*Article* **A Possible Mode of Action of Methyl Jasmonate to Induce the Secondary Abscission Zone in Stems of** *Bryophyllum calycinum***: Relevance to Plant Hormone Dynamics**

**Michał Dziurka 1,\*, Justyna Góraj-Koniarska 2, Agnieszka Marasek-Ciolakowska 2, Urszula Kowalska 2, Marian Saniewski 2, Junichi Ueda <sup>3</sup> and Kensuke Miyamoto 4,\***


**Abstract:** Plants can react to environmental stresses through the abscission of infected, damaged, or senescent organs. A possible mode of action of methyl jasmonate (JA-Me) to induce the formation of the secondary abscission zone (SAZ) in the stems of *Bryophyllum calycinum* was investigated concerning plant hormone dynamics. Internode segments were prepared mainly from the second or third internode from the top of plants with active elongation. JA-Me applied to the middle of internode segments induced the SAZ formation above and below the treatment after 5–7 days. At 6 to 7 days after JA-Me treatment, the above and below internode pieces adjacent to the SAZ were excised and subjected to comprehensive analyses of plant hormones. The endogenous levels of auxin-related compounds between both sides adjacent to the SAZ were quite different. No differences were observed in the level of jasmonic acid (JA), but the contents of 12-oxo-phytodienoic acid (OPDA), a precursor of JA, and *N*-jasmonyl-leucine (JA-Leu) substantially decreased on the JA-Me side. Almost no effects of JA-Me on the dynamics of other plant hormones (cytokinins, abscisic acid, and gibberellins) were observed. Similar JA-Me effects on plant hormones and morphology were observed in the last internode of the decapitated growing plants. These suggest that the application of JA-Me induces the SAZ in the internode of *B. calycinum* by affecting endogenous levels of auxinand jasmonate-related compounds.

**Keywords:** auxin-related compound; *Bryophyllum calycinum*; indole-3-acetic acid; methyl jasmonate; plant hormone dynamics; secondary abscission

### **1. Introduction**

Plants encounter plentiful biotic and abiotic stresses, leading to shedding (separation) of no longer needed or damaged organs such as leaves, branches, flowers, and fruits, from the parent plants. This process is known as abscission, and it is strongly associated with plant growth and development [1–9]. In the process of abscission, mechanical weakening of cell walls at the abscission zone is brought about by the degradation of the middle lamella by multiple cell-wall-degrading enzymes such as cellulase, polygalacturonases, pectin methyl esterases, and so forth, resulting in shedding [4,9–15].

The position and the time of the formation of abscission zones are determined genetically in each organ, and abscission zones once formed commonly do not differentiate further. Contrarily,

**Citation:** Dziurka, M.; Góraj-Koniarska, J.; Marasek-Ciolakowska, A.; Kowalska, U.; Saniewski, M.; Ueda, J.; Miyamoto, K. A Possible Mode of Action of Methyl Jasmonate to Induce the Secondary Abscission Zone in Stems of *Bryophyllum calycinum*: Relevance to Plant Hormone Dynamics. *Plants* **2022**, *11*, 360. https://doi.org/10.3390/ plants11030360

Academic Editor: Tae-Hwan Kim

Received: 14 December 2021 Accepted: 25 January 2022 Published: 28 January 2022

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**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

in response to tissue injury or infection, differentiation of abscission zones in abnormal positions on stems, petioles, pedicles, and branches, designated as the secondary abscission zone (SAZ), can occur in vivo [1,16]. The secondary abscission has been observed primarily in various in vitro systems involving pedicels of *Malus sylvestris* [17] and *Pyrus communis* [18], stems of *Impatiens sultani* [10,11,19], *Morus alba* [20], *Citrus sinensis* [21], and *Phaseolus vulgaris* [16], and petiole explants of *P. vulgaris* [22], *Pisum sativum* pedicle, or *Euphorbia pulcherrima* flower [23].

The SAZs are induced by some signals, especially plant hormone cues, between neighboring cells [8,24–26]. According to histological analyses, the formation of the SAZ in the stems of *Bryophyllum calycinum* was characterized by the presence of newly synthesized cell plates resulting from periclinal cell division within one layer of mother cells in stems [27].

Plant hormones are well known to play an essential role in plant growth and development, including the abscission or induction of transdifferentiation in mature cortical cells. Ueda et al. [28,29] reported that jasmonic acid (JA) and methyl jasmonate (JA-Me) as senescence-promoting substances promoted the abscission of bean petiole explants. JA-Me also promotes leaf abscission in intact *Kalanchoe blossfeldiana* [30] and *B. calycinum* plants [31]. Furthermore, Saniewski et al. [31] have reported that JA-Me at a concentration of 0.5% (*w*/*w*) applied as a lanolin paste in different stem explants or the debladed petiole induced the formation of the SAZs in *B. calycinum*. These suggest that JA and JA-Me (designated as jasmonates, JAs) have a powerful effect of inducing the SAZ and developing an abscission zone that has already been initiated in plant tissues, resulting in leaf abscission. Ito and Nakano [32] have suggested that a decrease in auxin levels might be considered to provide the first signal for abscission in pedicel abscission in tomatoes. In the stem of *B. calycinum,* indole-3-acetic acid (IAA) applied to a decapitated shoot or internode explants totally prevented the formation of the SAZ in the stems induced by JA-Me [31,33]. However, it should be mentioned that only IAA application substantially induces the formation of the SAZ not only in internode explants, petiole segments, and the petiole after excision of the leaf blade but also in decapitated stems in intact plants of *B. calycinum* [31,33]. It is suggested that in mechanisms of the SAZ formation induced by exogenously applied IAA in the internode of *B. calycinum*, an auxin gradient is vital, and the gradient results from polar IAA transport from the application site [27,31,33]. However, those phenomena induced by JA-Me have not been reported.

As mentioned earlier, plants belonging to the Crassulaceae family show fascinating phenomena, such as leaf abscission and secondary abscission zone formation, easily inducted. This was the reason we chose for experiments the important medicinal plant *Bryophyllum calycinum* (syn. *Kalanchoe pinnata*) [34].

To clarify JA-Me's possible mode of action to induce the formation of the SAZ in terms of its plant hormone dynamics, we focused on differences in plant hormone dynamics between adjacent tissues to the SAZ induced by JA-Me in stem segments, as well as decapitated growing plants of *B. calycinum.* In this paper, comprehensive analyses of plant hormones in JA-Me treated stems, mainly internodes segments, of *B. calycinum* were reported.

#### **2. Results**

#### *2.1. The Effect of JA-Me on Induction of the Secondary Abscission in Internode Segments and Decapitated Plants of Bryophyllum calycinum*

In *Bryophyllum calycinum*, JA-Me application (0.5%; *w*/*w* in lanolin) to the middle of internode segments induced formation of the SAZ, observed at length from 0.5 to 2 cm above and below the JA-Me treatment, 5–7 days after the treatment (Figure 1). JA-Me application induced senescence or loss of chlorophylls in the internode segments in both acropetal and basipetal directions. Treatment with JA-Me (0.5%, *w*/*w* in lanolin) at the middle of the last internode in decapitated growing plants also induced the SAZ below the treatment (Supplementary Figure S1).

**Figure 1.** Secondary abscission zone (SAZ) induced by the application of methyl jasmonate (JA-Me) in internode segments of *Bryophyllum calycinum*. The treatment was made in the middle of internode explants. Photograph was taken 8 days after treatment. Red and blue arrows indicate the SAZ and JA-Me treatment place, respectively. Stem pieces (ca. 3–4 mm in length) above and below the SAZ were subjected to comprehensive plant hormone analyses.

#### *2.2. Changes in the Levels of Endogenous Plant Hormones in Relation to the Formation of the Secondary Abscission Zone Induced by JA-Me*

Comprehensive analyses of the endogenous plant hormones and their related compounds concerning the induction of the SAZ were performed in the internode segments of *B. calycinum*. At the appropriate time or 6 or 7 days after the treatment, small pieces of the internode segments adjacent to the SAZ were harvested for plant hormone analyses, as illustrated in Figure 1. Similar internode pieces above and below the SAZ in decapitated growing plants of *B. calycinum* were also subjected to the plant hormone analyses (Supplementary Figure S1).

#### 2.2.1. Effect of JA-Me on Auxin-Related Compounds

As shown in Figure 2, the following auxins and their related compounds were successfully identified in internode segments of *B. calycinum*: indole-3-acetic acid (IAA), indole-3 acetamide (IAM), indole-3-acetonitrile (IAN), 2-oxindole-3-acetic acid (OxIAA), indole-3 carboxylic acid (ICA), indole-3-acetyl-aspartic acid (IAAsp), indole-3-acetyl-glutamic acid (IAGlu), and indole-3-propionic acid (IPA).

In the internode segment treated with JA-Me, endogenous levels of IAA, IAGlu, IAAsp, OxIAA, and IAM, in the above (senescent side, yellow color) and below pieces (non-senescent side, green color) adjacent to the secondary abscission in internode segments were similar. However, the contents of IAN and ICA were lower in the senescent than in the non-senescent side (Figure 2). These results suggest that the SAZ formation induced by JA-Me is closely related to the modification of IAA biosynthetic pathways via IAM, IAN, and ICA from tryptophan.

