a polylinker and a transcription terminator between the *Sal*I and *Eco*RI sites of the pCambia1300 binary vector. cDNA encoding the C-terminal fragment of *A. thaliana* clathrin heavy chain 1was PCR-amplified using primers 5′-CCAGGATCCAAGAAGTTTAACTTAAATGTTCAG-3′ and 5′-GTTGGTACCTTAGTAGCCGCCCATCGGTG-3′. Hub1 cDNA (*c*. 1,870 bp long) was then inserted downstream of and in frame with the mRFP gene between the *Nco*I and *Kpn*I sites of the pCambia1300EX binary vector. The EGFP-LTI6b-encoding plasmid was a gift from M. Taliansky (The James Hutton Institute, UK).

### Agroinfiltration and Protein Fractionation

The obtained plasmid constructs were introduced into *Agrobacterium tumefaciens* C58C1 or GV3101 cells. Transformed agrobacteria were infiltrated into *N. benthamiana* leaves using a blunt syringe or vacuum-infiltrated into seedlings (see below) in combinations described in legends (**Figures 1–7**). Agrobacteria carrying the empty vector (pCambia1300 or pLH7000) were used as a control, and were also added to the infiltration mix in the case of co-expression experiments to equalize plasmid ratio and bacterial load. At the indicated days post-infiltration (p.i.), leaves were examined by confocal fluorescence microscopy. Where indicated, treatment of leaves with antimycin A was performed by vacuum infiltration with water containing 10 μM antimycin A (Sigma, from 20 mM stock solution in ethanol). Control leaves were infiltrated with distilled water supplemented with an equivalent amount of ethanol. After an overnight incubation, confocal microscopy images were taken to determine fluorescence distribution in leaf tissue.

In parallel, leaves were subject to protein fractionation for subsequent determination of phytaspase activity. Apoplastic washes were obtained by low-speed (2,000 *g*) centrifugation of leaves at 4°C for 10 min. The leaf material was re-extracted with 20 mM MES buffer, pH 5.5, containing 100 mM NaCl and 25 μg/ml AEBSF, 2 μg/ml aprotinin, 5 μg/ml E-64 (all from Sigma), and 6 μg/ml leupeptin (MP Biomedicals) protease inhibitors, by vacuum infiltration and centrifugation, and the apoplastic washes were combined. After the separation, the residual leaf material was frozen in liquid nitrogen and disrupted in Minilys homogenizer (Bertin Instruments) using 1.6 mm ceramic beads by two 10 s bursts. An additional 10 s burst was performed after suspending the samples in B1 buffer (20 mM MES, 2 mM dithiothreitol, 0.1% Tween 20, 5% glycerol), pH 5.5, containing 50 mM NaCl and protease inhibitors (225 μl of the buffer per 25 mg of leaves). Debris was eliminated by 10 min centrifugation at 10,000 *g* at 4°C, and the supernatants (as well as the apoplastic washes) were taken for phytaspase activity determination. Leaves without prior separation of the apoplastic liquid were processed in an analogous fashion to obtain "total protein" samples. For western blot analysis and for glucose 6-phosphate dehydrogenase (G6PDH) activity determination, leaves were infiltrated with 20 mM MES buffer, pH 5.5, containing 100 mM NaCl and protease inhibitors. After separating the apoplastic protein fraction, the remaining leaf material (as well as the total/unwashed tissue) was ground in liquid nitrogen and extracted with 10 mM Tris-HCl buffer, pH 9.0, containing 0.2 M KCl, 30 mM MgCl2, 0.2 M sucrose, 10 mM 2-mercaptoethanol, and protease inhibitors. Western blot analysis of the EGFP- and mRFP-fused proteins was performed using monoclonal anti-EGFP 3A9 antibody (Sukhacheva et al., 2002) and polyclonal anti-mRFP antibodies (Abcam) as described (Chichkova et al., 2010).

For G6PDH activity determination (Yang et al., 2019), the intracellular protein and apoplastic fractions obtained from approximately 12 mg of leaf tissues were diluted 10- to 100-fold with 33 mM HEPES buffer, pH 7.5, containing 5 mM MgCl2, and 5 mM glucose 6-phosphate. As the protein fractions were obtained in different buffers, to equalize buffer conditions, the reaction mixtures were supplemented with the corresponding amounts of the complementary buffer prior to starting the reaction by the addition of NADP to a final concentration of 0.5 mM. G6PDH activity was determined spectrophotometrically, by measuring change in absorbance at 340 nm.

### Phytaspase Activity Quantification

Proteolytic activity of phytaspases in apoplastic washes, intracellular protein fractions, and in total *N. benthamiana* leaf extracts was determined using either the Ac-VEID-AFC [AFC, 7-amino-4-(trifluoromethyl) coumarin] fluorogenic peptide substrate (for *Nt*Phyt and for endogenous *N. benthamiana* phytaspase), or for *At*Phyt, the Ac-YVAD-AFC substrate (both from California Peptide). Fluorogenic peptide substrates Ac-VAD-AFC, Ac-VDVAD-AFC, Ac-LEHD-AFC, Ac-WEHD-AFC, Ac-DEVD-AFC (all from Calbiochem), Ac-VNLD-AFC (California Peptide), Ac-STATD-AFC (Bachem), and Ac-IETD-AFC (Anaspec) were used to assess cleavage specificity. Peptide substrates were used at a final concentration of 30 μM. Protein samples were 10- to 15-fold diluted before activity measurements. Kinetic measurements of relative fluorescence increase were performed in B1 buffer, pH 5.5 (for tobacco phytaspases) or pH 6.5 (for *At*Phyt), containing 0.5 M NaCl and protease inhibitors at 28°C. Where indicated, N-ethylmaleimide (Sigma) was added to the reaction mixtures to a final concentration of 2 mM. The Fluoroskan Ascent reader (Thermo Fisher Scientific) equipped with 390 nm excitation and 510 emission filters was used to quantitate fluorescence intensities. Data are presented as means ± SD from three independent experiments. Statistical significance was analyzed using Student's *t*-test. *p* < 0.01 were considered significant.

### Treatment of *N. benthamiana* Seedlings

Seven-day-old sterile *N. benthamiana* seedlings grown in solid growth medium (2.17 g/L Murashige and Skoog basal salt mixture, 0.5 g/L MES, 10 g/L glucose, 0.8% agar, pH 5.7) were vacuum infiltrated with transformed agrobacteria. Three days after infiltration, seedlings were detached and submerged in water containing 10 μM antimycin A for 3 h. Control seedlings were submerged in water supplemented with an equivalent amount of ethanol. After that, seedlings were stained with Evans Blue (Sigma) according to Minina et al., 2013. Briefly, detached seedlings were vacuum infiltrated with fresh aqueous solution of 0.5% Evans Blue and incubated for 15 min at room temperature, then washed three times for 15 min with water, slightly shaking to remove unbound dye. Chlorophyll was removed by incubating samples in 96% ethanol for 10 min at room temperature twice, then ethanol was removed by washing with 70, 50, 20% ethanol and water, consequently.

### Confocal Fluorescence Microscopy

Samples were studied using Nikon C2+ confocal microscope based on Nikon Eclipse Ti (Nikon, Japan) inverted body equipped with 60× NA 1.2 plan-apochromat water immersion lens with the working distance of 300 μm and 402, 488, and 562 nm lasers used for excitation of DAPI, EGFP, and mRFP fluorescence, respectively. Fluorescence was detected using 560 nm dichroic mirror and 525/50 and 595/40 nm blocking filters for EGFP and mRFP-tagged proteins, respectively. FM4-64 fluorescence was also recorded using 595/40 nm filter. The same pinhole diameter setting of 60 μm was used for both channels, resulting in optical sectioning of 650 nm for green and red channels. Deconvolution was performed using Richardson-Lucy algorithm implemented in the microscope controlling software Nis Elements AR (Nikon, Japan). FM4-64 (Molecular Probes) was infiltrated into leaves at a 5 μM concentration in water. Images were taken 4 h p.i. Data were reproducible over at least three independent experiments.

# RESULTS

### Stress-Induced Internalization of *N. tabacum* Phytaspase Depends on Clathrin-Mediated Endocytosis and Is Specific

*N. tabacum* phytaspase (*Nt*Phyt) is known to accumulate in the apoplast of healthy leaves and to re-localize toward the cell interior in response to PCD-inducing triggers (Chichkova et al., 2010). To follow *Nt*Phyt trafficking, the *Nt*Phyt-EGFP protein was transiently produced in *N. benthamiana* leaves by agroinfiltration. Fluorescence microscopy examination of the infiltrated leaves confirmed the production of the target protein, with no fluorescence occurring in the cell interior (**Figure 1A**). Triggering oxidative stress-induced cell death in these leaves by treatment with 10 μM antimycin A resulted in the re-distribution of *Nt*Phyt-EGFP, visible as the formation of multiple small dots within the cell (**Figure 1B**). Western blot analysis of protein samples from control (mock-treated) and antimycin A-treated leaves did not reveal degradation of the *Nt*Phyt-EGFP protein in response to antimycin A treatment (**Figure 1C**). To verify the initial apoplastic localization of *Nt*Phyt, a plasma membrane protein marker EGFP-LTI6b (Cutler et al., 2000) was transiently produced together with *Nt*PhytmRFP in *N. benthamiana* leaves (**Figure 1D**). In plasmolysed leaf samples, the *Nt*Phyt-mRFP red fluorescence was visualized in the space between two plasma membranes (green) of the adjacent epidermal cells (**Figures 1E–G**). This observation was also supported by measuring fluorescence intensities across the boundaries of the two adjacent plant cells (**Figure 1H**). The *Nt*Phyt-mRFP protein responded to treatment with antimycin A similarly to the *Nt*Phyt-EGFP fusion, i.e., by forming small dots within the cell (**Figures 1I,J**).

To verify the involvement of clathrin-mediated endocytosis in the observed antimycin A-induced re-localization of *Nt*Phyt, mRFP-Hub1 was produced together with *Nt*Phyt-EGFP in *N. benthamiana* leaves. Hub1 represents the C-terminal fragment of *A. thaliana* clathrin heavy chain 1, which acts, upon over-production in *A. thaliana* and *N. benthamiana*, in a dominant negative fashion to specifically inhibit clathrin-mediated endocytosis (Liu et al., 1995; Dhonukshe et al., 2007; Kitakura et al., 2011; Li and Pan, 2017). In the presence of mRFP-Hub1, no stress-induced re-localization of *Nt*Phyt-EGFP was observed (**Figures 1K–P**). Thus, similar to results from previous studies (Chichkova et al., 2010; Trusova et al., 2019), oxidative stressinduced PCD caused retrograde transport of *Nt*Phyt-EGFP from the apoplast to the cell interior, which could be efficiently blocked by inhibiting clathrin-mediated endocytosis.

One could imagine that internalization of *Nt*Phyt by means of clathrin-mediated endocytosis could be achieved either through specific recognition of the cargo or non-specific capturing of the apoplastic fluid by the newly forming clathrin-coated pits. Availability of an inert soluble apoplastic protein marker would be helpful to distinguish between these possibilities. To construct such a marker, we fused the *Nt*Phyt signal peptide (SP, amino acid residues 1–24) to the N-terminus of the mRFP protein to drive secretion of the fluorescent protein into the apoplast and produced the resultant SP-mRFP protein in *N. benthamiana* leaves by agroinfiltration. **Figure 2A** shows that mRFP fluorescence was detectable at the cell borders in the infiltrated leaves. After separating the proteins into apoplastic and intracellular fractions, mRFP was visualized as an apoplastic protein by western blot analysis (**Figure 2B**). This was in sharp contrast to the behavior of free mRFP (without signal peptide) produced in *N. benthamiana* leaves. While mRFP fluorescence was located to the cell periphery and the nucleus (**Figure 2C**), separating the proteins into apoplastic and intracellular fractions expectedly revealed free mRFP as an intracellular protein by western blot analysis (**Figure 2D**). Of note also, the results of western blot analyses (**Figures 2B,D**) demonstrate the absence of appreciable cross-contamination between apoplastic and intracellular fractions. To further confirm this notion, enzymatic activity of glucose 6-phosphate dehydrogenase (G6PDH), an intracellular protein (Debnam and Emes, 1999), was quantified in both fractions. **Figure 2E** shows that the majority of the G6PDH activity was present in the intracellular fraction, with less than 5% of the total activity observed in the apoplastic fraction.

As production of SP-mRFP was found to generate a reliable and soluble apoplastic protein marker, we aimed to use it to assess specificity of phytaspase internalization. Co-production of *Nt*Phyt-EGFP and SP-mRFP in *N. benthamiana* leaves resulted in co-localization of both proteins in the apoplast under non-stressed conditions (**Figures 2F,H,J**). Antimycin A-induced cell death caused the expected shift of the *Nt*Phyt-EGFP fluorescence toward the cell interior (**Figure 2G**). In contrast, mRFP fluorescence retained its extracellular localization (**Figures 2I,K**).

We concluded that clathrin-mediated endocytosis of phytaspase proceeds through a specific recognition step, most likely involving a phytaspase receptor at the plasma membrane of plant cells, rather than by a non-specific fluid-phase uptake.

