UV-B Radiation as a Novel Tool to Modulate the Architecture of In Vitro Grown Mentha spicata (L.)

Abstract

In vitro culturing can generate plants with a distorted morphology. Some distortions affect the plant’s survival after transfer to an ex vitro environment, while others can affect the aesthetic value. Therefore, exogenous hormones are often applied in in vitro cultures to modulate plant architecture. In this study, it was hypothesised that regulatory effects of UV-B radiation on plant morphology can be exploited under in vitro conditions, and that UV exposure will result in sturdier, less elongated plants with more branches and smaller leaves, mediated by changes in plant hormones. Plants were grown in tissue-culture containers and exposed to ~0.22 W m⁻² UV-B for 8 days. Subsequently, plants were transferred to soil and monitored for a further 7 days. Results show that UV induced a marked change in architecture with a significant increase in axillary branches, and reductions in leaf area, plant height and root weight. These changes were associated with significant alterations in concentrations of hormones, including IAA, GA7, GA3 and iP–9–G. Changes in hormone concentrations suggest a regulatory, rather than a stress response to UV-B. Therefore, it is proposed that the application of UV in in vitro culture can be an innovative approach to manipulate plant architecture.

1. Introduction

Since its inception in the early 1900s, plant tissue culture or plant in vitro culture has developed into a commercially important propagation method that is widely applied by the horticultural industry [1]. In vitro propagation may refer to the cultivation of protoplasts, suspensions of individual cells, callus, explants from various plant organs, embryos or seeds. The commonality between these distinct cultures is the controlled environment, which is typically aseptic and characterised by a high relative humidity, constant temperature, and media enriched with nutrients and/or plant growth regulators [2]. The use of in vitro techniques facilitates rapid clonal propagation, with a high success rate, which in many cases cannot be achieved under less controlled conditions and/or on soil [3]. However, as a consequence of imposing these controlled and unnatural environmental conditions, in vitro raised plants are characterised by physiological, anatomical and morphological adjustments, that differentiate these plants from those grown using non-in vitro conditions [4,5]. Generally, in in vitro-generated plants, leaves are thinner, stomatal opening and epicuticular wax deposition are altered, the photosynthetic apparatus is not well developed, and the vascular system is not efficient in transferring water and assimilates. Some of these alterations make plants more vulnerable to stress when they are transferred to an ex vitro environment [6]. To stimulate development of a less aberrant morphology, media have been supplemented by adding exogenous plant hormones (plant growth regulators) such as 2,4-Dichlorophenoxyacetic acid (2,4-D) and/or 6-benzyl adenine (BA) [7,8,9]. Furthermore, for the same reason it has also been attempted to modulate the in vitro environment through alterations in temperature, humidity, and/or light [4,10,11]. Light in particular has a strong influence on the physiology and morphology of in vitro raised plants, and this effect depends both on light quantity and quality (i.e., spectrum). Light can affect a variety of physiological processes such as the development of photosynthetic competence, accumulation of secondary metabolites, cell wall development, and plant and leaf morphology [12]. While the importance of the light spectrum for in vitro culture is well established [12,13], knowledge is largely limited to visible wavelengths (380–700 nm). In contrast, very limited information is available on the influence of ultraviolet A (315–400 nm) and B (280–315 nm) wavelengths. These wavelengths are part of the natural solar spectrum, and plants have evolved to sense and respond to such wavelengths using dedicated photoreceptors and signalling cascades [14]. For example, UV-B wavelengths are sensed through the UVR8 photoreceptor, while both phototropin and cryptochrome photoreceptors are active in the UV-A part of the spectrum. UV-B wavelengths in particular can trigger a wide array of plant responses, including accumulation of various metabolites, increases in antioxidant defences and changes in plant morphology [15,16,17]. Indeed, it has been argued that UV-B radiation can be used as a method to control plant morphological parameters [18]. Noted alterations in the architecture of UV-B-exposed plants include leaf thickening and a reduction in leaf area, shortening of petioles, a reduction in stem length, increased axillary branching and alterations in root morphology [18,19,20,21,22]. While the mechanisms underlying such morphological responses are not fully understood, it is accepted that the photoreceptor UVR8 can play a role in this process. UVR8-mediated signalling can affect hormone-related signalling, ultimately impacting on organ differentiation and causing UV-induced morphological changes [23]. UV-B affects hormonal pathways through two possible mechanisms. The first one involves UVR8-mediated effects on growth-related hormones such as auxins, cytokinins (CK) and gibberellins (GA). The second pathway involves a more generic UV-induced stress response, altering concentrations of defence hormones such as abscisic acid (ABA) [23,24]. This study explored whether UV radiation can be exploited as a tool for the in vitro culture of plants. We specifically asked the question whether UV radiation can be used to modulate plant morphology under sterile, tissue culture conditions. To develop such an innovative application of UV radiation, we used UV-transmitting containers, and used sterile, seed-based culture of mint (Mentha spicata L.) as a model system. It was hypothesised that the well documented effect of UV-B radiation on plant morphology can be exploited under in vitro conditions, and that UV exposure would result in a sturdier, less elongated plant with more branches and smaller leaves, mediated by changes in plant hormones.