It should be mentioned that the endogenous level of IPA is much higher in the senescent than in the non-senescent side, suggesting that IAA metabolism to IPA is possible to be related to the SAZ induced by JA-Me (Figure 2).

Similar results of the effect of JA-Me on the endogenous levels of auxin-related compounds in the internode segments were obtained in the last internode of decapitated growing plants of *B. calycinum* (Supplementary Figure S2). These results suggest that

the application of JA-Me substantially affects the IAA metabolism in the internode of *B. calycinum* and then might induce secondary abscission.

**Figure 2.** Endogenous levels of auxin-related compounds in the senescent and non-senescent sides of the SAZ induced by JA-Me in the internode explants of *Bryophylum calycinum*. IAA: indole-3 acetic acid; IAM: in-dole-3-acetamide; IAN: indole-3-acetonitrile; IPA: indole-3-propionic acid; ICA: indole-3-carboxylic acid; OxIAA: 2-oxindole-3-acetic acid; IAAsp: indole-3-acetylaspartic acid; IAGlu: indole-3-acetylglutamic acid. Values are the mean with standard error (n = 6). Different letters on the column (a, b) indicated statistically significant at *p* < 0.05 after ANOVA.

2.2.2. Effect of JA-Me on Jasmonate-Relating Compounds, Abscisic Acid, Salicylic Acid and Benzoic Acid

The contents of 12-oxo-phytodienoic acid (OPDA) and *N*-jasmonyl-leucine (JA-Leu) were substantially lower in the stem above the senescent than in the non-senescent side, but the content of jasmonic acid (JA) was similar in the stem pieces below and above the SAZ (senescent and non-senescent) in the internode explants (Figure 3).

An almost similar tendency was observed in the decapitated growing plants of *B. calycinum* (Supplementary Figure S3), suggesting that the application of JA-Me substantially increases endogenous levels of JA.

The endogenous levels of abscisic acid (ABA), salicylic acid (SA), and benzoic acid (BA) occurred in similar amounts in the stem pieces below and above the SAZs induced by JA-Me both in stem explants and in the internode of decapitated growing plants of *B. calycinum* (Figure 4 and Supplementary Figure S4).

**Figure 3.** Endogenous levels of jasmonate-related compounds in the senescent and non-senescent sides of the SAZ induced by JA-Me in the internode explants of *Bryophyllum calycinum*. JA: jasmonic acid; OPDA: 12-oxo-phytodienoic acid; JA-Leu: *N*-jasmonyl-leucine. Values are the mean with standard error (n = 6). Different letters on the column (a, b) indicated statistically significant at *p* < 0.05 after ANOVA.

**Figure 4.** Endogenous levels of abscisic acid (ABA), salicylic acid (SA), and benzoic acid (BA) in the senescent and non-senescent sides of the SAZ induced by JA-Me in the internode explants of *Bryophylum calycinum*. Values are the mean with standard error (n = 6). Different letters on the column (a) indicated statistically significant at *p* < 0.05 after ANOVA.

2.2.3. Effect of JA-Me on Cytokinins

The contents of identified cytokinins such *trans*-zeatin (t-Z), *cis*-zeatin (c-Z), *trans*-zeatin riboside (t-ZR), and *cis*-zeatin riboside (c-ZR) were similar in the senescent and non-senescent sides of SAZ induced by JA-Me both in internode explants and in the internode of decapitated growing plants of *B. calycinum* (Figure 5 and Supplementary Figure S5).

**Figure 5.** Endogenous levels of cytokinins in the senescent and non-senescent sides of the SAZ induced by JA-Me in the internode explants of *Bryophylum calycinum*. t-Z: *trans*-Zeatin; c-Z: *cis*-Zeatin; t-ZR: *trans*-Zeatin riboside; c-ZR: *cis*-Zeatin riboside. Values are the mean with standard error (n = 6). Different letters on the column (a) indicated statistically significant at *p* < 0.05 after ANOVA.

#### 2.2.4. Effect of JA-Me on Gibberellins

Thirteen gibberellins (GAs), gibberellin A1 (GA1), GA3, GA4, GA5, GA6, GA7, GA8, GA9, GA15, GA19, GA20, GA44, and GA53, were also successfully identified in the internode segments of *B. calycinum*. Similar levels of these GAs were found in both the senescent and non-senescent sides of SAZ induced by JA-Me, except that GA8 was lower above the SAZ (senescent side; Table 1). A similar tendency was observed in the decapitated growing plants of *B. calycinum* treated with JA-Me (Supplementary Table S1).

**Table 1.** Endogenous levels of gibberellins in the senescent and non-senescent sides of the SAZ, induced by JA-Me in the internode explants of *Bryophylum calycinum*. Values are the mean with standard error (n = 6). Different letters (a, b) on the column indicated statistically significant at *p* < 0.05 after ANOVA.


#### **3. Discussion**

As mentioned in the Introduction (Section 1), many plant species develop the secondary abscission zone that extends between organs and the main body of the plants to shed. Plant hormones may play an essential role in the transdifferentiation in mature cortical cells to induce the SAZ. JA-Me and JA (designated as jasmonates, JAs) show the powerful effect of inducing the SAZ in stems and developing an abscission zone that has already been initiated in plant tissues in *B. calycinum* [31,33]. JAs were applied in lanolin

paste, where lanolin alone did not affect morphological changes in the internode segments. This situation was demonstrated in previous works [27,31,35]. In the stem of *B. calycinum*, IAA applied to a decapitated shoot or internode explants prevented the formation of the SAZ induced by JA-Me [31,33]. Contrarily, IAA application has also been demonstrated to substantially induce the formation of the SAZ not only in internode explants, petiole segments, and petiole after excision of the leaf blade in intact plants but also decapitated stems in intact plants of *B. calycinum* [31,33]. A decrease in auxin levels might be considered as providing the first signal for abscission, as suggested in Arabidopsis [36] and tomatoes [32,37]. JAs, together with the disruption of endogenous auxin status by the decapitation or excision, may trigger the formation of the SAZ. The results confirm our previous observations [27,31,33], indicating that JA-Me is translocated in stem explants of *B. calycinum* in both ways, acropetally and basipetally, from the place of treatment. The SAZ development place is considered the final result of the stem's secondary abscission formation and senescence. Thus, it could be asserted that fresh, green tissues of the stem below the SAZ are not affected by JA-Me and can also be treated as a control.

What kinds of hormonal control factors are responsible for the formation of the SAZ induced by JA-Me? The SAZ formation by JA-Me has already been reported to be closely related to auxins [27,31]. Therefore, it is worthwhile to study the dynamics of plant hormones, especially auxins in the senescent and the non-senescent sides of the SAZ induced by JA-Me. Notably, the IAA gradient was not observed in the explants between the induced SAZ on both sides. The same situation occurred in the internode of the decapitated growing plant (Figure 2 and Supplementary Figure S2).

It has been reported that JA-Me is converted into JA and jasmonyl-isoleucine (JA-Ile), activating the jasmonates signaling pathway and emission of volatile organic compounds in *Achyranthes bidentate* [38]. The application of JA-Me resulted in the differences in endogenous levels of auxin-related compounds such IAN, ICA, and IPA in the senescent and non-senescent sides of the SAZ (Figure 2 and Supplementary Figure S2). Endogenous levels of OxIAA, which is one of IAA metabolites, were also different. These results suggest that the SAZ induced by JA-Me is closely related to the disruption of IAA metabolism in the stem adjacent to the SAZ.

IPA and IBA, other auxins that share similar structural scaffolds, are strongly conjugated and hydrolyzed with enzymes with similar or even higher activities than with IAA or IAA conjugates [39]. The occurrence of IBA has been reported in various plants, including *B. calycinum* [27]. In the present study, we report for the first time the occurrence of IPA in *B. calycinum*. The natural occurrence of IPA is scant, and until now, little is known about the physiological activity of IPA compared to IAA [39]. The content of IPA was relatively high in the stem of *B. calycinum*, and evidently, the content of IPA further increased on the stem side of JA-Me treatment, suggesting that IPA is responsible for the SAZ formation in *B. calycinum*.

Jasmonates (JAs) might function as a core signal in the plant hormone signaling network, a signal of JAs interacting with other hormone signaling to regulate plant growth, and abiotic and biotic stress tolerance [40–43]. Evidence for a close functional relationship between JAs signaling and auxin homeostasis has been well documented [44,45]. Du et al. [46] showed that biosynthesis and signaling of JA and IAA are differentially regulated by different abiotic stresses in rice, suggesting that the balance between JA and IAA homeostasis and their signaling are critical for plant development and stress responses. The application of JA-Me substantially induces an increase in the endogenous levels of JA in the stem of explant and internode of the decapitated growing plant of *B. calycinum*, as well as the disruption of auxin metabolism, but negligibly affected dynamics of ABA, cytokinins, and GAs (Figures 3–5, Table 1, Supplementary Figures S3–S5 and Table S1). Thus, cross-talk between JAs and auxin might be essential for the induction of the SAZ formation.

Based on the results of comprehensive analyses of endogenous plant hormones, Marasek-Ciolakowska et al. [27] strongly suggested that GAs and cytokinins did not contribute to the formation of the IAA-induced SAZ in *B. calycinum*. In this experiment, JA-Me

also little affected the endogenous level of GAs, ABA, and cytokinins in stems above and below the SAZ (Figures 3 and 4, Table 1, Supplementary Figures S3 and S4, and Table S1). Thus, these plant hormones seem not to contribute to the formation of the JA-Me-induced SAZ in *B. calycinum* as the IAA-induced one [27].

