All experiments were carried out with Lotus japonicus ecotype B-129 F9 GIFU. Sterilized seeds were sown on H₂O agar plates and left over night at 4°C cap side down. After 24 h in the dark in the growth chamber, Petri dishes were exposed to light and kept in a vertical position. Care was taken to maintain the young emerging roots in contact with the filter paper. In vitro cultures were maintained in a growth chamber with a light intensity of 200 μmol m⁻² s⁻¹ at 23°C with a 16 h/8 h day/night cycle. Solid growth media had the same composition as that of B5 medium (Gamborg, 1970), except that, when needed, (NH₄)₂SO₄ and KNO₃ were omitted and replaced by ammonium nitrate (NH₄NO₃). KCl was added to the medium to replace the potassium source. The media contained vitamins (Duchefa catalog G0415) were buffered with 2.5 mM MES and pH was adjusted to 5.7 with KOH.

For RWC analyses, plants from in vitro cultures were transferred to the growth chamber with a 16 h/8 h day/night cycle in pots (2 plant per pot) containing 70 g of a sand/vermiculite mixture (3:1). The plants were watered every 2 days with 30 ml of half strength BandD (Broughton and Dilworth, 1971) solution containing 5 mM NH₄NO₃ for three additional weeks until the starting of drought treatment by stopping irrigation.

Plants were grown under greenhouse conditions for Scanalyzer 3D platform in pots of 2 L (16 cm diameter, 19 cm height; one plant per pot), containing 1.5 kg of soil consisting of a 50:50 mixture of peat moss and river sand. Growth conditions were 23/17°C day/night temperature, 75%, relative humidity, and 16 h per day photoperiod. Plants were irrigated with 100 ml of water every 3 days during the analyses.

Phenotyping through image analysis was performed with a LemnaTec Scananalyzer 3D System. Images of Lotus plant chlorophyll fluorescence were taken under illumination of light having wavelengths greater than 500 nm and with a filter in front of the camera to remove wavelengths shorter than 500 nm. The images of fluorescent light from the Lotus plants were then converted from the red/green/blue (RGB) color space to the hue/saturation/value color space. The color pixels of the plants were divided into a scale of 13 classes. When a plant is placed under greater levels stress more fluorescent light of higher energy (or shorter wavelength) is released and this change might be measured through pixel distribution. To quantify this shift a photosynthetic health index is calculated with the formula (F_x − F_y)/(F_x + F_y), where F is the number of pixels in a specific color class while x and y are the color classes. The color classes chosen were determined experimentally for each experiment by examining the hue histogram.

Bio-volume parameters were measured as described by Eberius and Lima-Guerra (2009).

We followed the procedures previously described (Lombari et al., 2003; Barbulova et al., 2005).

Oligonucleotides and strategies for prLjGLB1-gusA, LjGLB1-GFP fusions and PII over-expressing T-DNA constructs were described in D'Apuzzo et al. (2015).

Confocal microscope analyses were performed using a Nikon PCM2000 (Bio-Rad, Germany) laser scanning confocal imaging system. For GFP and RFP detection, excitation was at 488 nm and detection between 515 and 530 nm. For the chlorophyll detection, excitation was at 488 nm and detection over 570 nm.

Total RNA was prepared from Lotus leaves using the procedure of Kistner and Matamoros (2005). The samples were treated with DNAse I (Qiagen) to remove contaminating DNA the absence of which was subsequently confirmed by PCR. One microgram of total RNA was annealed to random decamers and reverse-transcribed with reverse transcriptase (Qiagen) to obtain cDNA. Real time PCR was performed as described in Omrane et al. (2009) with a DNA Engine Opticon 2 System, MJ Research (MA, USA) using SYBR to monitor dsDNA synthesis. The ubiquitin (UBI) gene (AW719589) was used as an internal standard. The concentration of primers was optimized for each PCR reaction and each amplification was carried out in triplicate. The PCR program used was as follows: 95°C for 3 min and 39 cycles of 94°C for 15 s, 60°C for 15 s, and 72°C for 15 s. Data were analyzed using Opticon Monitor Analysis Software Version 2.01 (MJ Research). The qRT-PCR data were analyzed using comparative Ct method. The relative level of expression was calculated with the following formula: relative expression ratio of the gene of interest is 2^−ΔCT with ΔCT = Ct_AMT1 minus CT_UBI. The efficiency of the different AMT1 primers was assumed to be 2.