2. Materials and Methods

2.1. Experimental Set-Up

2.1.1. Plant Material and Germination Stage

Mint (Mentha spicata L.) seeds were obtained from a commercial grower (Moles Seeds Ltd., Stanway, UK). Seeds were surface sterilised by incubating in 0.05% Triton X-100 for 5 min, followed by 10 min in 50% sodium hypochlorite in 0.05% Triton X-100. Afterwards, the seeds were rinsed in distilled water. Sterilised seeds were incubated for 3 days at 4 °C, then 20 seeds per box were sown in solid Murashige and Skoog medium without added sucrose (MS) without hormones and grown for 30 days in plastic boxes (RA40 plastic micro boxes, Sac 0₂, Deize, Belgium) in a growth room under 180 μmol m⁻² s⁻¹ PAR (Figure 1).

2.1.2. UV—Exposure Stage

Prior to UV-treatment, the aforementioned boxes were randomly divided into 2 groups and the lids were replaced with either a UV-B blocking filter Mylar (125 µm thickness, Polyester film, Tocana Ltd., Ballymount, Ireland) or a UV-B transmitting Cellulose Acetate (CA) filter (95 μm thickness; Kunststoff-Folien-Vertrieb GmbH, Hamburg, Germany). Furthermore, the external sides of the boxes were covered with Mylar or CA. Filters were previously sterilised with 70% ethanol and changed after every 20 h of UV exposure (Figure 1). All plants, UV-exposed and non-UV-exposed, were kept in the same room.

2.1.3. Recovery Stage

After 8 days of UV exposure, a random subset of the plants was harvested, while another subset was transplanted to John Innes II compost (William Sinclair Horticulture Ltd., Lincoln, UK) and moved to a growth room. Conditions in the growth room (i.e., PAR, temperature, and relative humidity) were similar to those applied during the seedling growth and UV-exposure phases. Plants were harvested for analysis 7 days after transplantation to soil conditions (Figure 1).