Until now, four tryptophan (Trp)-dependent pathways of IAA biosynthesis, namely the indole-3-acetamide (IAM) pathway, the indole-3-pyruvic acid (IPyr) pathway, the tryptamine pathway, and the indole-3-acetaldoxime (IAOx) pathway, were identified in plants [39,47–50], although biosynthesis pathway(s) of IAA in plants of the Crassulaceae family (succulents) is unknown. The Trp-independent IAA biosynthesis from indole was also documented in some plants [50]. In *Arabidopsis thaliana*, indole-3-carbaldehyde and indole-3-carboxylic acid (ICA) are synthesized from Trp via intermediates such IAOx and IAN, although ICA can also be attributed to the degradation of IAA [51]. Whether ICA can be converted to IPA and vice versa, as indole-3-butyric acid (IBA) and IAA interconversions, has not been shown as yet [39]. ICA has been identified in *Pinus sylvestris* needles, in the leaves of *Ginkgo biloba*, and in the stem of *B. calycinum* [27,31,52].

The occurrence of IAM and IAN in the stem of *B. calycinum* may suggest that biosynthesis of IAA in the plant is going through the IAM and IAOx pathways since IAM and IAN are downstream intermediate metabolites of IAOx [50,53]. The IAOx-dependent IAA biosynthesis pathway was indicated in some plants, but it is not a common pathway [50]. Other pathways of IAA biosynthesis are also possible in *B. calycinum*. Intensive studies on JAs-dependent changes in metabolism or biosynthesis of IAA and the physiological function of ICA, related to the secondary abscission formation, will be needed in the future.

#### **4. Materials and Methods**

#### *4.1. Plant Materials and Hormone Treatment*

Three- to six-month-old plants of *Bryophyllum calycinum* Salisb. (Crassulaceae), propagated from epiphyllous buds arising in the marginal notches of the leaves, were used in the experiments. Stem segments and decapitated stems of growing *B. calycinum* plants were used in methyl jasmonate (JA-Me) treatment.

Internode segments at the length of ca. 4–5 cm with two nodes (leaves removed) were excised, from mainly the second or third internodes from the top of growing plants with active elongation. The segments were treated with JA-Me at 0.5% (*w*/*w*) in lanolin paste in the middle of the internode and kept vertically in a 50 mL glass chamber with moistened papers at the bottom of these explants under natural light conditions in a greenhouse, as shown in Figure 1. In June, August, and September, experiments were repeated three times with 15 to 20 explants.

A similar experiment with decapitated growing plants was carried out. After decapitation of the apical part of the growing plant shoot, JA-Me (0.5%, *w*/*w* in lanolin) was applied in the middle of the last internode, as shown in Supplementary Figure S1. The experiment was repeated twice from August to October with 20 explants.

#### *4.2. Analyses of Plant Hormones in Relation to the Formation of the Secondary Abscission Zone Induced by Methyl jasmonate*

Analyses of plant hormones were performed according to the methods reported previously [27,35,54–56]. At 6 to 7 days after treatment with JA-Me, the below (nonsenescent, green) and above (senescent, yellow) parts of ca. 3–4 mm internode pieces adjacent to the SAZ formed by JA-Me application in the stem of *B. calycinum* were excised, respectively. Excised samples were immediately frozen in liquid N2 and then lyophilized. Lyophilized materials in each piece of internode were combined, and an aliquot of a small amount (ca. 10 mg DW) was used for comprehensive plant hormone analyses. Lyophilized materials with appropriate amounts of a mixture of each stable isotope-labeled plant hormone as an internal standard were extracted with an organic solvent consisting of methanol/water/formic acid = 15: 4: 1 (*v*/*v*/*v*) three times. Respective extracts were combined and then evaporated under N2. The extract obtained was re-suspended in 3% methanol in 1 M formic acid and then cleaned up on hybrid SPE cartridges (BondElut Plexa PCX, Agilent, Santa Clara, CA, USA). Qualitative and quantitative analyses of plant hormones and other related compounds were performed on a HPLC-MS/MS system with UHPLC apparatus (Agilent Infinity 1260, Agilent, Waldbronn, Germany) coupled to a triple quadruple mass spectrometer ESI-MS/MS (6410 Triple Quad LC/MS, Agilent, Santa Clara, CA., USA). Plant hormones were separated on an Ascentis Express RP-Amide analytical column (particle size: 2.7 μm; 2.1 mm × 150 mm; Supelco, Bellefonte, PA., USA) at 60 ◦C, at a linear gradient of water vs. acetonitrile both with 0.01% of formic acid. As internal standards, [15N4] dihydrozeatin, [15N4] kinetin, [2H5] *trans*-zeatin riboside (t-ZR), [2H5] indole-3-acetic acid (IAA), [2H4] indole-3-acetonitrile, [2H4] salicylic acid (SA), [2H2] gibberellin A1 (GA1), [2H2] gibberellin A4 (GA4), [2H2] gibberellin A5 (GA5), [ 2H2] gibberellin A6 (GA6), [2H6] *cis*, *trans*-abscisic acid (ABA), [2H5] benzoic acid (BA), [ 2H5] jasmonic acid (JA), and [2H5] dinor-12-oxo-phytodienoic acid (dinor-OPDA) were used. All standards, except for [2H5] JA supplied by CND Isotopes (Quebeck, Canada) and [2H5] dinor OPDA supplied by Cayman Chem. Comp. (Ann Arbor, USA), were from OlChemim (Olomouc, Czech Republic) at the highest available purity. Multiple reaction monitoring (MRM) transitions were used to identify and quantify all compounds of interest. Quantitation was based on calibration curves obtained with each pure standard compound taking account of the recovery rates of an internal standard used. Further technical details are given by the references cited above.

#### *4.3. Statistical Analysis*

The analysis of variance (ANOVA) was conducted using STATISTICA software (StatSoft, Kraków, Poland). To compare the means, Duncan's multiple range test was used. Values of *p* < 0.05 were considered to be statistically significant. Values are expressed as the mean with standard error. Different letters in the columns in the figures and tables indicate statistical differences.

#### **5. Conclusions**

A comprehensive study of the dynamics of plant hormones in the stem pieces above and below the SAZ induced by the application of JA-Me in *B. calycinum* revealed that the application of JA-Me substantially affected auxin metabolism and the endogenous status of JAs. However, it negligibly affected the endogenous IAA levels. These suggest that the mode of JA-Me action to induce the SAZ is different from that of IAA, whereas IAA also induces the SAZ. JA-Me functions as a trigger modifying metabolism of IAA and JAs to induce the formation of the SAZ in the stem of *B. calycinum*.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/plants11030360/s1, Figure S1: The secondary abscission zone induced by the application of methyl jasmonate (JA-Me) in the last internode of decapitated growing plants of *Bryophyllum calycinum*, Figure S2: Endogenous levels of auxin-related compounds in the stem pieces above and below the secondary abscission zone induced by JA-Me in the last internode of decapitated growing plants of *Bryophyllum calycinum*, Figure S3: Endogenous levels of jasmonate-related compounds in the stem pieces above and below the secondary abscission zone induced by JA-Me in the last internode of decapitated growing plants of *Bryophyllum calycinum*. Figure S4: Endogenous levels of abscisic acid, salicylic acid, and benzoic acid in the stem pieces above and below the secondary abscission zone induced by JA-Me in the last internode of decapitated growing plants of *Bryophyllum calycinum*. Figure S5: Endogenous levels of cytokinins in the stem pieces above and below the secondary abscission zone induced by JA-Me in the last internode of decapitated growing plants of *Bryophyllum calycinum*. Table S1: Endogenous levels of gibberellins in the stem pieces above and below the secondary abscission zone induced by JA-Me in the last internode of decapitated growing plants of *Bryophyllum calycinum*.

**Author Contributions:** Conceptualization, A.M.-C., K.M. and M.S.; Methodology, M.D., A.M.-C. and M.S.; Software, J.G.-K.; Investigation, M.D., A.M.-C. and U.K.; Writing—Original Draft Preparation, A.M.-C., J.G.-K., K.M. and M.S.; Writing—Review and Editing, M.D., A.M.-C., K.M., J.U. and M.S.; Visualization, A.M.-C., K.M. and M.S.; Supervision, M.S.; Funding Acquisition A.M.-C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was partly supported by the Polish Ministry of Science and Higher Education through statutory funds of the Research Institute of Horticulture, Skierniewice, Poland (Grant ZBS/7/2021).

**Data Availability Statement:** The data sets generated for this study are available in this article and Supplementary Material.