### Internalized Phytaspase Retains Proteolytic Activity

To determine if phytaspase retains its proteolytic activity upon PCD-induced internalization, we analyzed the behavior of endogenous *N. benthamiana* phytaspase (*Nb*Phyt) first. Apoplastic washes were obtained from *N. benthamiana* leaves containing endogenous phytaspase only (in the absence of *Nt*Phyt-EGFP production) and from leaves producing mRFP-Hub1, either treated with antimycin A or untreated. Phytaspase activity was quantified in these apoplastic washes ("apoplast"), as well as

in residual leaf tissue ("intracellular fraction") and in the total (unwashed) tissues ("total"). The preferred fluorogenic peptide substrate of *Nt*Phyt, Ac-VEID-AFC (Chichkova et al., 2010), was used for proteolytic activity measurements. In the "endogenous *Nb*Phyt only" leaves in the absence of stress, the majority of phytaspase activity was detected in the apoplast. Antimycin A-induced oxidative stress resulted in a dramatic re-localization of the phytaspase proteolytic activity to inside the cell (**Figure 3A**, upper panel). Production of mRFP-Hub1 did not have an appreciable effect on the level and distribution of phytaspase activity in healthy *N. benthamiana* tissues. However, upon the induction of oxidative stress, the inhibition of clathrinmediated endocytosis precluded the accumulation of phytaspase activity inside the cells (**Figure 3A**, lower panel).

Analogous fractionation and proteolytic activity measurements were then performed for *N. benthamiana* leaves transiently producing *Nt*Phyt-EGFP, leaves producing *Nt*Phyt-EGFP together with mRFP-Hub1, either treated with antimycin A or not. The distribution of proteolytic activity of the ectopically produced *Nt*Phyt-EGFP was found to be similar to that of endogenous enzyme in regards to the stress-induced re-localization to inside the cells (**Figure 3B**, upper panel) and the dependence of this retrograde trafficking on clathrin-mediated endocytosis (**Figure 3B**, lower panel). The only variance was the apparently less pronounced difference between the proteolytic activity levels in the apoplast versus the intracellular protein fractions observed upon the *Nt*Phyt-EGFP overproduction.

An increase in the phytaspase-specific proteolytic activity observed upon *Nt*Phyt-EGFP overproduction suggests that this activity belongs to *Nt*Phyt-EGFP. To further verify this assumption, total extracts were prepared from leaves infiltrated with agrobacteria carrying either the wild type *Nt*Phyt-EGFP – encoding plasmid, or the *Nt*Phyt-S537A-EGFP – encoding plasmid driving production of catalytically inactive phytaspase mutant (Chichkova et al., 2010), or the empty vector. Determination of the Ac-VEID-AFC – hydrolyzing activities in total extracts demonstrated a marked increase of the proteolytic activity in the case of wild type *Nt*Phyt-EGFP production, relative to the empty vector control (**Figure 3C**). Production of the inactive *Nt*Phyt-S537A-EGFP protein, on the other hand, failed to increase proteolytic activity over the background level, although both proteins were overproduced to a similar level judging by western blot analysis of the extracts (**Figure 3D**) and fluorescence intensities of infiltrated leaves (**Figures 3E,F**).

Based on the results obtained both with the endogenous and ectopically produced phytaspases, we conclude that induction

(S537A, ~ 120 kDa) proteins. The lower panel, the loading control depicting the Coomassie blue-stained Rubisco band.  Bar, 20 μm.

of cell death does not markedly change the overall level of phytaspase activity in plant tissues. Rather, it causes the re-distribution of proteolytically active phytaspase from the apoplast to inside the cells through the retrograde trafficking step that is critically dependent on clathrin-mediated endocytosis.

### Correlating *Nt*Phyt Activity and Trafficking With Plant Cell Death

Previously, by using transgenic tobacco plants with up- and downregulated phytaspase activity, *Nt*Phyt has been demonstrated to promote plant cell death induced by biotic and abiotic stresses, with concomitant retrograde transport of the enzyme (Chichkova et al., 2010). We, therefore, strived to evaluate whether transient *Nt*Phyt overproduction and/or inhibition of *Nt*Phyt internalization could affect plant cell viability. To address these issues, sevendays-old *N. benthamiana* seedlings were vacuum-infiltrated with agrobacteria carrying either empty vector, or the *Nt*Phyt-EGFP – encoding plasmid, or the mRFP-Hub1 – encoding plasmid, or a combination thereof, or the *Nt*Phyt-S537A-EGFP – encoding plasmid. Cell death in the cotyledons was examined by staining with Evans Blue both with and without treatment of seedlings with antimycin A. As shown in **Figure 4**, antimycin A-induced oxidative stress caused extensive cell death (**A,B**), as expected.

agroinfiltrated to produce mRFP-Hub1 (C,D), *Nt*Phyt-EGFP (E,F), both proteins together (G,H), or neither of them (empty vector, A and B), and *Nt*Phyt-S537A-EGFP (I,J). Three days after agroinfiltration, the seedlings were treated with 10 μM antimycin A for 3 h (the lower row) or mock-treated (buffer only control, the upper row). Subsequent staining with Evans Blue revealed marked cell death in the cotyledons upon antimycin A treatment (B,F,J), which was prevented by the mRFP-Hub1 production (D,H). Overproduction of *Nt*Phyt-EGFP caused moderate damage by itself (E) and again, co-expression of mRFP-Hub1 nullified this effect (G). Overproduction of proteolytically inactive *Nt*Phyt-S537A-EGFP mutant failed to cause cell death in the absence of antimycin A treatment (I). Data were reproducible over three independent experiments.

On the other hand, transient production of *Nt*Phyt-EGFP predisposed cells to death even in the absence of antimycin A treatment (**Figure 4**, compare images **A** and **E**), whereas the proteolytically inactive phytaspase mutant failed to do so (**Figure 4**, compare images **E** and **I**). The pro-death effect of *Nt*Phyt-EGFP production was further enhanced by the subsequently applied oxidative stress (**Figure 4F**). Notably in the presence of mRFP-Hub1, both the antimycin A-induced and *Nt*Phyt-EGFP-promoted cell death was attenuated (**Figures 4D,G,H**). These results are in line with pro-death proteolytic activity of *Nt*Phyt, and furthermore they highlight the importance of clathrin-mediated endocytosis for accomplishment of stress-induced and *Nt*Phyt-promoted death of plant cells.

### Clathrin-Mediated Internalization of *A. thaliana* Phytaspase in a Model System

The *A. thaliana* phytaspase (*At*Phyt) is distinct from the characterized phytaspases in other plant species in a number of ways (Chichkova et al., 2018). In particular, Trusova et al., 2019 recently documented the unexpected mobility of *At*Phyt-EGFP produced in *N. benthamiana* leaves. The recombinant protein expressed in *N. benthamiana* epidermal cells by agroinfiltration initially localizes along cell borders. However after 2 days post infiltration (p.i.), *At*Phyt-EGFP was spontaneously (in the absence of any additional treatment) re-imported into *N. benthamiana* cells (**Figures 5A,B**). This differs significantly from the behavior of the same protein in *A. thaliana* epidermal cells and of *Nt*Phyt-EGFP protein in the non-stressed *N. benthamiana* cells. Notably, EGFP fluorescence in tissues rapidly declined on days 4 and 5 p.i. (**Figures 5C,D**). Spontaneous *At*Phyt-EGFP internalization in this system occurred in the absence of death symptoms in leaf tissue (Trusova et al., 2019) and was critically dependent on clathrin-mediated endocytosis (**Figures 5E–P**). When clathrin-mediated endocytosis was suppressed by co-production of the mRFP-Hub1 inhibitor, no internalization of the enzyme occurred at days 3 and 4 (**Figures 5F,G**), and apoplastically "arrested" *At*Phyt-EGFP persisted outside the cells at day 5 p.i. (**Figures 5E–H**).

Co-production of the SP-mRFP protein together with *At*Phyt-EGFP in *N. benthamiana* leaves revealed that the secreted mRFP retained its apoplastic localization when the internalization of *At*Phyt-EGFP was essentially complete (**Figures 5Q–T**). This indicates that the spontaneous internalization of *At*Phyt-EGFP occurs specifically, similar to the stress-induced internalization of *Nt*Phyt-EGFP described above.

To determine whether the internalized *At*Phyt-EGFP protein retains its proteolytic activity, intracellular proteins were obtained from the *At*Phyt-EGFP-producing leaves on different days following infiltration. *At*Phyt proteolytic activity was quantified using the preferred *At*Phyt fluorogenic peptide substrate Ac-YVAD-AFC (Chichkova et al., 2018), which is also a sub-optimal substrate for the tobacco phytaspase. Internalization of *At*Phyt-EGFP occurring at days 3 and 4 p.i. was followed by an increase of *At*Phyt activity inside the cells (**Figure 6A**). At day 5 p.i. this intracellular proteolytic activity was still clearly detectable, although somewhat decreased.

Western blot analysis with an anti-EGFP antibody of the total protein extracts obtained from the *At*Phyt-EGFP-producing leaves on different days following infiltration revealed the accumulation of the *At*Phyt-EGFP protein at days 2 and 3 p.i., which was followed by a sharp decline of the protein level at days 4 and 5 p.i. (**Figure 6B**). Concomitantly with the *At*Phyt-EGFP disappearance, a protein of approximately 28 kDa was accumulated during days 3–5 p.i. The absence of EGFP fluorescence in leaf tissues at day 5 p.i. (**Figure 5D**) may indicate that this newly formed product corresponds to a non-fluorescent EGFP fragment. Meanwhile, the presence of the YVADase activity in the 5 day p.i. sample suggests that either active *At*Phyt survives after the EGFP detachment, or the observed proteolytic activity does not belong to *At*Phyt.

To clarify this issue, a panel of 10 synthetic fluorogenic peptide substrates used previously to characterize *At*Phyt cleavage specificity (Chichkova et al., 2018) was employed. In full agreement with the established *At*Phyt specificity, protein samples obtained at days 3, 4, and 5 following infiltration hydrolyzed Ac-YVAD-AFC and Ac-IETD-AFC

substrates most efficiently (**Figure 6C**), whereas low level of cleavage or no hydrolysis at all was observed with other fluorogenic peptides.

As an alternative, the YVADase activity in the intracellular protein extracts could belong to vacuolar processing enzyme (VPE), a cysteine-dependent proteinase known to recognize YVAD-based substrates and inhibitors (Hatsugai et al., 2004; Rojo et al., 2004). However, hydrolysis of the Ac-YVAD-AFC substrate by protein extracts was not affected by the presence of 2 mM N-ethylmaleimide, a VPE inhibitor (Hiraiwa et al., 1999) (**Figure 6D**). Finally, when the catalytically inactive *At*PhytS553A-EGFP mutant was transiently produced in *N.*

*benthamiana* leaves (see **Figure 6B** for the relative level of production), no proteolytic activity above the background level was observed with the Ac-YVAD-AFC substrate using total leaf extracts obtained at any time point (**Figure 6A**). Fluorescence microscopy examination of agroinfiltrated tissues confirmed efficient production of both proteins as well (**Figures 6E,F**).

Taken together, these results suggest that the enhanced intracellular YVAD-AFC-hydrolyzing proteolytic activity belongs

Bar, 20 μm.

to the transiently produced *At*Phyt, and that the activity survives within the cell for several days.

To further examine the retrograde vesicular membrane trafficking of phytaspase at the healthy cell background, *At*Phyt-EGFP-producing *N. benthamiana* leaves at 3 days p.i. (when the *At*Phyt-EGFP protein was largely intact) were treated with FM4-64 fluorescent membrane-staining dye. Prolonged incubation with FM4-64 allowed the dye to become endocytosed and stain intracellular membranous compartments (Jelínková et al., 2010), whereas *At*Phyt-EGFP was visualized as intracellular green dots (**Figure 7A**). Measuring green and red fluorescence intensities along the line drawn through phytaspase-positive dots (direction is shown by the arrow in **Figure 7A**) revealed that the internalized *At*Phyt-EGFP (green) is entrapped within the membranous (FM4-64-positive, red) structures (**Figure 7B**). This is consistent with the vesicular membrane mechanism of delivery of active phytaspase into plant cells.

### DISCUSSION

Clathrin-mediated endocytosis was recently shown to drive PCD-induced retrograde transport of phytaspases, plant cell death-related proteases, from the apoplast into the plant cells (Trusova et al., 2019), raising new important questions regarding mechanisms and functional consequences of this unanticipated trafficking pathway. What happens to the proteolytic activity of the enzyme upon internalization? Is the uptake of phytaspases specific, or any soluble apoplastic protein will become internalized upon induction of cell death? Will interference with the phytaspase uptake compromise cell death? Here, we addressed these questions using *N. benthamiana* leaves either containing endogenous phytaspase only, or overproducing *Nt*Phyt tagged with EGFP or mRFP that allowed to follow phytaspase re-distribution with the aid of fluorescence microscopy in parallel with the determination of a peculiar Asp-specific activity of phytaspases. *N. benthamiana* leaves either treated with antimycin A to trigger oxidative stress-induced PCD and phytaspase internalization, or mock-treated were used in these studies.

By separating proteins into apoplastic and intracellular fractions and by quantifying phytaspase proteolytic activity using fluorogenic peptide substrates of phytaspases, we showed that retrograde transport of phytaspases is accompanied by the shift of phytaspase activity toward the cell interior. When internalization of phytaspases was blocked by co-production of Hub, the inhibitor of clathrin-mediated endocytosis, phytaspase proteolytic activity was retained within the apoplast. It is important to note that the overall level of phytaspase activity in leaf tissues was not diminished after phytaspase internalization. Efficient degradation within the vacuole is a frequent outcome for endocytosed proteins (Kleine-Vehn et al., 2008; Korbei and Luschnig, 2013; Claus et al., 2018; Reynolds et al., 2018). However, the internalized phytaspases are evidently able to deviate from the degradation pathway.

Therefore, our results indicate that clathrin-mediated endocytosis accomplishes the delivery of active phytaspases inside the plant cell. The process of phytaspase internalization appears to be specific, as the apoplastic mRFP protein (originating from the signal peptide-mRFP precursor) retained its extracellular localization both before and after the induction of cell death. This suggests that a receptor for phytaspases exists in the plant cell plasma membrane. Indeed, clathrin-mediated endocytosis targets proteins that localize at the plasma membrane and possess cytoplasmic domains, to which clathrin is recruited with the aid of adapter proteins (Traub, 2009; Jackson et al., 2010; Chen et al., 2011; Traub and Bonifacino, 2013; Paez Valencia et al., 2016). Phytaspases, however, are soluble proteins that can be easily obtained in apoplastic wash. Therefore, an interface between phytaspases and clathrin endocytic machinery appears to be necessary. Identification of such a receptor would be an interesting task for the future.