2.1.4. Light, Humidity and Temperature Conditions

Plantlets were exposed to ~180 μmol m⁻² s⁻¹ PAR background provided by LED lamps (AP673L, Valoya, Finland), at a temperature of 20 ± 2 °C and ~50% relative humidity. FAR-RED LEDs tubes (L18C, Valoya, Finland) were used to achieve the R: FR ratio of 1.6. The light regime was set up for 14 h light/10 h dark for PAR (7:00 to 21:00) and 16 h light/8 h dark (06:00 to 22:00) for FAR- RED. UV-B radiation was provided by fluorescence tubes (TL20W/12 Philips, Germany) wrapped with a single layer of CA to block the UV-C radiation. Each sheet of CA was changed after 20 h of UV exposure. Boxes were exposed to ~0.5886 W m⁻² total UV (with 0.3661 W m⁻² UV-A and ~0.2225 W m⁻² UV-B) under the CA filter and ~0.2498 W m⁻² total UV (with 0.2464 W m⁻² UV-A and 0.004 W m⁻² UV-B) under Mylar for 4 h around mid-day (12:00 to 16:00). These irradiances are several fold lower than would be observed at noon in the summer in the temperate zone. The daily biologically effective UV-dose was equivalent to 2.7937 kJ/m² (0.2830 kJ/m² for UV-A and 2.5258 kJ/m² for UV-B) under CA and 0.3250 kJ/m² (0.1797 kJ/m² for UV-A and 0.1453 kJ/m² for UV-B) [25]. The light spectra, the UV intensity and the Red: Far Red ratio were determined using an optical fiber spectroradiometer Flame-S equipped with a cosine corrector (Ocean Optics, Duiven, The Netherland) using the manufacturer’s software, Oceanview (version 1.6.7). The PAR intensity was measured using a PAR meter (PAR special sensor, Skye Instrument Ltd., Powys, United Kingdom). The R: FR ratio was calculated as photon irradiance between 655 and 665 nm/photon irradiance between 725 and 735 nm [26]. A portable data logger (EL-USB-2, Lascar Electronics, Whiteparish, United Kingdom) was used to determine and monitor the temperature and the humidity.

2.2. Biometric Analysis

All morphological parameters were measured at two different stages: (1) immediately after the UV exposure stage and (2) after the recovery stage. For each stage, 6 plants per treatment were evaluated. During the measurements leaf 1 (L1) is the oldest leaf and progressively leaf 6 (L6) is the youngest leaf. Parameters measured included the total number of leaves, number of branches, number of leaves on branches, stem thickness, plant height, internode length, leaf area, dry weight, and specific leaf area. Stem thickness was measured using a calliper at the soil level, while plant internode length was calculated as plant height divided by the number of leaf pairs on the main stem. Total leaf area (TLA) was measured based on photographs of the first six leaves and processed using ImageJ software (version 1.52a) (Wayne Rasband, National Institute of Health, United States). Leaf dry weight (LDW) for the oldest six leaves was recorded after drying leaves at 60 °C for 5 days. Specific Leaf Area (SLA) for the oldest six leaves was calculated as a ratio between leaf area and leaf dry weight (LA/LDW). Leaf area, leaf dry weight and leaf specific area were also calculated for individual leaves, albeit pooling together L1–L2, L3–L4 and L5–L6. The results were obtained from 5 independent replicates.

2.3. Root Morphology

Root morphology refers here to the number and length of the primary roots and the number and length of the secondary roots. These were measured with the help of SmartRoot [27] a plugin of ImageJ software (version 1.52a) (Wayne Rasband, National Institute of Health, United States). Root fresh weight was recorded at the end of each experimental stage after the removal of the stems and agar or soil residuals. Root dry weight was recorded after drying roots at 60 °C for 5 days. Analyses were conducted directly after the UV exposure stage and once more at the end of the recovery stage on 6 samples for each stage and on 3 biological replicates.