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

#### **References**


### *Article* **Cold Acclimation in** *Brachypodium* **Is Accompanied by Changes in Above-Ground Bacterial and Fungal Communities**

**Collin L. Juurakko 1,\*, George C. diCenzo <sup>1</sup> and Virginia K. Walker 1,2**


**Abstract:** Shifts in microbiota undoubtedly support host plants faced with abiotic stress, including low temperatures. Cold-resistant perennials prepare for freeze stress during a period of cold acclimation that can be mimicked by transfer from growing conditions to a reduced photoperiod and a temperature of 4 ◦C for 2–6 days. After cold acclimation, the model cereal, *Brachypodium distachyon*, was characterized using metagenomics supplemented with amplicon sequencing (16S ribosomal RNA gene fragments and an internal transcribed spacer region). The bacterial and fungal rhizosphere remained largely unchanged from that of non-acclimated plants. However, leaf samples representing bacterial and fungal communities of the endo- and phyllospheres significantly changed. For example, a plant-beneficial bacterium, *Streptomyces* sp. M2, increased more than 200-fold in relative abundance in cold-acclimated leaves, and this increase correlated with a striking decrease in the abundance of *Pseudomonas syringae* (from 8% to zero). This change is of consequence to the host, since *P. syringae* is a ubiquitous ice-nucleating phytopathogen responsible for devastating frost events in crops. We posit that a responsive above-ground bacterial and fungal community interacts with *Brachypodium*'s low temperature and anti-pathogen signalling networks to help ensure survival in subsequent freeze events, underscoring the importance of inter-kingdom partnerships in the response to cold stress.

**Keywords:** *Brachypodium distachyon*; cold acclimation; microbiome; amplicon and shotgun sequencing; metagenomics; *Pseudomonas*; *Streptomyces*

### **1. Introduction**

As sessile organisms, plants are at the mercy of an array of abiotic stresses, and, as winter approaches in mid- to high-latitudes and altitudes, one such stress is low temperature. Plants employ various strategies that allow them to recognise and cope with the cold [1]. As autumn progresses, perennials undergo a period of cold acclimation, which in a few days of low temperature exposure allows them to physiologically prepare for freezing conditions. Such preparations include changed levels of hundreds of proteins, the accumulation of fatty acids, lipid remodelling for plasma membrane protection, increased production of cryoprotective metabolites, such as soluble sugars and amino acids, as well as chaperones and reactive oxygen scavengers [2]. This acclimation process also appears to coincide with changes in host-associated microbial communities. Such a turnover in microbiota could assist plants in preparing for sub-zero temperature conditions and their vulnerability to psychrophilic pathogens. Indeed, winter seasonality in the plant microbiome has been previously reported [3–5]. Although the impact of cold acclimation on the microbiomes of perennial grass has not been hitherto explored, the identification of their bacterial and fungal communities offers the promise of understanding how the battle against coming winter conditions can be won by partnerships.

The perennial grass and model cereal, *Brachypodium distachyon* (hereinafter, *Brachypodium*), is capable of cold acclimation, reaching peak freezing tolerance after two days

**Citation:** Juurakko, C.L.; diCenzo, G.C.; Walker, V.K. Cold Acclimation in *Brachypodium* Is Accompanied by Changes in Above-Ground Bacterial and Fungal Communities. *Plants* **2021**, *10*, 2824. https://doi.org/10.3390/plants10122824

Academic Editor: Ewa Muszy ´nska

Received: 30 November 2021 Accepted: 16 December 2021 Published: 20 December 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

at 4 ◦C, and is associated with changes in the abundance of multiple plasma membrane proteins at 2–6 days [6]. In turn, these proteins are involved in complex crosstalk networks that prime the *Brachypodium* defensive response to a variety of abiotic and pathogenic stresses. Studies of cold acclimation have, for the most part, ignored the host-associated microbiota [1,7,8]. Nevertheless, the plant microbiome is emerging as an important factor in stress responses, including symbiont-mediated tolerance [9,10].

The general beneficial effects of microbes on plant fitness under a variety of stressful conditions have recently come to be known as the "Defence Biome" [5,10–17]. Symbiontmediated fitness benefits may be a collective result of microbial exudates and function, for example, by facilitating early stress sensing and more efficient nutrient uptake and transfer, as well as by the induction of plant stress genes [9,10]. Specifically, symbiontmediated cold tolerance has been directly demonstrated with some plant species and plant growth promoting bacteria (PGPBs) [9]. For example, *Burkholderia phytofirmans*inoculated grape vines expressed cold stress-responsive genes earlier than non-inoculated vines [18] and *Streptomyces neyagawaensis* J6-inoculated turfgrass showed enhanced cold tolerance over non-inoculated plants [19]. Microbes thus have a demonstrated role in plant protection. They excrete a variety of products to benefit host plants, including anti-pathogenic microbial compounds and osmolytes, including proline and trehalose, as well as scavengers of reactive oxygen species, such as superoxide dismutase, catalase, and peroxidases [9,10,20]. Taken together, plant-associated microbial communities undoubtedly help plants survive cold stress.

The identification of host-associated microbiota that enhance freezing tolerance may lead the way to the development of synthetic cocktails of species that could eventually be used to inoculate crops or seeds to enhance cold tolerance [21]. Here, shotgun sequencing and metagenomic analysis of the phyllosphere/endosphere and rhizosphere in cold-acclimated *Brachypodium* is an important first step towards this goal. Our experimental inoculation of a commercial growing mix with old pasture soil allowed for the exposure and subsequent identification of bacterial and fungal taxa that thrived after transfer of the growing plants to low temperatures and thus are prospective native partners in the cold acclimation process. In addition, we contribute to the general appreciation of the robustness of the plant abiotic stress response, which employs communities of diverse organisms for survival.

#### **2. Materials and Methods**

#### *2.1. Soil Inoculation and Preparation*

Commercial potting soil (Sun Gro Horticulture, Agawam, MA, USA) was autoclaved twice and sealed in a double layer of plastic autoclave bags before being inoculated with bulk field soil (5% *w*/*v*). Bulk field soil was sampled using a sterilized trowel from the active layer (3–7 cm depth) in autumn (29 October 2020) after 96 h of day and night temperatures of ~5 ◦C and ~0 ◦C, respectively. The sampled fallow field had been left unfertilized and unplowed for 26 years and without domestic grazing animals for 15 years (Figure S1). It was characterized by grasses, including orchard grass, brome, and timothy (*Dactylis*, *Bromus*, and *Phleum* species, respectively) on clay soils and was located north of Sydenham, Ontario, Canada (44◦24 26" N, 76◦36 1" W). Soils were thoroughly mixed for 15 min using a cement mixer that had been rinsed with 70% ethanol, with the inoculated soil then stored in a lidded container that had also been rinsed with 70% ethanol. The inoculated soil mixture was kept at room temperature until use.

#### *2.2. Plant Material and Growth Conditions*

Surface-sterilized *Brachypodium* seeds of an inbred line (ecotype: *Bd*21) (RIKEN, Wako, ¯ Japan) were sown in the inoculated potting soil and grown in a temperature-controlled chamber (Conviron GEN2000, Queen's University Phytotron, Kingston, ON, Canada) on a 20 h light (~100 μmol m−<sup>2</sup> s−1; 22 ◦C) and 4 h dark (22 ◦C) light cycle. *Brachypodium* that had been grown under standard conditions for three weeks (Figure S2) were then

cold acclimated by transferring the plants to a low temperature chamber (Coldmatic Refrigeration, Etobicoke, ON, Canada) (4 ◦C, 12 h light as indicated above; 12 h dark) for 6 days [6]. Plants maintained at standard conditions until time of use were considered the non-acclimated controls.

#### *2.3. Microbiome Extraction and Preparation*

Microbiome extractions were performed under sterile conditions. Above-ground extractions were from tissue excised from the tips of primary leaves. Phyllosphere microbes are found on the leaf surface and endosphere microbiota include communities that enter the plant through the leaves, as well as those that circulate within the xylem. Rather than separate these, we reasoned that both phyllosphere and endosphere communities would be driven by the changing environmental conditions, in addition to plant interactions. Accordingly, these leaf microbiota were extracted together using a DNeasy Plant Pro Kits (Qiagen, Hilden, Germany), following the manufacturer's recommended directions, using 10 mg of leaf tissue per plant (10 plants per replicate for a total of 100 mg of tissue) and three replicates.

Extractions of the below-ground, tightly bound root soil of the rhizosphere (Figure S3) were performed as previously described [22] using a DNeasy PowerSoil Pro Kit (Qiagen, Hilden, Germany), following the manufacturer's recommendations. Adhering root soil (25 mg per plant) was released from the roots following careful removal of the plants from the pots and gentle shaking. Extra care was taken to remove any root tissue, or non-soil material from samples, such as wood or perlite. Three replicates were performed, each using 10 individual plants. DNA purity and concentration was quantified using a Synergy H1 microplate reader with a Take3 Micro-Volume Plate (both BioTek Instruments Inc., Winooski, VT, USA).