The known involvement of phytaspases in the accomplishment of stress-induced plant cell death (Chichkova et al., 2004, 2010; Reichardt et al., 2018) has pushed us to explore whether prevention of phytaspase internalization would promote cell viability under the unfavorable conditions. We found that interruption of phytaspase uptake through inhibition of clathrinmediated endocytosis correlated with alleviation of cell death induced by oxidative stress (antimycin A treatment) and by *Nt*Phyt overproduction. These findings appear to provide the first indication of the importance of clathrin-mediated endocytosis for stress-induced death of plant cells. Also, inability of the catalytically inactive *Nt*Phyt mutant, in contrast to the wild type enzyme, to cause cell damage further emphasizes the importance of the *Nt*Phyt proteolytic activity for promoting plant cell death. However, although the obtained data are in line with the current model of phytaspase participation in plant PCD (Vartapetian et al., 2011; Chichkova et al., 2012; Trusova et al., 2019), further efforts are obviously required for detailed characterization of the underlying mechanisms.

The second experimental system used in this study, the "*At*Phyt-EGFP in *N. benthamiana*" model, can of course be regarded as artificial and results obtained with this system should be interpreted with some caution. Indeed, we do not yet understand why internalization of the *At*Phyt-EGFP protein begins spontaneously in *N. benthamiana* epidermal cells after a 2-day lag period. A requirement for high level pre-accumulation of the *At*Phyt-EGFP protein within the apoplast to drive internalization could possibly account for the observed phenomenon. Also, what is the difference between *At*Phyt-EGFP and *Nt*Phyt-EGFP that allows the former to enter cells spontaneously, while the latter requires a PCD-inducing stimulus for internalization?

Despite these open questions, the behaviors of *Nt*Phyt and *At*Phyt in *N. benthamiana* leaves share a number of features, as shown by the employment of clathrin-mediated endocytosis for their specific internalization. As spontaneous uptake of *At*Phyt-EGFP does not appear to be associated with *N. benthamiana* cell death (Trusova et al., 2019), the *At*Phyt-EGFP model may offer an opportunity to address details of phytaspase internalization in live plant cells, in the absence of major complicating perturbations occurring in dying cells. With this approach, co-localization of the internalized *At*Phyt-EGFP protein with

### REFERENCES

intracellular membranous vesicles was observed, which is consistent with the endocytic entry pathway for phytaspases.

The described localization dynamics of phytaspases is atypical for plant subtilisin-like proteases, for which secretion into the apoplast was considered to be the end point of their trafficking. In this regard, it would be interesting to learn whether phytaspases represent a rare exception in their retrograde vesicular trafficking, or perhaps other plant subtilases behave in a similar manner under certain conditions.

### DATA AVAILABILITY

The raw data supporting the conclusions of this manuscript will be made available by the authors, without undue reservation, to any qualified researcher.

### AUTHOR CONTRIBUTIONS

AV, NC, and ST designed the study and directed the research. ST, AT, SG, EM, RG, and NC performed experiments. NC, ST, AT, SG, EM, RG, and AV analyzed and interpreted the data. AV wrote the manuscript with contribution from NC. All the authors read and approved the manuscript.

### FUNDING

This work was supported by the Russian Science Foundation (grant nos. 16-14-10043 and 19-14-00010).

### ACKNOWLEDGMENTS

We thank M. Taliansky (The James Hutton Institute, UK) for providing the EGFP-LTI6b construct. We are grateful to the reviewers for their helpful comments and suggestions.

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analysis of the *Arabidopsis thaliana* subtilase gene family. *PLoS Comput. Biol.* 1:e40. doi: 10.1371/journal.pcbi.0010040

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

*Copyright © 2019 Trusova, Teplova, Golyshev, Galiullina, Morozova, Chichkova and Vartapetian. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Phosphorylation of the Chloroplastic Metalloprotease FtsH in *Arabidopsis* Characterized by Phos-Tag SDS-PAGE

*Yusuke Kato and Wataru Sakamoto\**

*Institute of Plant Science and Resources (IPSR), Okayama University, Kurashiki, Japan*

FtsH is an essential ATP-dependent metalloprotease for protein quality control in the thylakoid membrane of *Arabidopsis thaliana* chloroplasts. It is required for chloroplast development during leaf growth, and particularly for the specific degradation of photodamaged D1 protein in the photosystem II (PSII) complex to maintain photosynthesis activity. In the thylakoid membrane, the reversible phosphorylation of proteins is known to control the activity and remodeling of photosynthetic complexes, and previous studies implicate that FtsH is also phosphorylated. We therefore assessed the phosphorylation status of FtsH and its possible role in the regulatory mechanism in this study. The phosphorylation level of FtsHs that compose the FtsH heterohexameric complex was investigated by phosphateaffinity gel electrophoresis using a Phos-Tag molecule. Phos-tag SDS-PAGE of thylakoid proteins and subsequent immunoblot analysis showed that both type A (FtsH1/5) and type B (FtsH2/8) subunits were separable into phosphorylated and non-phosphorylated forms. Neither different light conditions nor the lack of two major thylakoid kinases, STN7 and STN8, resulted in any clear difference in FtsH phosphorylation, suggesting that this process is independent of the light-dependent regulation of photosynthesis-related proteins. Site-directed mutagenesis of putatively phosphorylated Ser or Thr residues into Ala demonstrated that Ser-212 may play a role in FtsH stability in the thylakoid membranes. Different phosphorylation status of FtsH oligomers analyzed by two-dimensional clearnative/Phos-tag SDS-PAGE implied that phosphorylation partially affects FtsH complex formation or its stability.

Keywords: FtsH, thylakoid, chloroplast, protein phosphorylation, protease, photosynthesis

### INTRODUCTION

FtsH is an ATP-dependent zinc metalloprotease with a transmembrane domain. The N-terminal transmembrane domain anchors FtsH to cellular membranes with their ATPase domain facing the membranes. Thus, FtsH protease pulls their substrates out of membranes in an ATP-dependent manner and degrades them to small peptides in the proteolytic chamber (Ito and Akiyama, 2005; Nishimura et al., 2016). FtsH protease is originally identified in a temperature-sensitive phenotype of *Escherichia coli*. FtsH protein homologues have been found in other prokaryotes as well as in organelles of bacterial origin, such as mitochondria and chloroplasts. The fundamental function of FtsH protease is the quality control of membrane proteins, but its contribution to stress responses

### *Edited by:*

*Mercedes Diaz-Mendoza, National Institute of Agricultural and Food Research and Technology, Spain*

### *Reviewed by:*

*Sweden*

*Zach Adam, Hebrew University of Jerusalem, Israel Christiane Funk, Umeå University,*

> *\*Correspondence: Wataru Sakamoto<EMAIL_ADDRESS>
### *Specialty section:*

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

*Received: 22 May 2019 Accepted: 08 August 2019 Published: 10 September 2019*

### *Citation:*

*Kato Y and Sakamoto W (2019) Phosphorylation of the Chloroplastic Metalloprotease FtsH in Arabidopsis Characterized by Phos-Tag SDS-PAGE. Front. Plant Sci. 10:1080. doi: 10.3389/fpls.2019.01080*

is also suggested (reviewed by Janska et al., 2013; van Wijk, 2015; Kato and Sakamoto, 2018).

In the *Arabidopsis thaliana* genome, 12 genes encoding members of the FtsH family have been identified. Nine of these proteins (FtsH1, 2, 5, 6, 7, 8, 9, 11, and 12) are located in the chloroplast (Sakamoto et al., 2003). Additionally, the *Arabidopsis* genome encodes five proteolytically inactive homologues (FtsHi) (Wagner et al., 2012); four of them (FtsHi1, 2, 4, and 5) form a complex with Ycf2 and FtsH12 at the chloroplast inner envelope membrane (Kikuchi et al., 2018; Schreier et al., 2018). This AAA-ATPase complex associates with the chloroplast TIC complex (Nakai, 2018), and participates in the translocation of chloroplast-targeted proteins as the import motor (Kikuchi et al., 2018). Much effort has been devoted to identify and characterize the FtsH protease in the thylakoid membranes. Five FtsH homologues (FtsH1, 2, 5, 6, and 8) function in the thylakoid membrane (Sakamoto et al., 2003; Yu et al., 2004; Yu et al., 2005; Wagner et al., 2011; Zaltsman et al., 2005a). Of them, four FtsH (FtsH1, 2, 5, and 8) form a heterohexameric complex (hereafter simply called FtsH complex) in the thylakoid membrane; these homologues are divided into two types, type A (FtsH1/FtsH5) and type B (FtsH2/FtsH8) (Sakamoto et al., 2003; Yu et al., 2004; Yu et al., 2005; Zaltsman et al., 2005b). Mutants lacking FtsH5 and FtsH2 are known as *yellow variegated1* (*var1*) and *var2*; these mutants show weak and strong leaf-variegated phenotype, respectively, whereas mutants lacking FtsH1 and FtsH8 do not show the leaf-variegated phenotype (Chen et al., 2000; Takechi et al., 2000; Sakamoto et al., 2002). Both types of subunits are essential for the active FtsH complex in the thylakoid membranes and the severity of the mutant phenotype depends on the level of FtsH complex in thylakoid membranes (Chen et al., 2000; Takechi et al., 2000; Yu et al., 2004; Zaltsman et al., 2005b; Miura et al., 2007; Kato et al., 2012a). Mass spectrometry analyses suggested that *Arabidopsis* FtsH complex contains two type-A subunits and four type-B subunits in a hexameric complex (Moldavski et al., 2012), but a study in cyanobacteria (Boehm et al., 2012) and a recent research of our group (Kato et al., 2018) suggested that the ratio between type A and type B subunits in the hexameric complex is 3:3.

Another relevant phenotype which characterizes mutants lacking FtsH is photosensitivity, with higher accumulation of reactive oxygen species (ROS) in the chloroplasts due to impairment of the repairing-capacity of photosystem II (PSII) from lightinduced damage (Lindahl et al., 2000; Bailey et al., 2002; Sakamoto et al., 2002; Sakamoto et al., 2003; Kato et al., 2009). Thylakoid FtsH mediates the degradation of the damaged D1 protein, which is part of the PSII reaction center (Kato et al., 2009; Kato et al., 2012b). The removal of damaged D1 protein is essential for the PSII repair cycle, which is required for the recovery of photosynthetic efficiency reduced by photoinhibition (reviewed by Nixon et al., 2010; Järvi et al., 2015). A possible regulation mechanism of FtsH activity by prohibitin-like proteins, which form a megacomplex with FtsH hexamers, was suggested in *E. coli* (Kihara et al., 1996; Saikawa et al., 2004). The interaction between prohibitin-like proteins and FtsH has been observed in cyanobacteria (Boehm et al., 2012). However, such prohibitin-like proteins have not been reported in chloroplasts, suggesting that thylakoid FtsH may have acquired other regulatory mechanisms along the endosymbiotic process.

Recent studies suggest that an increased turnover rate of FtsH during light irradiation is important for FtsH-mediated protein homeostasis in chloroplasts (Zaltsman et al., 2005a; Li et al., 2017; Wang et al., 2017; Kato et al., 2018). In the chloroplast of seed plants, thylakoid FtsH complex is rather unstable, and FtsH seems to exist in smaller complexes such as dimers (Yoshioka et al., 2010; Kato et al., 2018). The higher turnover rate of FtsH with flexible oligomerization seems to be required for the quality control of itself to access the damaged PSII complex, where ROS might generate at a high rate. On the other hand, disulfide bonds of FtsH controlled by the redox state are involved in the regulation of FtsH oligomerization and its proteolytic activity in *Chlamydomonas reinhardtii* (Wang et al., 2017).

Protein phosphorylation is one of the most important posttranslational modifications (PTMs) in thylakoid membranes (Grieco et al., 2016). For example, phosphorylation of PSII core proteins and light-harvesting antenna proteins (LHCII), which are the best-characterized phosphorylated proteins in the thylakoid membrane, undergoes light-dependent regulation. These proteins are dephosphorylated in dark condition and rapidly phosphorylated under low to moderate light irradiation. Under a high light condition, the phosphorylation of PSII core proteins shows a light-dependent increase, whereas LHCII phosphorylation drastically decreases (Rintamäki et al., 1997; Tikkanen et al., 2006). The phosphorylation of these photosynthetic proteins contributes to the fine-tuning of the photosynthetic apparatus to achieve rapid photosynthetic acclimation. The phosphorylation of LHCII, which needs for state transition, is regulated by the STN7 kinase and PPH1 phosphatase pair (Bellafiore et al., 2005; Bonardi et al., 2005; Pribil et al., 2010; Shapiguzov et al., 2010). On the other hand, the reversible phosphorylation of PSII core proteins is regulated by the STN8 kinase–PBCP phosphatase system, being responsible for both positive and negative modulation of D1 degradation (Bonardi et al., 2005; Tikkanen et al., 2008; Samol et al., 2012; Kato and Sakamoto, 2014; Puthiyaveetil et al., 2014a). Recent studies reported the potential phosphorylation of FtsH proteases in the chloroplasts (Stael et al., 2012 and PhosPhAt 4.0 database). In contrast to extensive studies related to the involvement of PSII core proteins in PSII repair, the role of FtsH phosphorylation in that process, however, has been scarcely addressed and remains unclear.