2.4. Hormone Analysis

Hormone concentrations were measured in different plant organs: leaves, stems and roots. For each replicate, material from different plants was pooled to reach the necessary amount of 100 mg per organ. Each measurement was independently replicated (i.e., independent plants, SPE extraction and chromatographic quantification for each aliquot) at least 5 times. After collection, plant material was immediately frozen in liquid nitrogen and stored at −80 °C until the extraction. Data were collected after the UV-exposure stage and after the recovery stage. The extraction procedure follows Qian et al. [21] with some modifications. Approximately 100 mg of frozen and homogenised plant material was mixed with 1 mL extraction solvent (80% (v/v) methanol (HiPerSolv CHROMANORM^®; VWR, Leuven, Belgium), 20% 0.8 mM formic acid (EMSURE^®; Merck, Darmstadt, Germany)) and extracted overnight at −20 °C. The following internal standards were added: 30 pmol 3-[phenyl-¹³C₆] indolylacetic acid (Cambridge Isotopes, Tewksbury, MA, USA), 30 pmol [²H₆](+)-cis, trans-abscisic acid, ([(S)-5-[²H₆](1-hydroxy-2,6,6-trimethyl-4-oxocyclohex-2-en-1-yl)-3-methyl-(2Z,4E)-pentadienoic acid] (Olchemim Ltd., Olomouc, Czech Republic), 30 pmol for each GA (²D₄-Gibberellin A1, ²D₄-Gibberellin A4, ²D₂-Gibberellin A8, ²D₂-Gibberellin A9, ²D₂-Gibberellin A15, ²D₂-Gibberellin A19, ²D₂-Gibberellin A20 (Olchemim Ltd.)), and 40 pmol each for the cytokinins ([²H₃]dihydrozeatin, [²H₃] dihydrozeatin riboside, [²H₅]trans-zeatin-7-glucoside, [²H₅]trans-zeatin-9-glucoside, [²H₆]N⁶-isopentenyladenine, [²H₆]trans-zeatin-O-glucoside, [²H₆]trans-zeatin-O-glucoside riboside, [²H₆]N⁶-isopentenyladenosine, [²H₆]N⁶-isopentenyladenine-7-glucoside, [²H₆]N⁶-isopentenyladenine-9-glucoside, (Olchemim Ltd.)), ([²H₇]N⁶-benzyladenine (D-BA), [²H₇]N⁶-benzyladenosine (D-BAR), [²H₇]N⁶-benzyladenine-9-glucoside (D-BA9G), ([¹⁵H₄]meta-topolin (15N-mT), [¹⁵N4]ortho-topolin (15N-oT), Olchemim Ltd.)). After overnight extraction, Oasis^® HLB (10 mg, Waters, Deerfield, IL, USA) was added in batches to bind pigments. After centrifugation (Eppendorf 5810R 14,000 rpm, 4 °C, 15 min; Eppendorf, Hamburg, Germany) to remove cell debris and Oasis sorbent, the extract was split with separate aliquots for the analysis of cytokinins (aliquot 1), indole-3-acetic acid, abscisic acid, gibberellins (aliquot 2), and IAA-conjugates (aliquot 3).

Aliquot 1 was filtered through a Chromafil^® AO-20/3 filter (nylon, pore size 0.20 μm, diameter 3 mm; Macherey-Nagel, Düren, Germany). The filtrate was collected in an LC-MS vial and kept at 4 °C. Isoprenoid cytokinins were analysed in aliquot 1 by ACQUITY UPLC coupled ES(+) TQD Tandem Quadrupole UPLC/MS/MS (Waters, Deerfield, IL, USA) in multiple reactions monitoring (MRM) mode using a BEH C18 Column (130 Å, 1.7 µm, 2.1 mm × 50 mm) coupled to a BEH C18 VanGuard Pre-column (130 Å, 1.7 µm, 2.1 mm × 5 mm), in combination with the following linear gradient: 95:5 A:B with A 0.1 M ammonium acetate (Molecular biology grade 7.5 M, Merck Life Science, Hoeilaart, Belgium) in water (HiPerSolv CHROMANORM^®; VWR, Leuven, Belgium) and B Methanol, isocratic 95:5 A:B for 0.4 min, 95:5 A:B to 76:24 A:B in 3 min, isocratic 76:24 A:B for 2 min, flow 0.39 µL min⁻¹, column temperature 50 °C, injection volume 6 µL. Aromatic cytokinins were analysed in a separate UPLC-TQD MS/MS run with the same analytical column at: 95:5 A:B with A 0.1M ammonium acetate in water and B Methanol, 99.9:0.1 A:B to 95:5 A:B to 72:28 A:B in 5 min, isocratic 72:28 A:B for 0.5 min, linear gradient 72:28 A:B to 0.1:99.9 A:B in 0.5 min, flow 0.4 µL min⁻¹, column temperature 48 °C, injection volume 6 µL.