Analysis of the melting curve of PCR product at the end of the PCR run revealed a single narrow peak for each amplification product, and fragments amplified from total cDNA were gel-purified and sequenced to assure accuracy and specificity. The oligonucleotides used for the qRT-PCR are the following: LjNR-forw 5′-TGCAGAACTTGCCAATGAAG-3′; LjNR-rev 5′-GTTTCTCCACCGTCCAGTGT-3′.

Histochemical staining of whole plant material was performed as described by Rogato et al. (2010) and Ferraioli et al. (2004).

To follow ABA induced stomatal closure, epidermal strips were prepared from well-watered 3- to 4-week-old plants at the 6th hour of the daily light period. Strips are incubated in 30 mM KCl, 10 mM MES-KOH, pH 6.5, at 22°C, and exposed to light (300 μmol m⁻² s⁻¹) for 3 h in the presence or in the absence of 10 μM ABA. Stomatal apertures were measured using a Nikon Optiphot-2 microscope (http://www.nikon.com/) fitted with a digital camera and a TG 1017 digitizing table (Houston Instruments; http://www.tms-plotters.com) linked to a personal computer. To study ABA induced inhibition of stomatal opening, leaves were harvested in darkness at the end of the night period, epidermal strips prepared and stomatal measurements performed as above with/out 10 μM ABA. To measure constitutive stomata aperture, homogeneous stem cuttings were obtained from well-watered 3- to 4-weeks-old plants at the 6th hour of the daily light period. The stem bases were immediately sealed with instant adhesive glue on a microscope cover glass and leaves were detached at T0, 1, and 2 h after sealing for epidermal strips preparation. For each treatment, at least 60 stomatal apertures were measured and data presented are the mean of at least 3 independent experiments (2 leaves per experiment).

Homogenous stem cuttings from 3 to 4 weeks old wild type and transgenic plants, carrying 4–5 trifoliates from the top second or top third internodes were detached. The stem bases were immediately weighted and sealed with instant adhesive glue on a microscope cover glass. Dehydration and weight measurements at the designated time intervals were performed in a dark room. Water loss is described as the percentage of initial fresh weight. Five replicates from different individual plants were analyzed.

To measure RWC, 24 leaves per replicate were used. RWC was calculated using the formula: RWC (%) = ((FW − DW)/(TW − DW)) × 100 (Hewlett and Kramer, 1962). Leaves of corresponding trifoils were detached from wild type and transgenic plants and immediately weighed to determine their FW (fresh weight). The same leaves were weighted after 24 h floating in deionized water and blotted dry to determine their TW (turgid weight). DW (dry weight) measurements were taken after the leaves were oven-dried (65°C) in paper bags for 24 h. Each biological replicate (four) consisted of a pool of leaves from two plants that were grown in the same pot.

The cell-permeable diacetate derivative diamino-fluorescein-FM (DAF-FM DA) (Sigma-Aldrich Cod. D2321) was used as a specific fluorescent probe for the detection of intracellular NO (Gould et al., 2003). Epidermal strips obtained from leaves of adult Lotus plants grown in pots were prepared accordingly to Behera and Kudla (2013) incubated for 10 min in the loading buffer (LB) (30 mM KCl, 10 mM MES-KOH). The buffer was then replaced with fresh LB supplemented with 5 μM DAF-FM-DA (0.1% v/v final DMSO) and kept for 15 min in the dark. The epidermal strips were then washed twice (2 × 5 min) with fresh LB before being visualized by microscopy.

For NO detection analysis, Lotus guard cells loaded with DAF-FM-DA were imaged by an inverted fluorescence microscope Nikon Ti-E (Nikon, JP, http://www.nikon.com/) with a 20 × A.N.0, 75 Plan Apo VC dry objective. Excitation light was produced by a fluorescent lamp Prior Lumen 200 PRO (Prior Scientific, UK, http://www.prior.com). The specimens were excited at 470/40 nm and the DAF-FM-DA fluorescence was collected with a bandpass filter of 505–530 nm. The fluorescence emitted was collected with an Hamamatsu Dual CCD Camera ORCA-D2 (Hamamatsu, Photonics, JP, http://www.hamamatsu.com/). Exposure time was 200 ms with a 1 × 1 CCD binning. The NIS-Element (Nikon, JP, http://www.nis-elements.com/) was used as platform to control microscope, illuminator, camera and post-acquisition analyses. Pixel intensity analysis of guard cells loaded with DAF-FM-DA was determined with FIJI (https://fiji.sc/). The DAF-FM-DA fluorescent signal was background subtracted.