In this study, we attempted to assess the effect of FtsH phosphorylation on this metalloprotease function. Aiming at that goal, we investigated the potential phosphorylation in FtsH by phosphate-affinity gel electrophoresis using a Phos-Tag molecule (Phos-tag SDS-PAGE) followed by immunoblot analysis. Both types of FtsH subunits were separated into phosphorylated and non-phosphorylated forms. We found that phosphorylation of thylakoid FtsH was neither dependent on light exposure nor the presence of the major thylakoid protein kinases, STN7 and STN8. Instead, we found that the phosphorylation status of FtsH varied with FtsH oligomerization degree. Furthermore, the leaf-variegated phenotype of *var2* was not rescued by expressing a mutated FtsH2 protein, harboring an amino acid substitution in the predicted phosphorylation site. These results imply that FtsH phosphorylation does not regulate FtsH function but FtsH complex formation or its stability.

# MATERIALS AND METHODS

### Plant Materials and Growth Conditions

*Arabidopsis* (*Arabidopsis thaliana*) Columbia ecotype was used as wild type. The mutant lines used in this study, *stn7-1*, *stn8-1*, and *var2-1*, were described previously (Bellafiore et al., 2005; Bonardi et al., 2005; Vainonen et al., 2005; Kato and Sakamoto, 2014). Plants were germinated and grown on 0.7% (w/v) agar plates containing Murashige and Skoog medium supplemented with Gamborg's vitamins (Sigma-Aldrich Corp.) and 1.5% (w/v) sucrose under 12-h light (approximately 100 µmol photons m–2 s–1), at a constant temperature of 22°C. The potential phosphorylation sites in FtsH2 were searched in the *Arabidopsis* Protein Phosphorylation Site Database (PhosPhAt 4.0; http://phosphat.mpimp-golm.mpg.de/).

For creating transgenic plants expressing mutated FtsHs under *var2-1* background, vector constructions were prepared as follows, using the primers shown in **Supplementary Table S1**. Amino acid substitutions in FtsH2 at the putative phosphorylation sites, S292A, T337A, S380A, and S393A, were introduced into the corresponding *VAR2* cDNA by PCRbased site-directed mutagenesis, using the appropriate set of primers (e.g. VAR2 Infusion-Fw/VAR2 S212A-Rv and VAR2 S212A-Fw/VAR2 Infusion-Rv, for S212A). The resulting PCR fragments were cloned into the BamHI-SacI site of a binary vector pBI121 using the In-fusion HD cloning kit (Takara Bio, USA). Final constructs were each subjected to sequencing, to confirm the accurate mutations inserted in the corresponding region. The constructs were transformed into *Agrobacterium tumefaciens* strain LBA4404 cells, and the positive LBA4404 strains were used to transform *var2-1* plants. Transgenic plants were selected by kanamycin resistance, and confirmed by PCR analysis (**Supplementary Figure S5**). At least three independent lines were obtained.

### Sample Preparation and Phos-Tag SDS-PAGE

Isolated thylakoid membranes were used for Phos-tag SDS-PAGE. Harvested 4-week-old seedlings were ground in a blender with ice-cold homogenization buffer (0.35 M sucrose, 50 mM HEPES, pH 7.0, 5 mM MgCl2, 10 mM NaCl, 10 mM NaF) containing EDTA-free complete protease inhibitor (Roche). Homogenates filtered through miracloth (Merck Millipore) were centrifuged at 2,380 × *g* for 10 min. The pellet was resuspended in the same buffer and centrifuged at 300 × *g* for 1 min. The supernatant was again centrifuged at 2,380 × *g* for 10 min. The pellet was resuspended in 1 x SDS-PAGE sample buffer and used for Phos-tag SDS-PAGE analyses. For phosphatase treatment, the thylakoid membranes were resuspended in 1 x Takara buffer for CIAP (supplied with CIAP) with cOmplete™ EDTA-free protease inhibitor (Merck) to a final chlorophyll concentration of 1 µg/ µl. Resuspended thylakoid membranes were incubated with or without CIAP phosphatase for 2 h at 37°C. Phosphorylation of thylakoid membrane proteins was analyzed by Phos-tag SDS-PAGE. Phos-Tag™ was purchased from FUJIFILM Wako Pure Chemical Corporation and used according to the manufacturer's protocol. In brief, gels for Phos-tag SDS-PAGE consisted of a separating gel [10% (w/v) acrylamide, 350 mM Bis-Tris, pH 6.8, 25 μM Phos-tag acrylamide, 100 μM ZnCl2, 0.1% (v/v) N,N,N',N'-tetramethylethylenediamine (TEMED), and 0.05% (w/v) ammonium persulfate (APS)] and a stacking gel [4.5% (w/v) acrylamide, 350 mM Bis–Tris, pH 6.8, 0.1% (v/v) TEMED, and 0.05% (w/v) APS]. Electrophoresis was performed at a constant current of ≤20 mA/gel with the running buffer [100 mM Tris, 100 mM MOPS, and 0.1% (w/v) SDS], to which 5 mM of sodium bisulfite was added. For immunoblot analysis, gels were washed in wash buffer [25 mM Tris, 192 mM glycine, 0.1% (v/v) SDS, 10 mM EDTA] for 10 min three times to remove metal ions, followed by one wash in transfer buffer [25 mM Tris, 192 mM glycine, 0.1% (v/v) SDS, 20% methanol] for 10 min. Then, the proteins were electroblotted to the polyvinylidene difluoride (PVDF) membrane (ATTO Corporation).

# Two Dimensional (2D) Phos-Tag SDS PAGE

Purified thylakoid membranes were used for two-dimensional Phos-tag SDS-PAGE. Purified thylakoid membranes were resuspended in buffer [25 mM Bis–Tris, 20% (w/v) glycerol, pH 7.5] to a concentration of 2 mg/ml chlorophyll. To solubilize thylakoid membranes, an equal volume of n-dodecyl ß-Dmaltoside was added to a final concentration of 1% (w/v). After centrifugation at 14,000 × *g* for 5 min, NativePAGE™ 5% G-250 Sample Additive (Thermo Fisher Scientific Inc.) was added to the supernatant according to the manufacturer's instructions, and samples were loaded onto a NativePAGE™ 4–16% Bis–Tris Protein gel (Thermo Fisher Scientific Inc.) Electrophoresis was performed at 4°C overnight at 50 V. The gel lane was then excised from the gel and incubated in equilibration buffer [50 mM Tris–HCl, 6 M urea, 2% (w/v) SDS, 0.05% (w/v) BPB, 10 mM dithiothreitol] for 30 min at 37°C. Then proteins were separated using Phos-tag SDS-PAGE or conventional SDS-PAGE.

### Immunoblot Analysis

Prior to immunoreaction, transferred membranes were blocked with 1% (w/v) bovine serum albumin (BSA) in 50 mM sodium phosphate buffer, pH 7.5, containing 155 mM NaCl and 0.05% (v/v) Tween 20 (PBST buffer) for 1 h. The membranes were then incubated with anti-D1 (dilution 1:5,000), anti-VAR2 (dilution 1:5,000), anti-VAR1 (dilution 1:5,000), anti-CP43 (Agrisera; dilution, 1:5,000), anti-CP47 (Agrisera; dilution, 1:5,000), and anti-Lhcb1 (Agrisera; dilution, 1:5,000). After two washes with PBST buffer, the membranes were incubated with secondary antibodies Amersham ECL Rabbit IgG, HRP-linked F(ab)2 fragment from donkey (GE Healthcare; dilution, 1:10,000 in PBST). Luminata Forte Western HRP Substrate (EMD Millipore Corp.) was used to develop blots, and chemiluminescence was detected on ChemiDoc XRS+ System (Bio Rad Laboratories, Inc.).

### Fluorescence Measurements

Chlorophyll fluorescence was measured in mature leaves of 6-week-old plants using FluorCam800MF (Photon Systems Instruments). To induce photodamage, leaves were incubated for 4 h under high-light conditions (White LED light; 1,200 µmol photons m–2 s–1). Before measurement, leaves were dark-adapted for 10 min to oxidize plastoquinone. Initial fluorescence yield of PSII (Fo) and maximal fluorescence yield of PSII (FM) were measured. Maximal PSII quantum yield (FV/FM) was calculated as (FM–Fo)/FM.

### RESULTS

### Detection of Phosphorylated FtsH in Thylakoid Membranes

To evaluate potential phosphorylation in FtsH, we employed a phosphate-affinity gel electrophoresis system using a Phos-Tag molecule and subsequent immunoblot analysis. We first tested Phos-tag SDS-PAGE with Mn2+ as a chelating divalent cation using purified thylakoid membranes. The detected signals by immunoblotting using anti-VAR1 and VAR2 specific antibodies, which recognize type A and type B subunits, respectively, showed blurred signals of FtsH proteins, whereas phosphorylated PSII core proteins (D1 and CP43) and a lightharvesting antenna protein (Lhcb1) were properly resolved and separated from their unphosphorylated forms (**Figure 1A**). To improve the mobility shift of FtsH proteins, we next tested the Phos-tag SDS-PAGE procedure using Zn2+ instead of Mn2+, with a neutral pH buffer system (Kinoshita and Kinoshita-Kikuta, 2011). The result shown in **Figure 1A** indicated that the improved Phos-tag method successfully enabled us to detect both the phosphorylated and non-phosphorylated forms of FtsH proteins. A majority of the signals detected by anti-VAR1 antibodies (corresponding to type A FtsH) was observed in the up-shifted band, suggesting that type A subunits are extensively phosphorylated. Meanwhile, the phosphorylated form of type B FtsH detected by anti-VAR2 antibodies was less than half respect to total immunoblot signals (**Figure 1A**). These results are consistent with the different susceptibility of recombinant FtsH5 and FtsH2 found in calcium-dependent phosphorylation assays in a previous report (Stael et al., 2012). In contrast, a non-phosphorylated PSII core protein CP47 did not show any mobility shift in both electrophoresis conditions. The mobility shifts of FtsH proteins were substantially reduced after treatment with calf intestine alkaline phosphatase (CIAP), suggesting that the up-shifted band in the gel indeed resulted from phosphorylation (**Figure 1B**). Additionally, we found unexpectedly that FtsH levels were decreased in phosphatase-treated samples as compared to untreated samples. Since the levels of the non-phosphorylated form of FtsH were not significantly altered after phosphatase treatment, the decrease in FtsH total content seemed to reflect degradation of dephosphorylated-FtsH.

### Phosphorylation of FtsH Under Various Light Conditions and in the Kinase Mutants

Since FtsH is involved in the PSII repair resulting from lightdependent photo-damage (Lindahl et al., 2000; Bailey et al., 2002; Sakamoto et al., 2002; Sakamoto et al., 2003; Kato et al., 2009), we assessed whether the phosphorylated state of FtsH was affected by light conditions. Thylakoid proteins obtained from 4-week-old seedlings preincubated in the dark overnight and exposed to low- (5 µmol photons m–2 s–1), growth- (100 µmol photons m–2 s–1), or high-light (800 µmol photons m–2 s–1) for 2 h were subjected to Phos-tag SDS-PAGE and immunoblotting analysis. The results demonstrated that the phosphorylation level of PSII core protein CP43 increased in a light-dependent manner (**Figure 2**). On the other hand, Lhcb1 had few detectable phosphorylations in the dark incubation; the phosphorylation form increased upon low-light illumination, whereas diminished in the high-light irradiation. These results were consistent with those obtained in the previous studies (Rintamäki et al., 1997; Vener et al., 1998; Tikkanen et al., 2006). However, no obvious change in the phosphorylation state of FtsH proteins could be detected (**Figure 2**), suggesting that phosphorylation of FtsH is independent of the light-dependent regulation of photosynthesis-related proteins.

To clarify the relationship between the phosphorylation state of FtsH and light-dependent kinase activities in thylakoid membranes, we further evaluated the effect of two major thylakoid kinases, STN7 and STN8, on FtsH phosphorylation. STN7 is a thylakoid-associated kinase responsible for LHCII phosphorylation, which is required for state transition (Bellafiore et al., 2005; Bonardi et al., 2005). On the other hand, the phosphorylation of PSII core proteins is mainly regulated by STN8 kinase, but some degree of PSII core proteins phosphorylation is observed in the *stn8* knockout mutant due to an overlap in substrate specificity of STN7 and STN8 (Bonardi et al., 2005; Vainonen et al., 2005). In 4-week-old seedlings exposed to growth light (100 µmol photons m–2 s–1), the major phosphorylation band of Lhcb1 in *stn7-*mutant thylakoids was drastically decreased below the detection level (**Figure 3**), while minor phosphorylation band of Lhcb1 in a lower position was not affected in *stn7* mutant. A previous study reported that there is STN7 independent phosphorylation of a serine residue in Lhcb1; the serine residue corresponds to Ser-48 in Lhcb1.1 protein (Ingelsson and Vener, 2012). It seems that Phos-tag SDS-PAGE allows for detection of serine phosphorylation in Lhcb1. On the other hand, the phosphorylation level of FtsH proteins was comparable with that detected in control samples. Additionally, the band intensity of phosphorylated FtsH proteins obtained from *stn8* mutant seedlings showed no difference compared with that of control seedlings under growth-light condition (100 µmol photons m–2 s–1). Phosphorylation of the PSII core protein D1 was still observed in this experimental condition, suggesting that the aforementioned phenomenon of overlapping activity of these two major thylakoid kinases effectively operated under this light intensity. However, under high-light condition (800 µmol photons m–2 s–1), the phosphorylation level of D1

proteins in the *stn8*-mutant was remarkably reduced (**Figure 3**), indicating that the phosphorylation of photosynthetic proteins was predominantly mediated by STN8 kinase in this irradiance situation. Nevertheless, the phosphorylation state of FtsH in *stn8*-mutant thylakoids showed no difference compared with that in control samples. These results suggested that STN8 is not involved in phosphorylation of FtsH.