Aliquot 2 was acidified with 5 mL 6% formic acid and loaded on a pre-conditioned Bond Elut C18 solid phase extraction cartridge (Agilent, Santa Clara, CA, USA). Compounds of interest were eluted with 4 mL of diethyl ether (AnalaR NORMAPUR^®, VWR, Leuven, Belgium). The eluent was dried under a nitrogen stream, dissolved in 60 µL methanol and derivatised with N-(3-Dimethylaminopropyl)-N′-ethyl carbodiimide, and kept at 4 °C until analysis. IAA, ABA and GA were analysed by ES(+)-UPLC-MS/MS in MRM mode, using the same combination of column and guard detailed for aliquot 1, and using the solvent gradient: 92:8 A:B with A 0.1% (v/v) FA in water and B 0.1% (v/v) FA in ACN to A:B 60:40 for 5 min under a linear gradient, followed by a linear gradient from 60:40 to 10:90 in 0.5 min, flow rate 0.420 µL min⁻¹, column temperature 40 °C.

Aliquot 3 was mixed with an equal volume of 12N NaOH, flushed with a water saturated nitrogen stream for 20 min and hydrolysed for 180 min at 100 °C in a Pierce Reacti-Therm III. After alkaline hydrolysis, the sample was titrated with 2M HCl and diluted to a final volume of 10 mL with deionised water. Further solid phase extraction and derivatisation are as described for aliquot 2. IAA conjugates were analysed as IAA-EDC using the chromatographic conditions described for IAA. All data are expressed in picomoles per gram fresh weight (pmol g⁻¹ FW).

2.5. Statistical Analysis

Statistical analyses were performed using IBM SPSS Statistics v28 (Armonk, New York, NY, USA). 2 tails t-test or Mann-Whitney test were used to determine the difference between +UV and −UV after the UV-exposure stage or after the recovery stage, separately. Data are shown as mean ± SE (Standard Error).

3. Results

3.1. Plant Morphology

M. spicata plants were grown in closed containers, morphological and biomass parameters were measured immediately after 8 days of UV exposure and again 7 days after transplanting to soil (Figure 2). The total number of leaves, which includes the leaves on branches, showed a significant UV-induced increase, both after the UV exposure phase (p = 0.02) and at the end of the transplanting phase (p < 0.001) (Figure 3A). A highly significant change in the number of branches and the number of leaves on branches was also observed. Thus, UV-treated plants developed a higher number of branches during UV exposure (p < 0.001) and this number further increased significantly during the following 7 days when the plants were not exposed to UV (p < 0.001) (Figure 3B,C). Moreover, the stem thickness decreased due to UV radiation exposure. Stems of the UV-exposed plants were significantly thinner after both UV exposure and recovery stages (p < 0.001) compared to the −UV plants (Figure 3D).

UV exposure leads to a reduction in plant size, indeed plants exposed to UV are significantly shorter than plants grown under the UV-blocking filter after both the UV exposure stage and the recovery stage (p < 0.001) (Figure 2A and Figure 4A). UV also affects the internode length causing a reduction both immediately after UV exposure (p < 0.001) and after transplanting (p < 0.001) (Figure 4B).

The total leaf area across all the main leaves on the plant showed a significant decrease for the UV-treated plants compared to −UV plants immediately after the UV exposure phase (p < 0.001). A significant decrease in total leaf area was also measured after transplanting to soil (p < 0.001) (Figure 2B and Figure 5A). The same trend was recorded for leaf dry weight where the change between the UV-exposed plants and the non-UV-exposed plants was both significant immediately after the UV exposure stage and after transplanting (p = 0.033, p = 0.001) (Figure 5B). On the contrary, the SLA does not show any significant variance after the UV-exposure stage, but the difference becomes both substantial and significant after the recovery stage (p = 0.001) (Figure 5C).