Leaf extracts of wild type and PII over-expressing plants were prepared in 100 mM Tris-HCl pH 8.0, 10 μM FAD, 1 mM EDTA, 1 mM dithiothreitol (DTT), 1 mM phenylmethylsulphonylfluoride (PMSF) and 0,1% Triton X-100. The extracts were assayed for Nitrate Reductase activity as described in Pajuelo et al. (2002).

Statistical analyses were performed using the VassarStats analysis of variance program.

We have recently described the GUS activity driven by the prLjGLB1-gusA promoter-fusion, localized in vascular bundle of Lotus japonicus transgenic hairy roots and nodules with a slight activity observed in root tips (D'Apuzzo et al., 2015). In order to gain further inside the profile of LjGLB1 expression in different organs, we have used the same T-DNA construct to obtain L. japonicus transgenic plants via A. tumefaciens-mediated transformation. Three different Lotus transgenic lines have been obtained after transformation with the prGLB1-gusA fusion containing the 980 bp fragment upstream of LjGLB1 ATG start codon. These three transgenic lines have been propagated to isolate homozigous T2 lines with consistent patterns of GUS activity. The analysis of the pLjGLB1-gusA confirms the expression previously described in vascular bundle of transgenic roots (D'Apuzzo et al., 2015), giving the opportunity to identify an additional striking spatial profile of expression in leaf tissues where GUS activity is localized in main and secondary veins but also in mesophyll and guard cells (Figures 1A,B). The GUS staining in guard cells is even better visualized in leaves after epidermis peeling (Figures 1C,D). The analysis of the 980 bp at the 5′ region of the LjGLB1 gene reveals some interesting features consistent with this spatial profile of GUS activity. This region contains eight (A/T) AAAG and three CCAAT elements, binding motifs for Dof zinc finger and NF-WA transcriptional factors (Figure S1) that have been associated to guard cell activity in potato, grapevine and A. thaliana (Li et al., 2008; Galbiati et al., 2011).

A. tumefaciens-mediated transformation-regeneration procedure has been also exploited to obtain Lotus trangenic plants transformed with LjGLB1-GFP fusion driven by the CAMV35S constitutive promoter and confocal laser-scanning fluorescence analysis in T2 transgenic lines indicated unambiguously a chloroplast localization of GFP in leaves, which co-localizes with chlorophyll red autofluorescence (Figure 2), confirming previous data obtained in Arabidopsis and rice (Hsieh et al., 1998; Sugiyama et al., 2004).

The clear-cut pattern of prLjGLB1-gusA activity detected in guard cells prompted us to investigate whether PII could play any role in the control of stomata movement. Therefore, we took advantage of the genetic tool we recently characterized, represented by two transgenic lines ectopically expressing LjGLB1 under the control of the CAMV35S constitutive promoter (D'Apuzzo et al., 2015), to test whether the PII overexpression could affect the guard cell functioning. Epidermal fragments have been prepared from leaves of wild type plants and the two PII-overexpressing 7–13 and 8–9 transgenic lines and analyzed for stomatal movement. We have first analyzed the effect of 10 μM abscisic acid (ABA) on stomatal closure phenotype in epidermal strips of detached wild type and PII-overexpressing transgenic leaves incubated for 3 h on light (300 μmol m⁻² s⁻¹) with or without addition of 10 μM abscisic acid (ABA). These measures indicate, as expected, an ABA dependent triggering of stomatal closure (40%) but no statistically significant differences are revealed in guard cells aperture of detached wild type and PII over-expressing leaves (Figure 3A). Therefore, PII overexpression is not affecting the level of the ABA-induced stomata closure in Lotus plants. Through a different approach, we have then tested the effect of PII overexpression on stomata closure in a time course experiment during a rapid leaf dehydration treatment. Stem cuttings 6–7 cm long, from 4 to 5 weeks old wild type and transgenic plants, carrying 4–5 trifoils, have been isolated at the 6th hour of the daily light period and immediately sealed at the wounded site with instant adhesive glue. Central leaves from corresponding trifoils detached at T0, 1, and 2 h have been utilized for epidermal strips preparation and immediate stomatal measurement. Data shown in Figures 3B,C indicate a progressive stomatal closure triggered by this rapid dehydration treatment in wild type and transgenic leaves but, a clear-cut increase of stomatal closure is observed in 8–9 and 7–13 transgenic leaves compared to wild type leaves and this effect is displayed from the T0 time point (71 and 68% of wild type stomatal aperture, respectively). The leaves for stomata measurement at T0 are harvested immediately after the stem cutting and sealing (less than 3 min), indicating that the detected differences in stomata apertures are likely associated to a constitutive condition of PII overexpressing leaves rather than to a differential early response to drought treatment. It should be also considered that the unchanged measures of stomatal aperture in the detached leaves of control plants not treated with ABA (white bars in Figure 3A) are due, in that experimental set up, to the preliminary treatment of epidermal strips that are exposed 3 h to direct light, inducing full aperture and hence equalizing the stomatal starting conditions.