#### *2.4. Shotgun Metagenomics Library Preparation and Sequencing*

Libraries were prepared using an Illumina DNA Prep (M) Tagmentation library preparation kit (Illumina Inc., San Diego, CA, USA), following the manufacturer's user guide. Initial DNA concentration was evaluated using the Qubit dsDNA HS Assay Kit (Life Technologies, Carlsbad, CA, USA). Eukaryotic DNA was depleted in leaf tissue samples using an NEBNext Microbiome DNA Enrichment Kit (New England Biolabs, Ipswich, MA, USA), following the manufacturer's user guide to decrease the probability of recovery of host genomic, chloroplast, and mitochondrial DNA sequences [23]. DNA (500 ng) was used for depletion of the eukaryotic DNA, as recommended by Molecular Research LP (MR DNA; Shallowater, TX, USA). The enriched microbial DNA was quantified using the Qubit dsDNA HS Assay Kit (Life Technologies, Carlsbad, CA, USA) (Table S1). Subsequently, 50 ng of DNA was used to prepare the libraries. The samples underwent simultaneous fragmentation and addition of adapter sequences, which were utilized during a limited-cycle polymerase chain reaction in which unique indices were added to the sample. Following library preparation, library concentration and mean library size were determined using the Qubit dsDNA HS Assay Kit (Life Technologies, Carlsbad, CA, USA) and the Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA), respectively. Libraries were pooled in equimolar ratios (0.6 nM), and sequencing was performed on a NovaSeq 6000 platform (Illumina Inc., San Diego, CA, USA) to a depth of 10 million 2 × 150 bp reads.

#### *2.5. Preprocessing and Quality Control*

Analysis of sequencing data was performed following the Sunbeam pipeline (v2.1.0) [24] with 26 available cores (15.425 Gb of memory each) on Ubuntu (v18.04.05). Raw fastq files of paired-end reads were quality controlled to remove adapter sequences using Cutadapt (v3.4.0) [25] and Trimmomatic (v0.3.9) [26], following which read quality was assessed using FastQC (v0.11.9) [27]. Low-complexity sequences were masked using Komplexity (v0.3.6) [24] and contaminating plant host reads were removed by Sunbeam following mapping of reads to the *Brachypodium* genome (RefSeq assembly accession GCF\_000005505.3)

using BWA (v0.7.17) [28]. Following initial host read decontamination, individual reads were interrogated using the National Center for Biotechnology Information (NCBI) BLAST *(blastn;* available at https://blast.ncbi.nlm.nih.gov/Blast.cgi; accessed on 18 August 2021), revealing numerous hits to mitochondrial genomic sequences. Subsequently, several mitochondrial genomic sequences (detailed below) were subsequently downloaded and added to the host genome path for removal of contaminating mitochondrial sequences. This process was repeated until a subset of individual reads did not return any mitochondrial genomes with high coverage.

Most mitochondrial genomes used to filter contaminating sequences were retrieved from NCBI from the following species with GenBank IDs: *Saccharum officinarum* cv. Khon Kaen 3 (NC\_031164.1), *Sorghum bicolor* (NC\_008360.1), *Triticum aestivum* cv. Chinese Yumai (NC\_036024.1), *Oryza sativa* (NC\_011033.1), *Zea mays* (NC\_007982.1), *Lolium perenne* (JX999996.1), *Oryza coarctata* (MG429050.1), *Sporobolus alterniflorus* (MT471321.1), *Aegilops speltoides* (AP013107.1), *Stipa capillata* (MZ161090.1, MZ161091.1, MZ161093.1, MZ161092.1), *Bambusa oldhamii* (EU365401.1), and a *Brachypodium* sequence (AC276583.1), suggesting a partial *Brachypodium* mitochondrial draft genome. In addition, the *Hordeum vulgare* mitochondria genome sequence was downloaded from Ensembl Plants (ID: IBSC\_v2, chromosome Mt). Pre-processing and quality control data is summarized in Table S2.

#### *2.6. Taxonomic Classification*

Taxonomic assignment was performed on the quality-controlled and host-decontaminated reads using a Kraken2 (v2.1.2) [29] database containing RefSeq libraries [30] of archaea (628 sequences), bacteria (58,811 sequences), fungi (1579 sequences), and protozoa (11,151 sequences) for a total of 72,217 sequences and ~110 billion bp (as of 24 June 2021). A Bayesian re-estimation of abundance with the Kraken (Bracken) (v2.6) [31] database was subsequently built with the Kraken2 database using the default 35 k-mer length and 150 bp read lengths. Kraken2 was run as an integrated module of Sunbeam using the development branch. Bracken was run on the Kraken2 output files, and the Bracken outputs were combined using the combine\_bracken\_outputs.py function for downstream analysis. Barplots were produced using the thresholds indicated in the legends to group together low abundant taxa for visual presentation. For diversity analysis, the kraken-biom tool (v1.0.1) (https://github.com/smdabdoub/kraken-biom; accessed on 27 September 2021) was used to convert Bracken outputs at the species level into .biom files for use with the Phyloseq (v1.36.0) [32] and Vegan (v2.5.7) [33] R packages.

#### *2.7. Core and Functional Microbiome*

To further characterize the microbiomes, PAST (Paleontological Statistics, v4.08, available at https://www.nhm.uio.no/english/research/infrastructure/past/; accessed on 15 November 2021) [34] was used for similarity percentage (SIMPER) analyses using the Bray–Curtis similarity matrix to compare leaf and rhizosphere-associated microbiota and to facilitate the identification of a core microbiome [35–37]. Core microbiomes were calculated based on species and ASVs present in 100% of the tissue-specific samples with >5% relative abundance.

Paired-end quality-controlled and decontaminated reads outputted by Sunbeam were concatenated using the command "cat sample\_R1.fq sample\_R2.fq > merged\_sample.fq" and inputted into HUMAnN (v3.0.0) [38] running MetaPhlan (v3.0) [38], Bowtie2 (v2.4.4) [39], DIAMOND (v2.0.11) [40], and SAMtools (v1.13) [41,42]. Sequences were processed using the default UniRef90 database and the following parameters for MetaPhlAn: –stat\_q 0, –bt2\_ps very-sensitive-local; the following parameters for HUMAaN 3: –nucleotide-subjectcoverage-threshold 5.0, –translated-subject-coverage-threshold 5.0; and the following parameters for and Bowtie 2: -D 20 -R 3 -N 1 -L 20 -i S,1,0.50 –local.

Gene families were regrouped and renamed to the uniref90\_Pfam database using the humann\_regroup\_table and humann\_rename\_table commands. Special features, including ungrouped genes and unintegrated pathways, were retained by skipping normalization in favour of downstream normalization using MaAsLin2 (v1.6.0) [43]. The final

renamed gene family and unnormalized pathway abundance tables were joined using the humann\_join\_table command and split into the stratified and unstratified tables using the humann\_split\_table command, the latter of which was used for differential abundance testing. Standard HUMAnN3 MetaCyc assigned metabolic pathways were used for analysis and were assigned classes based on the respective associated MetaCyc pathway superclasses. All scripts can be found in Supplementary File S1.

#### *2.8. Amplicon Sequencing*

Aliquots of the DNA extractions used for shotgun sequencing were sent to MR DNA for amplification and barcoded amplicon sequencing of the 16S rRNA V4 region using primers 515F (5 -GTGYCAGCMGCCGCGGTAA-3 ) [44] and 806R (5 -GGACTACNVGGG TWTCTAAT-3 ) [45], and of the ITS region using primers ITS1F (5 -CTTGGTCATTTAGAGG AAGTAA-3 ) and ITS2R (5 -GCTGCGTTCTTCATCGATGC-3 ) [46]. Peptide nucleic acid clamps pP01 (5 -GGCTCAACCCTGGACAG-3 ), as previously described [47], were used to reduce amplification of *Brachypodium-*contaminating sequences during the amplification of the 16S rRNA V4 regions. Blank kit controls for both Plant Pro and PowerSoil Pro kits were performed in triplicate and subjected to the same amplification and sequencing as the corresponding samples. Sequencing was performed on a MiSeq platform (Illumina Inc., San Diego, CA, USA) for ITS and NovaSeq 6000 platform (Illumina Inc., San Diego, CA, USA) for 16S.

#### *2.9. Amplicon Sequence Processing*

Sequences were processed using QIIME2 (v2021.4) [48]. Raw .fastq files were demultiplexed and non-biological sequences were removed, including primers, adapters, spacers, and linkers, using FASTqProcessor (v20.11.19). Sequences were trimmed and denoised to remove any chimeras and singletons using DADA2 (v1.18) [49] before being grouped into amplicon single variants (ASVs). ASVs were used for taxonomic classification with SILVA (v138) for 16S rRNA sequences and UNITE (v8) for ITS sequences [50–55]. In the leaf samples, any taxa classified as eukaryota, chloroplast, mitochondria, archaea, or unclassified were filtered out of the 16S rRNA feature tables. Shannon's diversity index was used as a measure for alpha diversity and Bray–Curtis dissimilarity distance was used as a measure for community dissimilarity. Principal coordinate analysis (PCoA) was performed using Bray–Curtis dissimilarity matrices and plots made in R using ggplot2. Differential abundance between cold-acclimated and non-acclimated samples and between blank kit controls and samples was also assessed at the genus taxonomic levels using ANCOM-BC in R (v1.2.2) [56]. All commands and codes used can be found in Supplementary File S1.

#### *2.10. Statistical Analysis*

All statistical analyses were performed in RStudio (v1.3.1073) running R (v4.1.1) and all scripts used are available in Supplementary File S1. All plots, when necessary, were cleaned up using Inkscape (v0.92.2). Alpha and beta diversity analysis was performed using the Vegan and Phyloseq packages and PCoA plots were performed using ggplot2 (v3.3.5). To find differentially abundant taxa between the two temperature conditions, ANCOM-BC was run on Bracken outputs with default parameters for shotgun data and feature tables for amplicon data. Output coefficients representing the natural log foldchange model were converted to log2 fold changes. ANCOM-BC outputs were parsed to remove any low abundant taxa from differential abundance results.