Leaf area, leaf dry weight and specific leaf area were also analysed for individual pairs of leaves, i.e., L1–L2, L3–L4, L5–L6. The leaf area of the oldest pair of leaves, L1–L2, was relatively similar for UV-treated and non-UV-treated plants immediately after the UV-exposure stage. In contrast, there was a significant decrease in the area of L2–L3 (p < 0.001) and L4–L5 (p = 0.023) in UV-exposed plants (Figure 6A). A comparison of leaves analysed after UV exposure, and those measured after recovery, showed that leaves did continue growing during the transplanting phase. After the recovery, the leaf area was significantly smaller among all the leaves that had previously been exposed to UV (p < 0.001, p < 0.001, p < 0.001) (Figure 6B).

In contrast, UV does not affect the leaf dry weight after the UV exposure stage, weights remaining relatively similar for L1–L2, and not significantly decreasing for L3–L4 and L5–L6 (Figure 7A). The same trend was recorded after transplanting with the same relative leaf dry weight for the first pair of leaves, irrespective of previous UV-exposure. However, a UV-induced decrease in leaf dry weight was noted for the second and third pair of leaves following recovery. This reduction was significant for L3–L4 (p = 0.001) but not significant for L5–L6 (Figure 7B).

Specific leaf area rises with UV exposure for all pairs of leaves with a progressive increase from the oldest to the youngest leaves, but the change was not significant at the end of the UV exposure stage (Figure 8A). After transplanting, the SLA recorded for previously UV-exposed plants tends to be lower than that of non-UV-exposed, decreasing from L1 to L6. This reduction is significant only for the youngest leaves L5–L6 (p = 0.006) (Figure 8B).

3.2. Root Morphology

Root fresh weight was significantly lower after UV exposure (p = 0.031). The difference is most marked after transplanting when the roots of non-UV-exposed plants are much heavier than the roots of the UV-treated plants (p < 0.001). In general, the change in fresh weight is larger in UV-exposed plants compared to non-UV-exposed plants, when comparing plants immediately after UV-exposure and plants transplanted to soil (Figure 9A). Root dry weight was similar across the treatments immediately after the UV exposure stage. However, the difference in dry weight becomes significant after the recovery stage with a higher root biomass in non-UV-exposed plants (p = 0.003) (Figure 9B).

The number of primary and secondary roots is not affected by UV exposure. Indeed, no differences in root numbers were recorded between UV-exposed and non-UV-exposed plants at either stage. An increase in secondary roots was noted after transplanting for both sets of plants (Figure 10A,B). Immediately after the UV-exposure stage, the primary roots were significantly shorter in UV-treated plants compared to non-UV-treated plants (p = 0.011). During the recovery stage, the primary roots of UV-exposed plants kept growing while the primary roots of non-UV-exposed plants did not elongate further. As a result, there was no significant difference in primary root length between the UV-treated and non-UV-treated (Figure 10C). UV also affected the length of the secondary root, and these are significantly shorter in UV-exposed plants (p = 0.006). In contrast to the main roots, secondary roots from non-exposed plants grow strongly after transplanting, while the UV-exposed ones do not grow when transplanted into the soil. The analysis revealed that this change is not significant after the recovery stage (Figure 10D).

3.3. Hormone Profile

3.3.1. Leaves

In order to determine the mechanism underlying the effect of UV-B on different plant organs, the concentration of different hormones was evaluated in leaves, stems and roots both immediately after UV exposure and 7 days after transplanting.

Most of the hormones measured immediately after UV exposure showed a decreasing trend, i.e., concentrations were lower in UV-exposed leaves. In detail, relative to the untreated plants, concentrations of BAP (−28%), BA-9-G (−61%), iP-9-G (−56%) (Figure 11A), ABA (−45%) (Figure 11D), IAA (−69%) (Figure 11G), GA3 (−30%) (Figure 11J), GA7 (−43%) were all decreased following UV treatment. Especially, the concentration of IAA–conjugates (IAA-C) significantly diminished in the UV—exposed plants (−99%, p = 0.016). Concentrations of iPA, GA19 and GA15 increased, but this was not significant. At the end of the recovery stage, similar trends were recorded as immediately after UV exposure. The concentration of BA-9-G and iPA in UV-exposed leaves were even below the detection limit. Decreased concentrations in UV-exposed leaves were measured for BAP (−25%), ABA (−78%) (Figure 11D), IAA-C (−74%), and GA19, but the difference is statistically different only for iP-9-G (−51%, p = 0.038) (Figure 11A) and GA3 (−63%, p = 0.009) (Figure 11L). IAA (Figure 11G) and GA7 slightly increased respectively by 10% and 34% (Supplementary Table S1).