### Phosphorylation State and FtsH Oligomerization

Unlike bacterial FtsH megacomplexes, the FtsH hexamer complex in *Arabidopsis* chloroplasts was considered to be unstable, giving rise to smaller oligomers detected in a native PAGE (Li et al., 2017; Kato et al., 2018). To investigate the relation, if any, between phosphorylation and the oligomerization of FtsH, thylakoid membrane preparations solubilized in 1% n-dodecyl-β-D-maltoside were subjected to two-dimensional (2D) gel electrophoresis. Protein complexes were separated by clear-native (CN) PAGE in the first dimension and further separated by Zn2+-Phos-tag SDS-PAGE or conventional SDS-PAGE in the second dimension. The migration of FtsH proteins was resolved by immunoblotting using anti-VAR2 (for Type B) and anti-VAR1 (for Type A) antibodies. Conventional 2D-CN-SDS-PAGE showed that FtsH proteins migrated broadly from the higher molecular weight position, that was around PSII dimer, approximately 650 kDa, to the lower molecular weight size in a range of less than 140 kDa, estimated by the size of LHCII trimer complex (**Figures 4A**, **B**). Since the predicted molecular mass of FtsH monomer is about 70 kDa, the immunoblot signal at the smallest molecular size and that around LHCII trimer seemed to be the monomer and the dimer of FtsH, respectively. On the other hand, the phosphorylation pattern obtained after 2D electrophoresis using Phos-tag showed a noticeable change, particularly in the ratio of non-phosphorylated forms to phosphorylated forms

FIGURE 2 | Phosphorylation of thylakoid proteins under various light conditions. Thylakoid membrane proteins were isolated from dark-adapted seedlings or seedlings exposed for 2 h to low (5 μmol photons m–2 s–1) (LL), growth (100 μmol photons m–2 s–1) (GL), or high (800 μmol photons m–2 s–1) (HL) light. Proteins were separated by Zn2+–Phos-tag SDS-PAGE and immunoblotted with anti-VAR1, anti-VAR2, anti-CP43, and anti-Lhcb1 antibodies. Conventional (Phos-tag-free) SDS–PAGE analysis was also carried out. The phosphorylated form (P) and non-phosphorylated form (N) are indicated. Proteins were loaded equally based on total chlorophyll content. (A Coomassie Brilliant Blue-stained gel of the same samples is shown at the bottom; LHC: light harvest complex). The band density was measured with the ImageJ and showed in Supplemental Figure S2.

in the Type B FtsH oligomers (**Figure 4A**). The immunoblot bands in the 2D CN PAGE showed that the phosphorylated form of FtsH migrated broadly, whereas high accumulation of the non-phosphorylated form of Type B FtsH was observed in the higher molecular weight and monomer positions. In addition, a portion of FtsH in the position corresponding to the dimeric and monomeric forms showed further retarded migration. Distribution of Type A FtsH assessed by anti-VAR1 antibodies showed the similar pattern to that of Type B FtsH, indicating that various oligomeric forms are detected in out 2D gel system (**Figure 4B**). In contrast to Type B, however, the non-phosphorylated forms strongly accumulated in the hexameric and monomeric regions were not noticeably

observed. We reasoned that Type A FtsHs are extensively phosphorylated (**Figures 1** and **3**) and are not detectable in our gel system.

### Amino Acid Substitution of Potential Phosphorylated Residues in FtsH2

In the database of *Arabidopsis* large-scale phosphoproteomic studies (PhosPhAt 4.0) (Heazlewood et al., 2008; Durek et al., 2010), four Ser or Thr residues at position Ser-212, Thr-337, Ser-380, and Ser-393 were reported to be phosphorylated in the FtsH2 mature protein (**Table 1**). Amino acid alignment is shown in **Supplementary Figure S4**. To evaluate whether these potential phosphorylated residues in FtsH2 influence FtsH function, we performed site-directed mutagenesis to create an amino acid substitution. As shown in **Figure 5A**, we made *FtsH2* constructs (driven by CaMV *35S* promoter) in which the sequences corresponding to each Ser or Thr residue were replaced with Ala, and the mutanted FtsHs. The resulting constructs were introduced into a FtsH2-knockout mutant *var2-1*. Transgenic lines were designated as *var2* (S212A), *var2* (T337A), *var2*  (S380A), and *var2* (S393A), respectively. Our previous work indicated that overexpression of FtsH2 restores leaf variegation successfully and accumulates FtsH levels comparable to the wild type (Zhang et al., 2010), suggesting post-translational control of FtsH heterocompex accumulation. As shown in **Figure 6A**, 4-week-old plants indicated that three of these transgenic

lines, *var2* (T337A), *var2* (S380A), and *var2* (S393S), rescued leaf variegated phenotype in *var2-1*, demonstrating that the corresponding amino-acid substitutions are tolerated to restore the defective activity of FtsH. However, we found that leaf variegation persisted in *var2* (S212A), suggesting that Ser-212 is important to maintain its activity (**Figure 6A**).

To examine if mutated FtsH2 proteins accumulated in the transgenic lines, thylakoid membrane proteins from 4-week-old seedlings were subjected to immunoblot analysis (**Figure 6B**). FtsHs accumulated to a similar extent in control, *var2* (T337A), *var2* (S380A), and *var2* (S393A) lines. By contrast, accumulation of FtsH in *var2* (S212A) was reduced considerably, whose levels appeared to be comparable to those in the original *var2* mutant. Since anti-VAR2 antibodies recognize type B FtsH, the remaining bands in *var2* samples were assumed to correspond to another type B protein, FtsH8. Therefore, it is likely that the lack of recovery from variegation in *var2* (S212A) was due to the impaired accumulation of FtsH2 in this line. On the other hand, Phos-tag SDS-PAGE demonstrated phosphorylation of FtsH in the three transgenic lines, *var2* (T337A), *var2* (S380A), and *var2* (S393S); their phosphorylation levels were comparable to that of control. To further evaluate whether these amino-acid substitutions affected FtsH activity in the thylakoid membrane, high-light sensitivity of PSII activity was measured using chlorophyll fluorescence analysis. As expected, an increased PSII photosensitivity was observed in *var2* (S212A), similar to that of *var2* mutant (**Figure 6C**). The FV/FM values in *var2* (T337A), *var2* (S380A), and *var2* (S393S) lines slightly decreased with respect to that of the wild type during highlight irradiation, but these differences were not statistically significant. These results using transgenic plants were thus unable to confirm that phosphorylation of the predicted target residues plays a critical role in FtsH phosphorylation. Nevertheless, our data demonstrated that S212, localized at the stromal edge of transmembrane domain connecting the ATPase domain, is important for FtsH stability.

### DISCUSSION

Protein phosphorylation of FtsH has been observed in large-scale comparative phosphoproteomics studies in *Arabidopsis* (**Table 1**).

In addition, previous studies reported the phosphorylation of FtsH in isolated chloroplasts of pea and *Arabidopsis* (Stael et al., 2012), and also in *Chlamydomonas* (Wang et al., 2014; Szyszka-Mroz et al., 2015). Although several reports suggested the potential phosphorylation of FtsH in thylakoid membranes, its regulatory role remains to be elucidated. Over the past decades, phosphorylation of thylakoid proteins has been characterized as one of the most important regulatory mechanisms of photosynthesis. However, characterization of phosphorylated proteins, focusing on the particular protein target in thylakoid membrane by using immunoblot analysis, appears to have several limitations; for example, conventional immunoblot approaches using anti-phosphothreonine antibodies exclusively detect major phosphorylated proteins such as PSII core proteins and LHCII, which hampers detection of minor phosphorylation. To overcome this drawback, we attempted to evaluate FtsH phosphorylation based on a Phos-tag approach. In PAGE gels containing Phos-tag molecules (Phos-tag SDS-PAGE), Phos-tag interferes with the migration of phosphorylated proteins, thereby exhibiting them as a retarded band, in addition to its nonphosphorylated forms. We found that the use of Zn2+ as divalent cation enabled us to identify the phosphorylated form of FtsH in thylakoid membranes successfully (**Figure 1**).

Since photodamage of D1 in the PSII complex increases with higher light intensity, we may predict the existence of lightinduced regulation that governs the proteolytic machinery for an efficient quality control of PSII activity. One of the lumenal serine-type Deg proteases, Deg1, which facilitates the effective D1 degradation by FtsH in photoinhibitory conditions, has a regulation mechanism dependent on thylakoid lumen acidification (Kley et al., 2011). However, our characterization under various light conditions showed that the phosphorylation state of FtsH in the thylakoid membranes did not undergo light-dependent regulation (**Figure 2**), suggesting that phosphorylation of FtsH is not important to increase proteolysis under high-light intensity. This idea was further supported by the finding that FtsH phosphorylation was not directly regulated by two major thylakoid kinases (**Figure 3**). FtsH phosphorylation is likely mediated by other ill-defined protein kinases localized in chloroplasts. Of note is that in several experiments, bands corresponding to phosphorylated-FtsH proteins from thylakoids of *stn7*-mutant seedlings slightly decreased than those observed in control seedlings; nevertheless, the reproducibility of this result was limited. Given that the loss of STN7 kinase affects both short and long-term photosynthetic acclimation (Pesaresi et al., 2009), we cannot exclude the possibility that indirect effects caused by the loss of STN7 influenced the phosphorylation state of FtsH.

It is unclear as to how FtsH function is regulated by light in the chloroplasts of *Arabidopsis*, apart from the light-induced transcription of FtsH8, which is likely to compensate for the rapid FtsH degradation caused by high-light (Sinvany-Villalobo et al., 2004; Zaltsman et al., 2005a; Wang et al., 2017; Kato et al., 2018). Previous studies demonstrated that FtsH selectively degrades D1 even in dark condition (Lindahl et al., 2000; Krynická et al., 2015), suggesting that light is not always a decisive factor for FtsH protease activity per se; rather, accessibility to their substrates may be crucial for FtsH-mediated protein degradation (Krynická et al., 2015). Together with our results, these observations raise the possibility that the structural change around D1 in the damaged PSII complex is probably more important for the increased proteolysis expected under high-light intensity than light-dependent induction of protease activity in FtsH (Lindahl et al., 2000; Krynická et al., 2015). This possibility well fits with the observation that

light-induced unstacking of the grana expands the grana margin region in the thylakoid membrane; the expansion of grana margin seems to increase the accessibility of photodamaged D1 to FtsH (Puthiyaveetil et al., 2014b; Yoshioka-Nishimura et al., 2014). The alternative possibility may be the redox regulation of FtsH mediated by the formation of Cys disulfide bonding, as suggested in *Chlamydomonas* under high-light irradiation (Wang et al., 2017). Further investigation is necessary to study post-translational regulation of FtsH in *Arabidopsis* chloroplasts.

Amino acid substitutions carried out in FtsH2, based on information provided at the *Arabidopsis* Protein Phosphorylation Site Database (PhosPhAt 4.0), showed that Ser-212 is important for FtsH accumulation, whereas these residues did not critically affect the phosphorylation state of FtsH assessed by Phos-tag analysis (**Figure 6**). The N-terminal Ser-212 residue was found to be conserved between type A and type B subunits among photosynthetic organisms, whereas the Ser-380 in the second region of homology (SRH) motifs of ATPase domain is conserved between type A and type B subunits, but it is not found in FtsH proteins of other photosynthetic organisms (**Supplementary Figure S4**). Besides, Thr-337 and Ser-393 in the ATPase domain are conserved in the type B subunit of FtsH protein between *Arabidopsis* and *Chlamydomonas*, in contrast to type A subunit. The structural model of FtsH hexamer lacks the surrounding transmembrane domain including the position corresponding to Ser-212 (**Figure 5B**). Given the sequence homology among FtsH proteases, Ser-212 would locate between the transmembrane domain and the first alpha-helix in the ATPase domain. This connecting area would be sandwiched between thylakoid membranes and the stromal region of FtsH and seems to be important for substrate translocation and recognition. The phosphorylation of this region was also observed in FtsH5, although it remained unclear which amino acid residue was phosphorylated (**Table 1**). The finding of decreased accumulation of thylakoid FtsH after phosphatase treatment (**Figure 1B**) suggests the possible effect of phosphorylation on FtsH stability in the thylakoid membranes. However, the possibility that in the transgenic line *var2* (S212A), designed for Ser-212 amino acid substitution, this condition itself could have influenced the translation process or the stability of FtsH protein, cannot be ruled out.

In thylakoid membranes, the FtsH functional complex is likely a temporary complex formed when the proteolysis is executed in the grana margin region (Yoshioka et al., 2010; Kato et al., 2018). Thus, FtsH in thylakoid membranes probably does not form a megacomplex with other proteins but is mainly present as smaller complexes. This flexible oligomerization capability of FtsH in the chloroplasts might contribute to the turnover of FtsH itself to maintain its activity under high-light condition (Wang et al., 2017; Kato et al., 2018). Interestingly, our two-dimensional PAGE approach showed different phosphorylation states among FtsH oligomers (**Figure 4**): greater phosphorylation degree of FtsH seems to be found in smaller oligomers, suggesting that FtsH phosphorylation was somehow related to the stability of its monomeric and dimeric forms in the thylakoid membranes.