#### **3. Results**

#### *3.1. Pre-Processing, Shotgun Sequencing, and Kit Controls*

Initial DNA samples representing the cold-acclimated (CA) leaf and rhizosphere were sent for shotgun sequencing without eukaryotic depletion, revealing high host contamination in the leaves (not shown). Subsequent replicate samples undergoing eukaryotic depletion proved successful as the classification of processed reads showed a full order

of magnitude better recovery of microbial sequences. DNA and library concentrations and average size, quality control, host read decontamination, and Kraken2 classification results are summarized in the Supplementary Materials (Figure S4, Tables S1 and S2). Although shotgun DNA library construction was attempted on the blank kit controls, a lack of sufficient DNA resulted in no results for this sequencing method. However, the same control samples were subject to amplicon marker gene sequencing. Following QIIME2 processing, it was determined through diversity analysis and PCoA using Bray–Curtis dissimilarities that the microbial compositions associated with the kits were significantly different than the *Brachypodium* leaf (*p* < 0.001 16S, *p* < 0.05 ITS, pairwise PERMANOVA) and rhizosphere microbiomes (*p* < 0.001 16S, *p* < 0.05 ITS, pairwise PERMANOVA) (Figure S5).

#### *3.2. Compatible Results with Shotgun and Amplicon Sequencing*

The correlation between taxa identified in both the shotgun data and the amplicon data was assessed at the genus level in order to compare the two methods. In the CA rhizosphere, the genera identified by shotgun metagenomic and 16S rRNA amplicon sequences, as well as shotgun metagenomics and ITS amplicon sequencing, were well correlated (R2 = 0.93 and R<sup>2</sup> = 0.88, respectively) (Figure S6). The non-acclimated (NA) rhizosphere shotgun and 16S rRNA, and the shotgun and ITS amplicon results (R<sup>2</sup> = 0.91 and R2 = 0.45, respectively) also correlated, but less well. It is notable that for the leaf microbiome, bacterial taxa in the CA shotgun and 16S rRNA samples, as well as for the NA leaf samples, showed mixed correlations (R<sup>2</sup> = 0.31 and R2 = 0.75, respectively). Insufficient fungal reads in the leaves following Bracken re-estimation resulted in no correlation between the shotgun and ITS reads in the leaves.

#### *3.3. Cold Acclimation and the Rhizosphere Microbiome*

In total, 4646 microbial species were identified in the rhizosphere shotgun data with 45 ± 3% of reads remaining unclassified. The majority of identified reads, 99.70 ± 0.06%, represented bacterial microbes with 0.15 ± 0.03% and 0.13 ± 0.02% representing fungi and archaea, respectively. Alpha diversity, assessed using Shannon's diversity index, across all rhizosphere samples was 4.98 ± 0.21 and was not significantly different between conditions with 5.07 ± 0.29 in the CA and 4.91 ± 0.94 in the NA samples. The rhizosphere was dominated by *Streptomyces* sp. M2, a PGPB, accounting for approximately one-third of the taxa in all samples. Rounding out the top abundant species across the rhizosphere samples were taxa present at 1–10% abundance, which included *Actinocatenispora sera*, *Actinocatenispora thailandica*, *Rhodanobacter denitrificans*, and *Rhodanobacter* sp. *FDA-ARGOS* 1247 (Figure 1A; Table S3). Nearly half of all species in the rhizosphere shotgun data were below a cut-off value (0.2%) for low relative abundance leaving a balance of 53% and 56% of species found in NA and CA samples, respectively.

The amplicon analysis identified 651 distinct ASVs at the genus level. Alpha diversity appeared similar in the NA and CA samples (6.79 ± 0.25 and 6.40 ± 0.16, respectively) and differences were not significant. Both conditions were dominated by the genera *Streptomyces*, *Actinocatenispora*, and *Rhodanobacter (*Figure 1B; Table S3*)*. After CA, low abundant taxa (<1% relative abundance) remained equal at 29%. Again, a similar number of ASVs were considered at low abundance under NA and CA conditions (20% and 15%, respectively). ITS analysis showed 25 distinct ASVs at the genus level (Figure 1C). *Ascomycota* and *Apiotrichum* each represented a third of the ASVs in the rhizosphere irrespective of conditions (Figure 1C; Table S3). Alpha diversity was significantly different (*p* < 0.05, two-tailed *t*-test) at 3.43 ± 0.06 in the CA and 3.05 ± 0.17 in the NA.

**Figure 1.** Average relative abundance of the taxonomies of the non-acclimated and cold-acclimated *Brachypodium distachyon* rhizosphere microbiomes: (**A**) species identified from shotgun sequencing and metagenomics classified using a custom Kraken2 database, (**B**) distinct amplicon sequence variants assigned down to the genus or lowest possible level by QIIME2 using the SILVA database for 16S rRNA sequences amplified using the V4 region of prokaryotes, and (**C**) distinct amplicon sequence variants assigned down to the genus or lowest possible level by QIIME2 using the UNITE database for ITS regions of eukaryotes.

Although there were few changes in the rhizosphere community following 6 days at 4 ◦C, differential abundance testing using ANCOM with bias control and parsed for taxa above the assigned low relative abundance thresholds (Figure 1) identified two modestly differentially abundant species (out of 143; 1.4%) in the shotgun data. *Kribbella qitaiheensis* (log2 fold change: 0.37) and *Kribbella flavida* (log2 fold change: 0.38) increased in relative abundance after CA (Figure 2A). In addition, the relative abundance of three fungal genera (out of 25; 12%) changed following CA, including a decrease in *Penicillium* (log2 fold change: −1.8) and *Phialemonium* (log2 fold change: −1.7) and a more substantial relative increase in *Pseudogymnoascus* (log2 fold change: 8.43) (Figure 2B).

**Figure 2.** Differentially abundant taxa between the non-acclimated and cold-acclimated *Brachypodium distachyon* rhizosphere microbiomes as determined by ANCOM-BC and showing their average relative abundance in both conditions and log2 fold changes with error bars representing standard error: (**A**) species identified by Kraken2 from shotgun sequencing data that are differentially abundant and above an average relative abundance threshold of 0.2%, and (**B**) ITS amplicon sequence variants that are differentially abundant. Only statistically significant changes are shown, as determined by ANCOM-BC.

Although shifts in the rhizosphere community appeared modest, the Bray–Curtis dissimilarity analysis showed that the shotgun rhizosphere communities were significantly different under the two temperature regimes (*p* < 0.01, pairwise PERMANOVA) (Figure 3A). In contrast, there were no differences in Bray–Curtis dissimilarity for the amplicon analysis, either for 16S (Figure 3B) or ITS data (Figure 3C). Taking all the results together, it appears that overall, the CA regime resulted in only a very minor shift in the rhizosphere microbial community. We speculate that a longer period of low temperature with concomitant changes in root exudates would be required for a more dramatic change in the rootassociated microbiota.

**Figure 3.** Principal coordinate analysis comparing non-acclimated and cold-acclimated conditions in each sample type for each sequencing method, for the following samples: (**A**) shotgun sequencing in the rhizosphere, (**B**) 16S rRNA sequencing of the V4 region in the rhizosphere, (**C**) ITS sequencing of the rhizosphere samples, (**D**) shotgun sequencing of the leaf samples, (**E**) 16S rRNA sequencing of the V4 region in the leaf samples, and (**F**) ITS sequencing of the leaf samples. Pairwise PERMANOVAs were conducted between conditions with significance as noted.

#### *3.4. Cold Acclimation and the Leaf Microbiome*

Although shotgun sequencing of the leaf, representing the endosphere and phyllosphere microbiomes, identified 143 microbial species with the most abundant taxa shown (Figure 4A; Table S4), an average of 92 ± 4% of the reads remained unclassified, with a portion of these likely attributable to as yet unsequenced host mitochondrial sequences (Figure S4C). Bacteria accounted for ~100% of the microbiota except in a couple of samples from which a few fungal sequences were recovered. Overall, alpha diversity was significantly lower (*<sup>p</sup>* < 5 × <sup>10</sup>−6, two-tailed *<sup>t</sup>*-test) in leaf samples (3.18 ± 0.36) compared to rhizosphere samples (4.99 ± 0.21).

Leaf alpha diversity did not significantly change after CA treatment (mean Shannon indices at 3.30 ± 0.29 in NA samples and 3.06 ± 0.47 in CA samples). However, the taxa profile changed with the cyanobacteria *Microcystis aeruginosa*, decreasing from ~27% to ~13% relative abundance after CA. *Streptomyces* sp. M2 showed the opposite profile, increasing from ~4% to ~15% average relative abundance after transfer to 4 ◦C. NA leaves were dominated by the plant pathogens *Pseudomonas syringae* and '*Candidatus* Liberibacter africanus', as well as the plant beneficial *Rhodococcus qingshengii,* whose levels substantially decreased in the CA conditions. Lower abundant reads (<1%) made up about a quarter of the taxa, similar to the CA samples.