3.3.2. Stems

The concentrations of most hormones measured in the stem compartment tended to decrease in plants exposed to UV, immediately after the end of the UV treatment. As for the leaves, the concentrations of iP-9-G (−20%) (Figure 11B), ABA (−82%) (Figure 11E), IAA (−77%) (Figure 11G), IAA-C (−96%), GA3 (−5%) (Figure 11K) were lower in stems of UV-exposed plants but not in a significant way. GA19 concentrations also slightly decreased (−2%) in stems, unlike leaves where a strong increase in concentration was observed. Concentrations of BAP (+22%), BA-9-G (+15%) iPA (+545%) and GA19 (+28%) slightly increased and of GA7 significantly increased (+79%, p = 0.016) in the stems. After the recovery stage, concentrations of BAP (−6%), iPG9 (−28%) (Figure 11B), ABA (−62%) (Figure 11E), IAA-C (−35%), GA3 (−15%), BAG9 (−23%) and iPA (−20%) were found to decrease in the stems of the plants previously exposed to UV. However, IAA (+11%) (Figure 11H) and the majority of gibberellin concentrations GA7 (+14%), GA15 (+28%), GA19 (8%) measured increased in + UV plants (Supplementary Table S2).

3.3.3. Roots

The effects of UV on hormone concentrations were also determined for the below-ground organs. As per the leaves and the stems, UV negatively influences concentrations of some hormones such as iPG9 (−50%) (Figure 11C), ABA (−51%) (Figure 11F), GA3 (−19%) (Figure 11L), BAP (−35%) and GA15 (−39%) in the roots. This effect is significant for both IAA (−91%, p = 0.032) (Figure 11I) and IAA-C (−96%, p = 0.008), when measured immediately after the UV exposure. In contrast, concentrations of BA-9-G (+102%), GA19 (+11%) and GA7 (+1%) increased in roots of UV-exposed plants. Measurements on roots 7 days after transplanting confirmed a reduction in GA3 (−26%, p = 0.026), GA15 (−6%), BAP (−26%), iPA (−35%) and iP−9-G (−19%) (Figure 11C), ABA (−51%) (Figure 11F), IAA-C (−70%) in roots of plants exposed to UV-B. The trend is similar to that seen in UV-exposed leaves and stems. A parallel increase in IAA (+34%) (Figure 11I) and GA7 (+6%) concentration was detected (Supplementary Table S3).

4. Discussion

In this study, we found that UV induces morphological adjustments in plants exposed to UV. This adjustment persists for some time, even when the UV exposure is ended, and is associated with changes in the concentrations of several growth regulating plant hormones.