FIGURE 6 | Complementation analyses of *var2* with mutated FtsH2 plants. (A) Photographs of 4-week-old Col, *var2-1*, *var2* (S212A), *var2* (T337A), *var2* (S380A), and *var2* (S393A) seedlings. Bars = 2 cm. (B) Accumulation and phosphorylation state of FtsH proteins. Thylakoid membrane proteins were isolated from 4-week-old seedlings and subjected to Zn2+–Phos-tag and conventional SDS–PAGE analyses. Immunoblot analyses were performed using anti-VAR1 and anti-VAR2 antibodies. Proteins were loaded equally based on total chlorophyll content. The phosphorylated form (P) and non-phosphorylated form (N) are indicated. A Coomassie Brilliant Blue-stained gel of the samples is shown at the bottom. (C) PSII photosensitivity during high light irradiation. Leaf discs obtained from mature leaves of Col, *var2-1*, *var2* (S212A), *var2* (T337A), *var2* (S380A), and *var2* (S393A) seedlings were exposed to high light (1,200 μmol photons m–2 s–1) and maximal fluorescence yields of PSII (FM) at 0, 60, 120, and 240 min after irradiation were determined. FV/FM values were calculated as indicated in *Materials and Methods* (Data were obtained from six replicates; means ± SD are shown).

However, the possibility that the phosphorylation state of FtsH affects the solubilization process of the FtsH complexes could not be excluded. On the other hand, it is still unclear how calcium-dependent phosphorylation of FtsH, reported in a previous study (Stael et al., 2012), influences FtsH complex formation and function.

### DATA AVAILABILITY

All datasets generated for this study are included in the manuscript/ **Supplementary Files**.

### AUTHOR CONTRIBUTIONS

YK and WS conceived the study and wrote the manuscript. YK performed the experiments. All authors reviewed the results and approved the final version of the manuscript.

### REFERENCES

### FUNDING

This work was supported by KAKENHI grants from Japan Society for the Promotion of Science (16H06554 and 17H03699 to WS; 18K06290 to YK) and from the Oohara Foundation (to WS and YK).

### ACKNOWLEDGMENTS

We would like to thank Rie Hijiya and Kenji Nishimura for technical assistance.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fpls.2019.01080/ full#supplementary-material

cycle of photosystem II in *Arabidopsis* thylakoid membranes. *Plant Cell* 15, 2843–2855. doi: 10.1105/tpc.017319

near photosystem II at grana for the swift repair of damage. *J. Biol. Chem.* 285, 41972–41981. doi: 10.1074/jbc.M110.117432

Zhang, D., Kato, Y., Zhang, L., Fujimoto, M., Tsutsumi, N., Sodmergen, et al. (2010). The FtsH protease heterocomplex in *Arabidopsis*: dispensability of type-B protease activity for proper chloroplast development. *Plant Cell* 22, 3710–3725. doi: 10.1105/tpc.110.079202

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

*Copyright © 2019 Kato and Sakamoto. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Dealing With Stress: A Review of Plant SUMO Proteases

### *Rebecca Morrell and Ari Sadanandom\**

*Department of Biosciences, Durham University, Durham, United Kingdom*

The SUMO system is a rapid dynamic post-translational mechanism employed by eukaryotic cells to respond to stress. Plant cells experience hyperSUMOylation of substrates in response to stresses such as heat, ethanol, and drought. Many SUMOylated proteins are located in the nucleus, SUMOylation altering many nuclear processes. The SUMO proteases play two key functions in the SUMO cycle by generating free SUMO; they have an important role in regulating the SUMO cycle, and by cleaving SUMO off SUMOylated proteins, they provide specificity to which proteins become SUMOylated. This review summarizes the broad literature of plant SUMO proteases describing their catalytic activity, domains and structure, evolution, localization, and response to stress and highlighting potential new areas of research in the future.

### *Edited by:*

*Mercedes Diaz-Mendoza, National Institute of Agricultural and Food Research and Technology, Spain*

### *Reviewed by:*

*Andreas Bachmair, University of Vienna, Austria Anna-Maria Botha-Oberholster, Stellenbosch University, South Africa*

### *\*Correspondence:*

*Ari Sadanandom<EMAIL_ADDRESS>
### *Specialty section:*

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

*Received: 03 June 2019 Accepted: 14 August 2019 Published: 18 September 2019*

### *Citation:*

*Morrell R and Sadanandom A (2019) Dealing With Stress: A Review of Plant SUMO Proteases. Front. Plant Sci. 10:1122. doi: 10.3389/fpls.2019.01122*

Keywords: SUMO protease, SUMO cycle, Ubiquitin-like, Cysteine protease, Post-translational modification, Stress

# INTRODUCTION

SUMO (small ubiquitin-like modifier) proteins are critical for the normal function of eukaryotic cells; the protein is found in all eukaryotes from single-celled yeast *Saccharomyces cerevisiae* to all mammals and plants species. Deleting the only SUMO isoform, SMT3, from a yeast cell, causes a loss of cell viability (Meluh and Koshland, 1995; Giaever et al., 2002) and deletion in *Arabidopsis thaliana* of *sumo1sumo2-1* is embryonic lethal (Saracco et al., 2007; van den Burg et al., 2010), highlighting the critical role of SUMO in cell biology.

SUMO is an 11-kDa protein with one isoform in yeast and eight isoforms currently identified in *A. thaliana* through computational analysis (Novatchkova et al., 2004). In *Arabidopsis*, SUMO1, 2, 3, and 5 are expressed (Benlloch and Lois, 2018), but only SUMO1 and 2 are expressed at high levels (Castaño-Miquel et al., 2011). AtSUMO1 and 2 share 83% amino acid sequence identity and are orthologs of human SUMO2/3, based on sequence similarity. AtSUMO1/2 and human SUMO2/3 also have similar functions and are influenced by stress conditions. Conversely, AtSUMO3/5 are weakly expressed non-conserved isoforms. They are more distantly related to AtSUMO1/2 containing approximately 35% sequence similarity to AtSUMO1/2 (Castaño-Miquel et al., 2011).

SUMOylation is induced by heat, ethanol, drought stress, and oxidative stress; this is conserved in many species from *Arabidopsis, Drosophila,* and *Caenorhabditis elegans* to humans (Saitoh and Hinchey, 2000; Kurepa et al., 2003; Augustine et al., 2016). Indeed, there are many evolutionarily conserved SUMO targets and processes that SUMO is involved in throughout different species. Global proteomic studies in *C. elegans* have identified over 800 SUMO targets; based on these targets, *Arabidopsis* is predicted to have 5660 SUMOylated proteins. The breadth of SUMOylation is on par with other major PTMs (post-translational modifications) like phosphorylation and ubiquitylation (Drabikowski et al., 2018; Millar et al., 2019).

In addition to responding to stress, SUMO is also implicated in many essential cellular processes (Hannoun et al., 2010). Drabikowski et al. (2018) undertook a comprehensive analysis of all the proteins SUMOylated in *C. elegans*. They identified SUMOylated protein targets including proteins involved in genome stability, cell cycle progression, chromatin maintenance and modification, transcription, translation RNA splicing, and ribosome biogenesis. Many identified proteins were non-nuclear localizing in the mitochondria or extracellular matrix. Cytosolic proteins include proteasomal, ribosomal, metabolism, signaling, cell morphology, and motility (cytoskeleton, microtubules, intermediate filaments, and proteins connecting the cytoskeleton to the plasma membrane) proteins. In addition, transport and vesicular transport proteins were identified. The group predicted that at least 15–20% of the eukaryotic proteome can be SUMOylated and suggested that SUMO functions in three main areas: regulation of activity of individual proteins, biogenesis of macromolecular complexes, and SUMO-directed proteasomal degradation (Drabikowski et al., 2018).

A similar study was carried out by Rytz et al. (2018) looking into SUMOylated proteins in *Arabidopsis* found over 1000 SUMOylated targets; many of which were nuclear targeted. The proteins identified included major transcription factors, coactivators/repressors, and chromatin modifiers connected to abiotic and biotic stress defense (Rytz et al., 2018).

The SUMO cycle is likened to ubiquitin due to the similarities in the biochemical steps that catalyze SUMO conjugation and deconjugation of protein substrates (Kerscher et al., 2006). SUMO has a similar 3D structure to ubiquitin, adopting the signature β-grasp fold (characterized by β-sheet with five anti-parallel β-strands and a single helical element between strands β-4 and β-5) but only shares 20% sequence similarity with ubiquitin. SUMO is longer than ubiquitin and has a longer disordered C-terminal tail, which requires processing (Bayer et al., 1998). Like ubiquitin, SUMO can form polySUMO chains of multiple SUMO moieties attached to one lysine or a single-SUMO molecule can be attached to a single lysine residue.

Ubiquitin conjugation to substrate proteins targets them for degradation; conjugation of SUMO can aid the substrate protein for degradation through STUbLs (SUMO-targeted ubiquitin ligases). STUbLs are a novel class of ubiquitin E4 ligases that target proteins with polySUMO chains, ubiquitinating the proteins resulting in degradation. STUbLs have been identified in plants but await biochemical analyses (Elrouby et al., 2013). However, unlike ubiquitin, SUMOylation can have numerous other effects on target proteins, protecting a protein from degradation by protecting lysine residues prone to ubiquitylation, changing its localization, or altering the protein–protein interaction or protein–DNA interactions (Johnson, 2004); the effects of SUMOylation are summarized in **Figure 1**. The protein–protein interaction can occur *via* a non-covalent bond that forms with proteins harboring SIM sites (SUMO-interacting motifs). SIMs are characterized by hydrophobic residues flanked by acidic residues or residues that can be phosphorylated. Alternatively, SUMO can prevent interactions by masking partner-binding sites.

residues in substrates that may be ubiquitinated. D- SUMO binding to a protein can alter its structure activating the protein. E- SUMOylated proteins can signal to STuBL proteins to target for degradation *via* ubiquitination. F- SUMO can block interaction with proteins by blocking binding sites.

The outcome of SUMOylation is largely target dependent and altered by the location and number of SUMO substrates on a target. Indeed, target protein substrates can have single SUMO monomers covalently attached to a lysine, multiple SUMO monomers attached to multiple lysines, or a polySUMO chain from one lysine. This results in a high complexity of different SUMO patterns that can form on one protein altering the molecular consequences of the SUMOylation. For example, the SUMOylation at one site may stabilize the protein by protecting the lysine from ubiquitination, and the SUMOylation at a second site may promote interaction with a protein harboring a SIM site that usually the substrate does not interact with. Despite the complexity of SUMO modification target proteins can experience, it has been observed that, commonly, only a small percentage of target proteins are SUMO modified at a given time entitled the "SUMO enigma" by Hay (2005).

# SUMO CYCLE

SUMOylation is a highly dynamic cyclical process with SUMO changing between conjugated and non-conjugated free SUMO forms. SUMO conjugates are dynamic, changing during the cell cycle and in response to stimuli. The SUMO system accomplishes accurate, rapid, specific responses to stimuli. It has a series of biochemical steps, which are similar to ubiquitination, and involves activation, conjugation, and ligation (**Figure 2**).

Firstly, mature SUMO is generated by a SUMO peptidase cleaving 10 amino acids, exposing a carboxyl-terminal diglycine motif (Johnson, 2004). SUMO is activated by an E1 activation

heterodimer of SAE1/2 (SUMO-activating enzyme). SAE1 has two isoforms SAE1a and SAE1b; either can be used to create the E1 heterodimer with SAE2, using ATP the complex forms SUMO-AMP. The AMP is released from the SUMO-AMP resulting in the formation of a high-energy thioester bond, between the sulfhydryl group of the catalytic cysteine residue in SAE2 and the carboxyl group of glycine in SUMO.

Activated SUMO is transferred from the SAE2 to a cysteine residue in SCE1 (SUMO conjugating enzyme), an E2 conjugation enzyme, in a transesterification reaction to form SUMO-SCE1 thioester complex. This complex catalyzes the reaction of SUMOylation onto a lysine in the target substrate *via* an isopeptide bond between the SUMO carboxyl terminal glycine and the ε-amino group of lysine (K). The lysine, typically, is part of the SUMOylation consensus motif *ψ*KXE/D; *ψ* denotes a large hydrophobic residue, K the acceptor lysine, X any amino acid, and E/D glutamate or aspartate.

SUMO E3 ligases are SCE1-interacting proteins that also help aid the transfer of SUMO from SCE1 to the target substrate. There are two identified E3 ligases in *Arabidopsis* SIZ1 (SAP and MIZ1) and HPY2 (high ploidy2), these complex proteins require a number of domains including nuclear localization and SUMOinteracting domains. However, SUMOylation of a target residue can occur without the presence of E3 ligases; it is as yet unclear how essential E3 ligases are.

An additional catalytic step has been identified involving E4 ligases. These ligases form SUMO chains; two E4 ligases have been identified in *Arabidopsis* PIAL1 and 2 (protein inhibitor of activated STAT-like 1/2).

using ATP and a heterodimer of SAE1 and SAE2. The SUMO is then passed to SCE in a conjugation step, and using a SUMO ligase is ligated onto the substrate. This substrate can then be SUMOylated at more than one SUMO site or form a polySUMO chain. Lastly, in the deSUMOylation step, SUMO is removed from the substrate using a SUMO isopeptidase to generate free SUMO.

Finally, SUMOylation is a cyclical process due to SUMO proteases, which carry out two main functions in the SUMO system. First, they cleave SUMO from target substrates, providing a pool of free SUMO, making SUMO a reversible modification (isopeptidase activity). Secondly, they mature newly synthesized SUMO by cleaving a c-terminal peptide from the immature SUMO (hydrolase/peptidase activity). These sources of SUMO are believed to be critical in the SUMO cycle as the cellular pools of unconjugated SUMO are very low (Johnson, 2004). In yeast, there are two SUMO proteases ULP1 and 2 (ubiquitinlike protease 1/2); the *ulp1-1* mutant is lethal, demonstrating the critical role of SUMO proteases in cell function.

In the ubiquitin system, it is assumed that the ubiquitin E3 ligases provide the specificity in the system due to the large number of E3 ligases. However, in the SUMO system, few SUMO E3 ligases have been identified. Conversely, a relatively larger number of SUMO proteases have been identified, which display specificity to the target proteins suggesting that they may provide specificity in the SUMO system (Chosed et al., 2006; Colby et al., 2006; Yates et al., 2016; Benlloch and Lois, 2018).