L. japonicus lines with LORE1 retrotransposon element inserted within the LjGLB1 sequence are not available in the large collection of insertion lines recently released (Fukai et al., 2012; Urbanski et al., 2012; Malolepszy et al., 2016). Therefore, in order to test the possible physiological involvemen of PII protein in the guard cell functioning we have analyzed the stomatal aperture phenotype of an Arabidopsis thaliana knock out mutant already characterized for different phenotypes (Ferrario-Mèry et al., 2005). The T-DNA insertion line N521878 (Salk collection) carrying a double insertion event in the fourth intron of the AtGLB1 gene and previously characterized with an undetectable expression of the AtGLB1 gene (PIIS2 mutant; Forchhammer and Hedler, 1997), has been obtained from the Nottingham Arabidopsis Stock Center (http://arabidopsis.info). We have confirmed by PCR the genotype of homozygous insertion mutants and tested the stomata phenotype by comparison with leaves of wild type A. thaliana Columbia ecotype. This analysis did not reveal any alteration neither of ABA-induced nor constitutive stomata aperture in the PIIS2 plants (Figure S2).

Stomata serve as major gateways both for transpirational water loss and CO₂ influx of plant leaves. Therefore, we have investigated whether the altered stomatal movement observed in Lotus PII overexpressing plants could affect two phenotypes correlated to those physiological functions such as water use efficiency and photosynthetic performances.

In order to test whether the constitutive reduced aperture of stomata in the PII overexpressing leaves induce a favorable response to a rapid dehydration treatment, the stem cuttings material described above, has been used for the analysis of water loss. The bases of stem cuttings are immediately sealed with instant adhesive glue on a microscope cover glass to prevent water loss by gravity and fresh weight measured in the dark, throughout the desiccation treatment. As shown in Figure 4A, both 7–13 and 8–9 transgenic lines display an early significant reduced transpirational water loss in response to this rapid drought stress treatment. In the following analysis, we have tested whether wild type and transgenic PII overexpressing plants display differences in the gradual long term response to cessation of irrigation. Plants have been grown individually in the growth chamber, in pots with soil composed by a mixture of sand/vermiculite and watered with half strength B&D medium supplemented with 5 mM NH₄NO₃. The plants in vegetative growth conditions, either wild type, –9 and 7–13, after 12 days of desiccation showed severe wilting and turned brown with no differences observed in the recovery capacities during re-watering treatments (data not shown). This observation has been quantified through the analysis of leaf Relative Water Content (RWC) that progressively decrease with overlapping curves, in both wt and transgenic leaves during drought stress treatment (Figure 4B).