4.1. UV- Induced Changes in Morphology and Possible Applications for In Vitro Culturing

UV-B has been reported to induce morphological responses in plants [18]. For example, it has been reported a dwarfed phenotype of cucumber, with a reduction in leaf area, petiole length and stem elongation, following UV-exposure [21]. Here, we observe that similar morphological responses can be induced under in vitro conditions. The results show that the oldest leaves respond weakly to UV, while younger leaves are strongly affected (Figure 6). This may occur through the activation of a specific signalling cascade initiated by the UVR8 photoreceptor [28]. Alternatively, the response could be a generic, so-called stress-induced morphogenic response (SIMR) [29]. SIMRs have been characterised as redistribution, rather than cessation of growth, and this can be seen in the observed phenotype. On one side, UV promotes plant growth seen as an increase in axillary branching; on the other side, UV acts as a growth inhibitor seen as the reduction of stem elongation and leaf elongation [29]. This phenotype is recognised as typical for plants acclimated to low doses of UV-radiation [15,18]. Thus, it is concluded that environmentally relevant UV-doses can be used under in vitro conditions to similarly adjust plant morphology. Overall, the UV-induced plant architecture may not only make for a commercially more attractive plant, but may also contribute to a better acclimation to the new growth conditions, for example under ex vitro conditions [4]. The data observed in this study are consistent with work by Metwally et al. [30] who showed that a daily UV exposure (15–45 min) of in vitro spathiphyllum explants not only increased the survival rate of the explants but also contributed to the reduction of plant elongation. Thus, the observed UV-induced reductions in plant elongation are not limited to mint, but seem to apply to a wider range of species. A more compact phenotype can be directly beneficial for ornamental plants, yet can also help to reduce transport costs for a wider variety of crops.

4.2. Link Hormone Changes to Morphological Changes

In this study, architectural changes were accompanied by marked changes in the concentrations of several phytohormones. For example, it is noted that increased axillary branching is accompanied by increased concentrations of cytokinin (BAP, BA9G and iPA) in the stem and strongly decreased concentrations of auxin (IAA). This observation supports or is consistent with the long-established antagonistic relationship of auxin and cytokinin in controlling axillary buds [31]. Typically, endogenous cytokinin concentrations tend to decrease when a plant is exposed to abiotic stress [32], thus the slight increase in cytokinin in the stem implies that the low doses of UV used in this study have a regulatory rather than a stress-inducing effect. Decreases in BAP in other organs may be linked with the quenching of oxyradicals by cytokines in response to UV -B exposure [33]. The observed changes in auxin are consistent with the observations of Hectors et al. [34], who reported a decreased IAA concentration in the leaf compartment after the UV exposure. Here we show both a significant decrease in free and conjugated IAA. The decrease in auxin is not limited to the leaf but is also noted in the stem, where it may be associated with increased axillary branching [35]. Additionally, a decrease in auxin was recorded in the roots where it may be associated with decreased root elongation [35]. The changes in auxin concentration are likely to be linked to the fundamental mechanism of UV-B responses. UV-B perceived via UVR8 and mediated via the HY5/HYH dependent pathway alters auxin synthesis (e.g., PIFs), transport (e.g., PIN proteins) and signalling (e.g., AXR2/IAA7) [23]. In the experiment, the altered auxin distribution after UV exposure is also paralleled by decreases in leaf area, and stem elongation. Effects of auxin on leaf area are consistent with the role of this hormone in leaf development [36], while effects on stem length are well documented, particularly in the context of the shade-avoidance response [37]. Simultaneous, UV-mediated decreases in GA3 are likely to have further slowed down elongation of leaf, stem and root. Finally, the amount of ABA in mint plants is reduced but is not significantly affected by the treatment across all the organs. A possible explanation can be found in the in vitro environment to which the plants were exposed. As reported by Hronková et al. [24], the closed container and consequently the high relative humidity can limit the production of ABA. ABA concentrations are often reported to increase under conditions of plant stress, where ABA can play a key role as regulator [38]. The observation that UV-does not induce increases in ABA, re-confirms the hypothesis that UV-B is a regulator rather than a stressor. Overall, it is concluded that subtle, regulatory differences in concentrations of key hormones such as cytokinin, gibberellic acid and auxin occur in UV-B-exposed plants, and it is hypothesised that these lead to a re-adjustment of the plant architecture under in vitro conditions. Although the link between changes in hormone concentrations and specific architectural adjustments is not conclusive, data give an insight into potential underlying mechanisms of UV-driven architectural adjustment.

5. Conclusions

In this paper, we show that UV supplementation during in vitro culture led to an alteration of plant morphology, characterised by a more compact phenotype and an increase in axillary branching. Thus, it is shown that UV-B supplementation is an alternative and innovative approach to alter plant morphology and obtain the desired plant architecture under in vitro conditions, without the application of exogenous PGRs.