### CYSTEINE PROTEASES

There are four major classes of proteases in plants, described in the MEROPS database: cysteine-, serine-, aspartate-, and metallo-proteases, with the protease superfamily comprising 2% of coding genes in plants (Hou et al., 2018). All identified SUMO proteases are cysteine proteases. Cysteine proteases are named after the cysteine residue in their active site which is used as a nucleophile for the formation of an acyl intermediate during proteolytic cleavage (Rawlings et al., 2008). Cysteine proteases are often specialized proteases and are widespread in eukaryotes and in plants, with most species having many types of cysteine proteases. The cysteine proteases are important in plants due to their functions in many processes from seed germination to plant senescence. Environmental cues trigger changes in the proteases enabling the plant to react to stimuli including in response to biotic and abiotic stresses. In plants, cysteine proteases are typically found in lytic vacuoles (Roberts et al., 2012). The superfamily of cysteine proteases is highly conserved and has diversified greatly (Gillies and Hochstrasser, 2012).

### SUMO Proteases Belong to Different Classes of Cysteine Proteases

Previously, the SUMO proteases have belonged to the same protein superfamily, the ULP superfamily, identified due to their similarity to yeast ULP1/2 proteases. Currently, eight ULPs have been predicted in *Arabidopsis*; six have been characterized as SUMO proteases. The active site has a characteristic papain-like fold found in all ubiquitin-specific and UBL (ubiquitin-like) specific cysteine proteases (Mukhopadhyay and Dasso, 2007; Hickey et al., 2012).

However, two new classes of SUMO proteases have since been identified. Shin et al. (2012) identified a novel type of SUMO protease in mouse; the two proteins were named DeSI1 (deSUMOylating isopeptidase1) and DeSI2, which lacked sequence similarity to ULP enzymes. Orosa et al. (2018) identified eight putative DeSI proteases in *Arabidopsis* based on sequence similarity to human DeSI1/2 and functionally characterized one protein, DeSI3a. Another SUMO protease recently identified in humans is called USPL1 (ubiquitin-specific protease-like1) (Schulz et al., 2012); currently, however, no homologues have been identified in *Arabidopsis*.

The three families of SUMO proteases identified are cysteine proteases. The cysteine proteolytic enzymes all contain a cysteine residue that acts as a nucleophile at the heart of the catalytic site triad or dyad capable of breaking the thioester bond between the SUMO and target protein. Also present in the active site in addition to the catalytic cysteine is a histidine that functions as a general base, and additionally, in some cases, there is an extra base that is required for stability. The orientation of the cystine, histidine, and stabilizing amino acid can differ. The SUMO proteases identified in yeast and *Arabidopsis* are listed in **Table 1**.

Surprisingly, the three families of SUMO proteases currently identified (ULP, DeSI, and USLP1) are all members of different clans, which show evolutionary relationships between a broad number of proteases, despite all being SUMO proteases. The different clan classifications are due to the different amino acids and the order of the amino acids required for the active site. Within the clan, the proteases are further characterized into families which shows a statistically significant relationship in amino acid sequence to at least one other family member. The ULP proteases are members of the CE cysteine clan, further characterized into the C48 cysteine protease family. The CE clan is characterized as having catalytic triad with residues in the order histidine, glutamine (or asparagine), and then cysteine. The DeSI proteases are from the CP clan in the C97 cysteine protease family. The CP clan has a catalytic dyad composed of histidine and cysteine: in the DeSI active site, no third residue is required to orientate the histidine ring (Suh et al., 2012).The third type of SUMO protease USPL1 is a clan CA C98 cysteine protease family member (Rawlings et al., 2018). The CA clan is characterized as having the catalytic triad in the opposite orientation to that of CE clan with the residues cysteine, histidine, and asparagine (or Aspartic acid). However, all the proteases contain a papain fold, which characterizes all ubiquitin-specific and UBL-specific cysteine proteases characterized so far (Gillies and Hochstrasser, 2012).

### Proteolytic Mechanism of Cysteine Proteases

Cysteine proteases are characterized by containing a nucleophilic cysteine thiol that is responsible for the peptide bond attack providing the mechanism of proteolytic activity. The adjacent histidine, which acts as a general base, donates a proton to the cysteine residue to enhance the nucleophilicity. The cysteine's anionic sulfur attacks the carbonyl carbon of the substrate, producing the first tetrahedral thioester intermediate in the reaction, releasing an amine or amino terminus fragment from the substrate. Additionally, the histidine residue in the catalytic triad is restored to its deprotonated form, and an intermediate is formed. The thioester bond is hydrolyzed, to produce a carboxylic TABLE 1 | List of currently identified SUMO proteases in yeast and *Arabidopsis*, giving all their known names, TAIR accession number, cysteine protease classification, and tissue expression.

acid moiety from the remaining substrate fragment helping form the oxyanion hole.

The catalytic action of the protease depends on the clan. In the CA clan, including USPL1, catalysis is caused by an acyl-enzyme intermediate, a glutamine amino acid that helps form the oxyanion hole that contains an electrophilic center. The electrophilic center helps stabilize the tetrahedral intermediate, and asparagine orientates the imidazolium ring of the catalytic histidine.

Whereas the ULP proteases are clan CE, the catalysis uses glutamine that, as in clan CA, helps form the oxyanion hole and glutamine that has a similar role to asparagine in CA clan of orientating the imidazolium ring of histidine. Additionally, an asparagine helps stabilize the histidine in the catalytic dyad. Many of the CE proteases show a preference for cleaving diglycine which may be due to a tryptophan following the catalytic histidine (Golubtsov et al., 2006). The tertiary structure for some of the CE proteases has been solved showing the active site is located between two structural subdomains, one being the beta barrel and the second a helical bundle.

In contrast, the DeSI proteases are the only yet identified members of the CP clan, which was identified when the crystal structure was solved (Suh et al., 2012). Unlike clan CA and CE proteases, there is no third active residue to orient the histidine ring; asparagine has been identified as a structurally important amino acid, but it has been shown to not interact with the catalytic histidine. The DeSI1 proteins forms a homodimer; this provides a prominent surface groove between the two monomers, similar to clan CA with the active site being located between a helix and a beta barrel, with the active sites directed toward the groove (Suh et al., 2012).

### IDENTIFICATION OF THE SUMO PROTEASES IN PLANTS

ULP1 (ubiquitin like protease 1) was the first isolated SUMO protease in an activity-based screen of yeast *S. cerevisiae* (Li and Hochstrasser, 1999). ULP1 has an essential role in the G2/M phase of the yeast cell cycle. It was identified by expressing yeast enzymes from a genomic DNA library in *Escherichia coli*. The transformed *E. coli* was also transformed to express a substrate composed of histidine-tagged ubiquitin fused to HA-tagged SMT3. Running the bacterial extracts on SDS-PAGE gels could determine if cleavage after the diglycine motif had occurred between the SMT3 and ubiquitin.

ESD4 (early in short days4) was the first characterized SUMO protease in *Arabidopsis*, firstly identified by Reeves et al. (2002) by conducting a general mutagenesis study. ESD4 was found due to its early flowering phenotype and the reduced mRNA abundance of floral repressor FLC (floral locus C). However, it was not identified as a SUMO protease until Murtas et al. (2003) sequenced ESD4 and searched databases for proteins with similar sequence homology. The search identified similar proteins including human SENP1 (sentrin-specific protease 1), yeast ULP1, and mouse SMT3IP1. The protein similarity was approximately 200 amino acids in the C-terminus forming the active site of a cysteine protease. To confirm ESD4 as a SUMO protease, ESD4 was purified and its peptidase activity monitored *in vitro.* Furthermore, the activity of ESD4 was blocked by thiol reagent (cysteine protease inhibitor) *N*-ethylmaleimide (NEM), which is also inhibited yeast ULP1. However, ESD4 was not inhibited by ubiquitin aldehyde which inhibits deubiquitinating protease. Additionally, a mutant of ESD4, with the catalytic cysteine in the catalytic triad mutated to a serine, was assayed for deSUMOylation activity and was found to be inactive. Finally, the double glycine motif at the C-terminus of the SUMO that linked to FLC was mutated to two alanine residues. No cleavage was detected by ESD4 between the SUMO and FLC. *Arabidopsis esd4-1* mutants also have reduced levels of free SUMO and an increased abundance of SUMO conjugates.

Kurepa et al. (2003) identified *A. thaliana* SUMO proteases by their sequence similarity to yeast ULP1. The study used BLAST to identify *Arabidopsis* proteins with sequence similarity to animal and yeast ULP1 catalytic domains. The catalytic domain is a conserved region of 200 amino acids, which surrounds a triad of histidine, aspartate, and cysteine residues. The search found 12 genes which were further classified into three subfamilies, with two subfamilies more related to yeast ULP1 and the third more similar to yeast ULP2. Of the SUMO proteases identified by Kurepa et al. (2003), they noted on the lack of similarity in the proteins outside of the ULP1 catalytic domain; hypothesizing this may provide substrate specificity.

ELS1 (ESD4-like SUMO protease1) (which was identified by Kurepa et al. (2003) and named ULP1a) was characterized due to its sequence similarity to ESD4. ELS1 contains the same ULP1 catalytic domain as in ESD4 and ULP1. The SUMO activity was also assayed *in vitro* in the same assay as ESD4, using purified ESL1 to cleave SUMO from an extension. As with ESD4, the SUMO protease activity was blocked with NEM and when the catalytic cysteine was mutated to serine. Despite the sequence homology between ESD4 and ELS1, they do not show functional redundancy (Hermkes et al., 2011).

ELS2 was identified in the initial blast search performed by Kurepa et al. (2003) that also identified ELS1, named ULP1b. FUG1 (fourth ULP gene) was identified by Lois (2010) using human SENP1 and was shown to have an expressed sequence tag. It was classified to fourth ULP gene class due to its different phylogeny. Both proteases have been identified *via* bioinformatic techniques but are yet to be functionally addressed.

OTS1 and 2 (overly tolerant to salt 1/2) were initially identified in *Arabidopsis* in the screen by Kurepa et al. (2003) and named ULP1d and ULP1c, respectively. Chosed et al. (2006) and Colby et al. (2006) demonstrated that OTS1 and OTS2 had *in vitro* SUMO protease activity. Chosed et al. (2006) designed an assay with SUMO with an HA (hemagglutinin) tag fused to the diglycine residues and incubated this substrate with purified SUMO proteases. If the SUMO-HA is cleaved, the product runs faster on an SDS/PAGE gel and can be identified. The assay demonstrated that OTS1 and OTS2 can cleave SUMO1 and SUMO2 to produce mature SUMO. They then tested for isopeptidase activity by purifying RanGAP (RanGAP [GTPase-activating protein]) substrate modified by various recombinant GST-SUMO proteins. Both OTS1 and OTS2 were capable of cleaving various SUMO variants from RanGAP. In a similar approach, Colby et al. (2006) also expressed purified OTS1 and OTS2 and assayed for isopeptidase activity using *in vitro* SUMOylation. Initially, they tested AtSUMO1, 2, and 3 conjugated to ScPCNA (proliferating cell nuclear antigen). OTS1 and OTS2 cleaved SUMO1 and SUMO2 from ScPCNA. Colby et al. (2006) also used a similar assay to determine that OTS1 and OTS2 in addition to possessing isopeptidase activity also possess peptidase activity capable of maturing SUMO. Conti et al. (2008) demonstrated that *ots1ots2-1* mutants had increased levels of SUMO conjugates.

SPF1 and SPF2 (SUMO protease related to fertility 1/2) were initially identified by Novatchkova et al. (2004) and called ULP2 like-2 and ULP2-like-1, respectively. They were identified by searching the *Arabidopsis* genome for a protein that was most similar to the yeast query protein, ULP2. The proteins identified however were not characterized as a SUMO proteases until 2017. Two groups characterized the proteins: Liu et al. (2017a) renamed the proteins SPF1 and SPF2, Kong et al. (2017) renamed the ULP2-like-2 ASP1 (*Arabidopsis* SUMO protease1). Kong et al. (2017) confirmed *in vitro* that SPF1 has endopeptidase activity by incubating purified SPF1 with SUMOylated FLC and demonstrating that SUMO was cleaved by WT (wild-type) SPF1 but not by a mutation to the cysteine catalytic site or when NEM was added to the incubation. Additionally, they showed that both SPF1 and SPF2 can process immature SUMO to the mature form, and both single and double mutant knockouts have higher levels of SUMO conjugates. Kong et al. (2017) showed that the number of SUMO conjugates remain higher after heat shock in the *spf1-1* mutant.

Unlike the ULP proteins, DeSI1 was not identified from yeast as it does not contain a homologue. Originally, the DeSI proteins were identified as PPPDE (peptidase-permuted papain fold peptidases) of dsRNA (double-stranded RNA) viruses and eukaryotes. The DeSI proteins were identified in mice by Shin et al. (2012) using BZEL (BTB-ZF protein expressed in effector lymphocytes) in a yeast two-hybrid screen as bait. It was predicted to function as a deubiquitinating peptidase, but no activity had been reported. DeSI1 was unable to deubiquitinate ubiquitinated BZEL but was capable of deSUMOylating SUMOylated BZEL; additionally, DeSI1 was capable of cleaving polymeric SUMO2/3 from targets in mouse. Mutating the catalytic cysteine in DeSI1 abolished the deSUMOylation capabilities of DeSI-1. However, the DeSIs do not have SUMO-processing peptidase activity (Shin et al., 2012, Suh et al., 2012).