**Figure 4.** Average relative abundance of the taxonomies of the non-acclimated and cold-acclimated *Brachypodium distachyon* leaf microbiomes representing the endosphere and phyllosphere: (**A**) species identified from shotgun sequencing and metagenomics classified using a custom Kraken2 database, (**B**) distinct amplicon sequence variants assigned down to the genus or lowest possible level by QIIME2 using the SILVA database for 16S rRNA sequences, amplified using the V4 region of prokaryotes, and (**C**) distinct amplicon sequence variants assigned down to the genus or lowest possible level by QIIME2 using the UNITE database for ITS regions of eukaryotes.

Amplicon sequencing of the 16S rRNA from the leaves identified 188 distinct ASVs at the genus level (with the most abundant shown in Figure 4B and Table S4). Again, alpha diversity was not significantly different between conditions (5.04 ± 0.25 and 4.60 ± 0.70 in the CA and NA samples, respectively). Taxa present under both conditions included the genera *Solimonas*, *Rhodanobacter*, and *Streptomyces*. *Pseudomonas* and *Rhodococcus* were abundant (21% and 15% average relative abundance, respectively) in NA conditions, but decreased in relative abundance after transfer of the plants to 4 ◦C with log2 fold changes of −4.18 and −5.41, respectively. The cereal growth-promoting genus *Nocardioides* and an unidentified genus from the same family, *Nocardioidaceae*, both increased in abundance to represent 11% of the taxa in CA plants. ASVs at low relative abundance (<1%) made up a similar 18% and 21% of CA and NA 16S samples, respectively. ITS analysis resulted in 20 distinct ASVs at the genus level (Figure 4C).

After shotgun sequence analysis, 3.5% (5/143) of the taxa were identified as differentially abundant between the NA and CA conditions (Figure 5A). After transfer to 4 ◦C, reads attributed to *P. syringae* (log2 fold change: −8.68) and *R. qingshengii* (log2 fold change: −8.33) decreased so that there was a change in the estimated average relative abundance of *P. syringae* and *R. qingshengii* from 8.2% and 5.0% to 0%, respectively. At the same time there was a corresponding increase in the relative abundance of *Streptomyces* sp. M2 (log2 fold change: 2.81), *A. sera* (log2 fold change: 3.20), and *A. thailandica* (log2 fold change: 3.87). In 16S CA samples, nine other taxa increased, including the genus *Solimonas*, which increased in relative abundance but was below the low abundance threshold. In total, 5.9% (11/188) of the identified sequences above the threshold were found to be differentially abundant. For the ITS analysis, the genus *Phialemonium* represented 5% (1/20) of the ASVs and decreased in relative abundance (log2 fold change: −10.6) (Figure 5C).

**Figure 5.** Differentially abundant taxa between the non-acclimated and cold-acclimated *Brachypodium distachyon* leaf microbiomes representing the endosphere and phyllosphere as determined by ANCOM-BC and showing their average relative abundance in both conditions and log2 fold changes with error bars representing standard error: (**A**) species identified by Kraken2 from shotgun sequencing data that are differentially abundant and above an average relative abundance threshold of 1%, (**B**) distinct 16S rRNA amplicon sequence variants assigned by QIIME2 and the SILVA database to the genus level that are differentially abundant, and (**C**) distinct ITS amplicon sequence variants assigned by QIIME2 and the UNITE database to the genus level that are differentially abundant. Only statistically significant changes are shown as determined by ANCOM-BC.

> Despite the apparent community differences, Bray–Curtis dissimilarity analysis suggested that the microbial communities identified with the shotgun sequencing approach were not significantly different, undoubtedly due to the low number of sequences (Figure 3D), similar to the leaf ITS communities. Supporting that conclusion, 16S rRNA communities were shown to be significantly different between conditions (*p* < 0.01, pairwise PERMANOVA) with the analysis supported by high ASV numbers (Figure 3E).

#### *3.5. Dissimilarity Comparisons and Core Microbiome*

The root and leaf-associated microbiomes were further independently characterized with SIMPER to identify taxa that contributed the most dissimilarity between NA and CA regimes (Table 1). For microbiota isolated from the rhizosphere, the taxa contributing to the top ~25% of dissimilarity were *Streptomyces* sp. M2, *A. sera*, and *A. thailandica* for the shotgun data, the genera *Actinocatenispora* and *Streptomyces* for the 16S data, and the genera *Phialemonium* and *Apiotrichum* for the ITS data. For leaf samples, taxa contributing to the top ~25% dissimilarity were *M. aeruginosa* and *Streptomyces* sp. M2 for the shotgun data, the genera *Pseudomonas* and *Rhodococcus* for the 16S data, and the genera *Aspergillus* and *Goidanichiella* for the ITS data.

Highly conserved taxa that are present in most samples, typically ~70%, can be considered part of the "core" microbiome that orchestrates the interactions between the host and the microbiota [57]. As described in the methods, we employed strict criteria that the taxa must appear in all of the samples for each condition (Table 2). In the rhizosphere, the core microbiota identified in the shotgun analysis included *Streptomyces* sp. M2 and

*Actinocatenispora sera.* Core taxa in the leaves included *Streptomyces* sp. M2 and '*Candidatus* Liberibacter africanus', both of which persisted across the two different conditions and all samples. The larger number of taxa associated with the rhizosphere ASVs were consistent with the microbes identified by shotgun analysis and indicated bacterial (*Streptomyces*, *Actinocatenispora*, and *Rhodanobacter*) as well as fungal taxa (*Ascomycota*, *Apiotrichum*, *Phialemonium*, and *Candida*) as contributors to the core microbiome. Leaf ASVs revealed that bacteria (*Streptomyces*, *Rhodanobacter*, and *Solimonas*), as well as a single unidentified fungal sequence, comprised the core.

**Table 1.** Similarity of percentage (SIMPER) analysis of microbiota contributing to the top ~25% of dissimilarity (Bray–Curtis) between non-acclimated (NA) and cold-acclimated (CA) samples (showing average relative abundance in %) in both leaf tissue and rhizosphere performed in PAST (v4.08).


**Table 2.** Core microbiota taxa (species or distinct ASVs as indicated) present in 100% of samples for each sequencing and analysis method of shotgun, 16S rRNA, and ITS sequencing methodologies with an average relative abundance >5%.



**Table 2.** *Cont.*

\* '*Candidatus* Liberibacter africanus'.

#### **4. Discussion**

The plant-microbiome partnership is responsive to stress, with the details of the signalling between the kingdoms of Eubacteria, Fungi, and Planta only beginning to be investigated [9,10,58,59]. Sub-zero temperatures are a particular challenge, resulting in cellular dehydration, membrane rupture, and increased vulnerability to psychrophilic pathogens and death, but some perennials respond to earlier non-freezing temperatures, and/or shortened day lengths to initiate a signalling response. This CA stress triggers changes in plant metabolism, resulting in cold-hardening and survival during subsequent freeze events and is accompanied by significant changes in the leaf microbiome community profile, but with less substantial community shifts in the rhizosphere (Figures 1 and 3).

#### *4.1. Little Change in Rhizosphere Communities after Cold Acclimation*

The different sequencing methodologies employed, either amplicon or shotgun analyses, generally yielded compatible results. As indicated, there were few changes in the rhizosphere community after the shift to low temperatures, as shown by the overlapping PCoA groupings with rare exceptions, and for the most part these did not make up a large proportion of the taxa. The rhizosphere communities from both NA and CA plants contained taxa previously reported in bound soils associated with *Brachypodium* and similar to those found in wheat [22]. Some species of the order *Burkholderiales* have been isolated from ryegrass rhizospheres and are associated with nutrient acquisition such that there is interest in their potential as beneficial probiotics for crop enhancement [60]. Ascomycota is dominant in grassland soils, which can be low in organic matter and nutrients, playing key roles in cyanobacteria-dominated soils as well as having important roles in cycling carbon and nitrogen in addition to nutrient transport [61]. The fact that these taxa are shared in wheat and *Brachypodium* underscores the co-evolution of the plant–host relationship, since microbiota in the dicot, *Arabidopsis*, is distinct [22]. As noted, neither the *Brachypodium* bacterial nor fungal communities changed significantly after the plants were moved to 4 ◦C, suggesting that there was insufficient time for the soil to reach that temperature. Indeed, investigations of cold-responsive rhizosphere microbiota in maize used 5 weeks exposure to "chilling" conditions compared to our 6-day treatment [17]. In addition, it is notable that the myriad of CA-dictated changes made in the above-ground portion of *Brachypodium* are not apparently signalled to the rhizosphere during the treatment regimen.

#### *4.2. Shifts in Leaf Communities Accompany Cold Acclimation*

Compared to the rhizosphere, which is relatively protected from rapid abiotic and biotic stresses, leaves are exposed to daily temperature fluctuations, visible and ultraviolet light, herbivore and mechanical damage, and arguably more pathogens. Within two days

of the shift to CA conditions, the *Brachypodium* leaf membrane is protected from freezeinduced electrolyte leakage, contains elevated levels of soluble sugars, and shows changes in the abundance profiles of hundreds of proteins [6]. The leaf community response was also rapid, as revealed by numerous abundance changes in the bacterial and fungal microbiota, as well as in the proportion of individual core taxa, as supported by the distinct groupings shown in PCoAs (Figures 3 and 5; Table 2). Similarly, cold-associated shifts occurred in leaves from European grasslands over winter while the rhizosphere was relatively unchanged [4]. As in the rhizosphere data, results from the two sequencing methods were generally consistent. However, a notable exception was for sequences corresponding to the toxic cyanobacteria *Microcystis aeruginosa*, which were abundant in NA and increased after CA, but only when using the shotgun methodology. It is possible that these sequences were misclassified as chloroplast DNA and were mistakenly filtered from the amplicon data. We speculate that the increase in relative abundance of cyanobacteria after CA is likely due to the reduction in evaporation on the leaf surfaces at low temperatures, consistent with their preference for aquatic habitats, and their known colonization of the phyllosphere [62].