The detected reduced stomata aperture of PII overexpressing plants is not associated to the display of macroscopic morphological changes when shoot growth parameters are compared in wild type and transgenic plants grown in axenic conditions (D'Apuzzo et al., 2015). We have confirmed this result through the analyses conducted on the normally watered control plants maintained in growth chamber conditions and used for the RWC analysis, as shoot fresh weight doesn't change between wild type and transgenic plants (Figure 5A). This phenotype has been even further investigated through a quantitative analysis of bio-volume parameters performed with a Scanalyzer 3D platform on wild type and transgenic plants, grown in greenhouse conditions and normally watered. Groups of 12 wild type, 8–9 and 7–13 plants have been transferred to pots and grown vegetatively for 50 days under greenhouse conditions. Leaf bio-volume parameters are obtained in a non-destructive way by calculating the sum of the projected areas (pixels) from the two orthogonal side views, during a time span of 10 days (from day 30 to day 40). The values plotted on Figure 5B confirm no significant differences between wild type and transgenic PII overexpressing plants. The plants grown under greenhouse conditions have been further analyzed by taking specific measures of their photosynthetic potential. Under visible light illumination, leaf pigments of the photosystems adsorb the energy of the photons in their electrons by elevating them to higher orbitals. There are three competing fates for this energy: (i) phytochemistry, the productive phase of photosynthesis, (ii) dissipated as kinetic energy, heat, or (iii) florescence, re-emitted as luminous energy at lower energy (Oxborough, 2004). The interplay of these three pathways for the photon energy depends on the state of the plant and normal healthy plants will be consuming the energy in photosynthesis, whereas stressed plants will be dumping the energy as heat or fluorescence. Therefore, leaves fluorescence is a quantitative parameter of their photosynthetic potential and these measures might be utilized for evaluation of the state of plant stress index (Muller et al., 2001). The values of the photosynthetic stress index plotted on Figure 5C and measured at the same time points used for the bio-volume analyses, should be considered only as relative levels when compared to other plants in the same experiment. The bins for the stress index calculation are chosen around the peak of the Hue histogram such that the calculated values, ranging from −1 to +1, indicate whether the median peak has shifted to the left (red shifted, toward −1) or to the right (blue shifted, toward +1) with respect to the wild type control. In this manner plants that are more stressed have higher values with respect to the reference plants. Interestingly, the measures fall all in a positive range of the scale values, confirming the absence of evident stressful conditions neither in wild type nor transgenic plants but nevertheless, both transgenic PII overexpressing lines show a significant higher stress index when compared to wild type plants (Figure 5C).

A possible role of PII in guard cells functioning is particularly intriguing as this protein has been already involved in A. thaliana on the control of NO₂⁻ and arginine content, two main sources of NO synthesis in leaves (Ferrario-Mèry et al., 2005, 2008). Therefore, we have tested whether the content of nitric oxide (NO), a key signal controlling the stomatal closure, is affected in leaves of trangenic overexpressing PII plants compared to wild type. Intracellular NO production has been compared in wild type and 8–9 transgenic leaves by means of the cell-permeable DAF-FM DA fluorescent probe. Peeled leaf epidermal fragments have been obtained from Lotus plants grown in the growth chamber and loaded with the dye. The excess of DAF-FM-DA probe is washed out and the epidermal strips are immediately analyzed by means of wide field fluorescence microscopy. As shown in Figure 6, guard cells of 8–9 leaves have a significantly increase of NO accumulation when compared to wild type cells (about 20%), thus indicating a striking correlation between NO generation and stomatal closure observed in the PII overexpressing transgenic plants.

In order to further characterize the altered biosynthetic pathways leading to the constitutive increased NO content in leaves of L. japonicus PII over-expressing plants (Figure 6), we have focused our attention on nitrate reductase (NR), a crucial enzyme for production of NO₂, an endogenous substrate for NO synthesis in leaf tissues. Nitrate reductase activities have been assayed in leaves of 3 weeks old Lotus plants grown in axenic cultures in the presence of 12 mM KNO₃ and 0.5 mM (NH₄)₂SO₄ as N sources (B5/2 Ganborg medium). NR activity is significantly increased in both 7–13 and 8–9 PII over-expressing lines when compared to wild type plants (Figure 7A; 63 and 70% increased activity, respectively). In a parallel analysis, we have also tested in the same plants used for characterizing the enzymatic activity, the LjNR transcript abundance. The L. japonicus gene coding for NR has been previously identified and characterized at the molecular level (Kato et al., 2003) and the blast analysis against the new version 3.0 of the Lotus japonicus genome database assembly (http://www.kazusa.or.jp/lotus) confirms the Lj0g3v0006719.1 as the unique LjNR gene. Data reported on Figure 7B show no significant differences of LjNR expression between wild type and PII over-expressing plants, indicating that overexpressed PII protein affects only NR enzymatic activity. The same analyses have been also conducted on leaves of plants grown in soil under growth chamber conditions with identical results (data not shown).

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.