Orosa et al. (2018) searched for *Arabidopsis* homologues using the mammalian DeSI1 active site as a search criteria identifying eight putative *Arabidopsis* DeSI proteins. One protein, named DeSI3a, was purified and assayed *in vitro* for deSUMOylation activity, compared to the same protein with the catalytic cysteine mutation to serine, which was incapable of deSUMOylation. DeSI3a WT (wild type) showed cleavage of the isopeptide-linked SUMO; the mutated DeSI3a did not show activity. Additionally, DeSI3a was shown to specifically reduce higher molecular weight SUMO-conjugated isoforms of the kinase domain of FLS2 (flagellin-sensitive2), which the mutated form of DeSI3a was unable to (Orosa et al., 2018).

Schulz et al. (2012) were encouraged to search for novel SUMO proteases due to the low number of identified SUMO proteases and the small number of different families of SUMO proteases compared to ubiquitin proteases. They used an activity-based search with suicide substrates that irreversibly cross-link with SUMO proteases. This technique has been used for the identification of ubiquitin proteases; they purified HA-tagged SUMO ligated to vinylmethylester and incubated the SUMO with human cell lysates. The HA-tagged proteins were immunopurified from the lysate and analyzed with mass spectrometry. This led to the identification of USPL1, which, when the catalytic cysteine was mutated to the serine, USPL1 was no longer capable of binding to the SUMO. USPL1 was shown to interact with SUMO2, but not ubiquitin, to have some peptidase activity, and it shows some chain editing activity. Fluorescencetagged USPL1 was found exclusively in Cajal bodies which are in the nucleus, associated with mRNA processing, and are highly dynamic, changing in number, size, and composition during cell cycle, development, and stress (Cioce and Lamond, 2005; Nizami et al., 2010). Interestingly, RNAi-mediated knockdown of USPL1 affected proliferation and COILIN localization; however, the phenotype was rescued by both USPL1 and the mutated form of USPL1, suggesting it has other functions (Schulz et al., 2012; Hutten et al., 2014). USPL1 highlights the importance of characterization of proteases, as USPL1 was originally misannotated as an ubiquitin protease due to sequence similarity.

Following previously outlined techniques to identify SUMO proteases in *Arabidopsis,* the catalytic site of USPL1 was blasted into the *Arabidopsis* genome. Two proteases were identified as potential matches: UBP6 and UBP7 (ubiquitinspecific protease6/7) shown in **Figure 3**. Both these proteases have already been identified as ubiquitin proteases through bioinformatic techniques. Moon et al. (2005) identified the protease as an interacting partner with CAM2 (Calmodulin2). However, they were unable to demonstrate in *E. coli* that UBP6 was capable of cleaving ubiquitin from substrates. It can only be speculated that UBP6 and its close homologue UBP7 are SUMO proteases currently. It will require functional characterization to prove their SUMO proteolytic activity. If, however, it is shown that UBP6/7 are SUMO proteases, it will be the third identified cysteine protease family present in *Arabidopsis*; there may be more yet.

### DOMAINS AND STRUCTURES OF SUMO PROTEASES

The domains and structures of plant SUMO proteases have not been well studied; they are largely based on similarity to yeast SUMO proteases. The ULP proteases typically have a 200–amino acid catalytic domain at the C-terminus of the protein (see **Figure 4**). The N-terminus of the protein is typically highly variable and is presumed to be responsible for specificity of SUMO protein– conjugate recognition and modulation of enzymatic activity and directing subcellular localization (Li and Hochstrasser, 2003; Gong and Yeh, 2006; Mukhopadhyay et al., 2006; Kroetz et al., 2009). The N terminal domain is also thought to contain SIM (SUMOinteracting motif) sites which may increase enzyme affinity for SUMOylated substrates; alternatively, they may help aid orientation of the SUMOylated proteins in the catalytic site (Hickey et al., 2012).

However, as can be seen in **Figure 4**, the structure of the ULP proteases varies between the proteases, helping to provide their

UBP7. The red boxes highlighting the conserved residues indicate the catalytic triad.

FIGURE 4 | The location of the active site and length of several different SUMO proteases. The orange box denotes the SIM site in ULP1. The blue oval depicts the location of the C48 active site, the green oval the C97 active site, and the purple oval the C98 active site.

individual specificity. The catalytic domain in ELS2 is closer to the N-terminus, and the catalytic domains in SPF1 and SPF2 are located in the center of the protein and form much larger proteins than the other ULP proteases. Chosed et al. (2006) examined the effect of truncating the OTS1, OTS2, ELS1, and ESD4, expressing just the catalytic domain *in vitro*. Truncated ESL1 and ESD4 were unable to function as SUMO proteases requiring the full length of the gene. Conversely, the truncated form of OTS2 was able to cleave more forms of SUMO than when the N-terminus of the protein was intact; the truncated form of OTS2 was able to cleave yeast and mammalian and tomato SUMO; this activity was not present in the full length of OTS2 (Chosed et al., 2006).

Additionally, the *Arabidopsis* ULP proteases have not had the crystal structure solved. It is assumed, however, that the structure will be similar to yeast ULP1. The crystal structure of yeast ULP1 catalytic domain interacting with SUMO has revealed a tight but shallow VDW tunnel that recognizes the Gly–Gly motif, stably orienting substrates to come into close interaction with the active site. The structure showed that side chains of residues other than glycine would sterically clash with the narrow tunnel, providing specificity (Mossessova and Lima, 2000). When the substrate is positioned in the VDW tunnel, the scissile bond after the double glycine is converted from a trans to a cis bond causing a kink in the SUMO C-terminal tail (Shen et al., 2006). This configuration is thought to be rare in proteins and is induced to destabilize the bond and promote cleavage (Hickey et al., 2012). The active site in ULP1 is in a narrow cleft structured to enable both large SUMO conjugates and single-SUMO molecules to access it (Mossessova and Lima, 2000). The active site is between two structural subdomains, one subdomain is a beta-barrel carrying the active site histidine and glutamine (or aspartic acid), and the second subdomain consists of a helical bundle, with one helix carrying the catalytic cysteine.

Compared to the ULPs, the DeSI proteins are smaller proteins with a larger component of their size comprising the active site (**Figure 4**), which is around 140 amino acids; the active site is also closer to the N terminus. As with the ULP proteases, the DeSI proteases have putative SIM sites (Hickey et al., 2012). The crystal structure for the DeSI proteases in *Arabidopsis* has not been solved but can be assumed to be similar to the solved structure of human DeSI1. It revealed the protein forms as a homodimer forming a papain groove between the two subunits forming the active site with the catalytic dyad. In the DeSI protein, the proteolytic groove forms between the cysteine at the N-terminal end of a helix, and the histidine on a strand that is part of a betabarrel. Surprisingly, the C-terminal tails of DeSI1 were shown to fold into the groove, seemingly blocking access to the active site. However, activity assays with a truncated C-terminal tail show no effect compared to WT, suggesting the structure may be an effect of crystallization (Suh et al., 2012).

The domains and structure of USPL1 have not currently been characterized but can be hypothesized to have similar functions to that of the ULP proteases containing SIM site signals for cellular localization and providing substrate specificity.

### EVOLUTION AND DIVERSIFICATION OF SUMO PROTEASES IN PLANTS

The ULP SUMO proteases are evolutionarily distinct from the DeSI SUMO proteases (**Figure 5**). The ULP SUMO proteases have a phylogenetic origin that can be traced to green algae and other eukaryotes including yeast ULP1 and ULP2 (Castro et al., 2018a). The evolutionary classification of the proteases has proved difficult due to the high amino acid sequence divergence. Initially, OTS1 and OTS2 were classified in a group that was more closely related to ESD4/ELS1/ELS2 believed to be related to *Sc*ULP1 due

to the active site being located at the C-terminus of the protein like *Sc*ULP1 (Novatchkova et al., 2004; Lois, 2010). However, based on amino acid conservation, they are more similar to *Sc*ULP2 (Castro et al., 2016). The current classification in use was carried out by Novatchkova et al. (2012), who conducted an in-depth phylogenetic analysis, including *Arabidopsis*, tomato, grapevine, and poplar genomes. They generated a novel grouping of the ULP proteases into four groups in *Arabidopsis*, namely, A, B1, B2, and C (Novatchkova et al., 2012). Benlloch and Lois (2018) suggested using the same classification system classing the ULP proteases on sequence similarity and organization of the ULP domain; however, they suggested classing the proteases independently of their similarity to yeast ULPs. Conversely, Castro et al. (2018a) used the classification system to divide the ULP proteases into evolution from ScULP1 and ScULP2; however, the naming system was changed and is being used in this review. Class I ELS type of homologues is believed to have evolved from ScULP1 including ESD4, ELS1, and ELS2. Three different subdivisions have evolved from ScULP2 including class II OTS-type homologues including OTS1 and OTS2 class III SPF-type homologues, including SPF1 and SPF2 and class IV FUG type that is yet to be characterized as a SUMO protease. However, the relationship of FUG1 to the other SUMO protease groups strongly suggests that it has the same activity (Novatchkova et al., 2012). FUG1 may be a relatively newer protease as it appears to be absent from early plant taxa, being present in flowering plants (Castro et al., 2018a).

The DeSI proteins are not as well studied evolutionarily due to their very recent identification in *Arabidopsis*; however, due to the similarities in some of their sequences, it can be hypothesized that they may share functional redundancy (Orosa et al., 2018), but this is yet to be determined.

If USPL1 homologues are found in *Arabidopsis*, it is likely that they will share more evolutionary similarity with the ULP proteases as clan CE proteases are hypothesized to share sequence similarity with clan CA proteases.

# LOCALIZATION OF SUMO PROTEASES IN THE CELL AND PLANT TISSUE

The subcellular localization of the SUMO proteases is thought to provide specificity to the SUMOylation machinery (Chosed et al., 2006). Li and Hochstrasser (2003) expressed in yeast cells lacking *Sc*ULP2 a truncated form of *Sc*ULP1 with just the catalytic domain expressed; this mutant was capable of suppressing defects of cells lacking ULP2, whereas full length *Sc*ULP1 was unable to. This suggests that the N-terminal region of ULP1 restricted activity of the protease; this may have been its cellular location to enable proteolytic activity. The known subcellular localization of the *Arabidopsis* SUMO proteases are summarized in **Figure 6**. The cellular localization is largely based on the N-terminal sequence; deletion of the localization domain alters the targets that are deSUMOylated (Li and Hochstrasser, 2003; Gong and Yeh, 2006; Mukhopadhyay et al., 2006; Kroetz et al., 2009). Yeast ULPs all localize to the nucleus, *Sc*ULP1 localizes to the nuclear pore complexes in the nuclear envelope (Panse et al., 2003), and *Sc*ULP2 localizes to the nucleoplasm (Li and Hochstrasser, 2000).

The ULP proteases largely all localize to the nucleus; ESD4 predominantly localizes to the periphery of the nucleus at the nuclear periphery and envelope (Murtas et al., 2003; Xu et al., 2007). Surprisingly, unlike the other ULP proteases, the closest homologue of ESD4, ELS1, is present in the cytosol (Hermkes et al., 2011). OTS1 and OTS2 localize to the nucleus, and OST2 is also found in nuclear foci (Conti et al., 2008). Additionally, the recent characterization of SPF1 and SPF2 showed that they are nuclear proteases (Kong et al., 2017; Liu et al., 2017a).

The DeSI1 proteases identified in human cells have been found to be located at both inside and outside the nucleus, localizing predominately in the cytoplasm, whereas DeSI2 localizes exclusively in the cytoplasm (Shin et al., 2012, Suh et al., 2012). The only DeSI currently characterized in *Arabidopsis*; DeSI3a was found to localize in the cell membrane by expressing mCherry-DeSI3a in *Nicotiana benthamiana*. Ultracentrifugation which separated the cytoplasmic nuclear and membrane fractions identified DeSI3a in the membrane fraction (Orosa et al., 2018).

The SUMO proteases are localized in all tissues in the plant; generally, their localization correlates with their phenotypes when knocked out. The highest concentration of ESD4 mRNA was detected in the inflorescence and flowers; however, it was also detected at lower concentrations in the seedlings, leaves, shoots, and roots of wild-type plants and was found to be constantly present throughout the 24-hour cycle (Murtas et al., 2003). An ELS1 promoter–GUS fusion showed that ELS1 is expressed ubiquitously in the plant with a higher accumulation in the vasculature and roots. RT-PCR demonstrated that there is also high expression in flowers and low levels of expression in the siliques and leaves (Hermkes et al., 2011).

Both OTS1 and OTS2 have a similar expression pattern as observed through GUS staining, both present from the early developmental stages, with high expression levels in vascular tissue in the root and shoot of seedlings and in the petioles. In mature plants, expression in the leaves was reduced, compared to the seedlings. The proteases were also identified in the flowers and siliques with OTS2 expression stronger than OTS1; for most other tissues, OTS1 expression was stronger (Castro et al., 2016).

The GUS reporter system for SPF1 expression found ubiquitous expression in 2-day-old seedlings; 4-day-old SPF1 was detected in the hypocotyl, cotyledons, and shoot and root apices of the seedlings. In older seedlings, it is present in newly developing leaves, shoot apex and root tips (Kong et al., 2017; Liu et al., 2017a). SPF1 and SPF2 strongest gene activity was detected in the reproductive organs, specifically localizing to embryo sacs, inflorescences, anthers, and developing seeds (Kong et al., 2017; Liu et al., 2017a). Despite SPF1 and SPF2 being expressed in the same tissues, both have different expression patterns in the respective tissues. Tissuespecific PCR revealed that SPF1 transcription is highest in inflorescences and cauline leaves, with intermediate expression levels in stems and rosette leaves. SPF2 transcription was seen to be at its highest in stems, cauline leaves, rosette leaves, and middle-length siliques, and interestingly, no expression of SPF2 was detected in root tissue (Liu et al., 2017a).

### SUMO PROTEASE SPECIFICITY