For other taxa, there was clear evidence of a change in relative abundance after CA that was generally consistent irrespective of the sequencing methodology. This included three prominent *Actinobacteria* species that increased in relative read numbers, including the grassland-associated *Actinocatenispora thailandica* and *Actinocatenispora sera*, as well as the mycelium-producing *Streptomyces* sp. M2, a known PGPB [63]. Although present in the rhizosphere samples under both conditions, *Streptomyces* sp. M2 increased 216-fold in relative abundance following CA in leaves. Presumably, it promotes plant growth with its extensive repertoire of antibiotics, plant growth hormones, siderophores, and insecticides [63–65]. Strikingly, this *Streptomyces* strain can inhibit the plant pathogen *P. syringae*, perhaps due to siderophores that chelate iron required by *Pseudomonas* [63]. Such inhibition could explain the disappearance of *P. syringae* after CA treatment, representing a log2 fold change of −8.7.

Other bacteria also showed inverse abundance profiles depending upon the condition, as described in the Results section. Fungal ascomycete taxa similarly exchanged their relative abundance, with a decrease in the genus *Goidanichiella* and an increase in the genus *Aspergillus* detected after CA. These changes may be related to the temperature regime since *Goidanichiella* was reported to dominate summer-collected wheat leaves whereas cold-tolerant *Aspergillus* are of interest as growth promoters likely due to their ability to solubilize phosphates [66,67].

#### *4.3. Leaf Cold Acclimation Associated with Low Temperature and Pathogen Responses*

After transfer to 4 ◦C, the leaf microbiome was impacted by the temperature shift and also showed changes in the relative abundance of potential pathogens. These observations reflect the results of network analysis of hundreds of plasma membrane proteome changes after CA that showed crosstalk between pathways for low temperature stress and disease and defence [6]. *Brachypodium* responds to CA by diverting resources away from growth and to the stress response. It appears then that the host–microbiome works together in a joint effort to prepare for the worsening conditions associated with winter.

One of the most obvious examples of the connection between low temperature and disease is found in the ice nucleation-active plant pathogen *P. syringae*, which can facilitate the formation of ice at temperatures just below 0 ◦C, presumably to lyse plant cells and thus access nutrients [68]. In NA leaves, *P. syringae* was a large contributor to the bacterial taxa (8% of the shotgun reads). However, as the temperature drops, such a large proportion of *P. syringae* in the leaf microbiota would surely present a grave risk to the host plant. Remarkably, after CA there was no evidence of this bacteria. This disappearance is undoubtedly fostered by *Brachypodium*'s defence pathways that lead to the production of multiple proteins, including antifreeze proteins, that target the ice nucleator, but we propose that the microbiome also supports this protective strategy.

Coincident with the collapse of the *P. syringae* population, there was a 216-fold increase in the relative abundance of *Streptomyces* sp. M2 (0.1% to 15.1%). It is important to note that this increase after CA cannot be explained by sensitivity to the NA growth conditions since it is routinely cultured at 30 ◦C [69]. Thus, the change in its abundance is independent of the temperature shift and may be fostered by *Brachypodium*. As mentioned, this PGPB secretes antibiotics and siderophores and is known to inhibit *P. syringae* [63]. *Rhodococcus* also decreased 40-fold in relative abundance, but to date there is no information on its interaction with *Streptomyces* or other plant beneficials. Nevertheless, as well as directly targeting *P. syringae,* it is likely that *Streptomyces* alerts plant defences against other phytopathogens since the inoculation of *Streptomyces* spp. induces the expression of defense-related genes—at least, so it was found to do in a pea crop [70]. This ability could also explain why *Streptomyces* spp. are not limited to inhibition of bacterial species but also inhibit fungal phytopathogens in planta [71,72].

Therefore, in addition to combating the cold-associated pathogen *P. syringae*, *Streptomyces* sp. M2 likely contributes to the overall cold tolerance of *Brachypodium* and thus would be central to the cold-acclimated microbiome. *Streptomyces* spp. have a variety of adaptations for cold resistance, including the production of cold shock proteins and small solutes for cryoprotection [73–75]. These products may assist host survival, since a strain of *Streptomyces* was shown to alleviate the effects of cold stress in turfgrass [19] and drought stress in maize [76]. In addition, BioCyc genome-wide predictions indicate that *Streptomyces* sp. M2 produces key oxidative stress enzymes that can be secreted in *Streptomyces* spp. [77–79]. In addition, *Streptomyces* sp. M2 synthesizes cryoprotective soluble sugars that coincidentally increase rapidly in CA *Brachypodium* [6,80]. The synthesis of the osmoprotectant proline may also benefit host plants, as inoculation of sugarcane with *Streptomyces* increased proline content and drought tolerance [81]. *Streptomyces* spp. are also reported to increase drought tolerance in maize and aid in the accumulation of soluble sugars [76].

Another bacterial taxon, the genus *Solimonas*, increased 3.3-fold after CA, and although these species have a wide temperature optimum, they are characterized by polar lipids and fatty acids, which are known to contribute to cold tolerance [82]. In parallel findings, *Brachypodium* shows changes in metabolic pathways leading to restructuring of the plasma membrane after CA, a common vulnerability for both microbes and their hosts [6,83,84]. Already mentioned was the cold tolerance of the plant-beneficial fungus *Aspergillus.* More insight could be revealed by an investigation of the functional microbiomes of CA *Brachypodium*. However, due to low reads and sequencing depths, our results can only be considered preliminary (see Supplementary File S2 and Figures S7–S9). Nevertheless, in parallel with the CA *Brachypodium* plasma membrane proteome [6], microbial proteins involved in pathways that intersect with low temperature tolerance, such as the synthesis of soluble cryoprotectants, oxidative stress, and pathogen resistance, were detected in the microbiome in response to cold stress. Again, this underscored the inter-dependent and symbiotic character of the CA response.

#### *4.4. Prospects and Conclusions*

Taken together, both the changes in microbial community profiles following CA and the functional role of these plant beneficials suggest that commercial growers could see some benefit from the inoculation of mixed community strains, including *Streptomyces* sp. M2, for protection against *P. syringae* and other phytopathogens, while at the same time benefiting from other plant growth-promoting characteristics as well as enhancing cold resilience. With the presentation of this first CA *Brachypodium* microbiome, it is hoped that the insights gained will inspire treatment options to enhance cold tolerance and other intersecting stresses tailored toward specific agriculturally important grain crops [1,9,85,86].

This special issue of *Plants* asks, "What makes the life of stressed plants a little easier?" The answer for *Brachypodium* undergoing acclimation to low temperature in preparation for the coming winter is very clear. It is the strong partnership with a shifting above-ground

bacterial and fungal community that works in concert with plant networks that intersect cold-, drought-, and antipathogen-signalling pathways to ensure that within only a few days host plants survive freeze events. Not only does it make the life of plants a "little easier", we also argue that it may very well be essential for survival. Therefore, we propose that the battle against winter condition stresses is won by important inter-kingdom partnerships.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/plants10122824/s1. Supplementary File S1: Contains all scripts and commands used. Supplementary File S2: Contains all Supplemental Tables, Supplemental Figures, and Supplemental Text describing functional classification of shotgun data. Figure S1: Bulk soil collection from a farm field. Figure S2: Representative three-week-old *Brachypodium distachyon*. Figure S3: Image showing an example of the tightly bound root soil still attached to the plant. Figure S4: Read statistics of the shotgun sequencing processing for averages of the cold-acclimated and non-acclimated leaf and rhizosphere samples. Figure S5: Principal coordinate analysis plots comparing the taxonomic communities from amplicon sequencing blank kit controls. Figure S6: Correlation plots comparing shotgun sequencing to amplicon sequencing results under both non-acclimated and cold-acclimated conditions in the leaf and rhizosphere samples. Figure S7: Heatmaps showing the average relative abundance of the Pfam domains. Figure S8: Heatmap showing the top 50 most abundant MetaCyc pathways in rhizosphere samples. Figure S9: Heatmap showing the top 50 most abundant MetaCyc pathways in leaf samples. Table S1: DNA, final library concentration, and average library size. Table S2: Summary of quality control and preprocessing of metagenomic reads from shotgun sequencing. Table S3: Summary of the top ten average relative abundant taxa for rhizosphere samples showing average relative abundance for each non-acclimated and cold-acclimated conditions of shotgun, 16S, and ITS sequencing. Table S4: Summary of the top ten average relative abundant taxa for leaf samples showing average relative abundance for each non-acclimated and cold-acclimated conditions of shotgun, 16S, and ITS sequencing.

**Author Contributions:** C.L.J. conducted all experiments, analyzed all data, and produced all figures. C.L.J. wrote the initial draft of the manuscript and all authors contributed to manuscript revision. G.C.d. and V.K.W. supervised the work. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by a Natural Sciences and Engineering Research Council of Canada Discovery grant to V.K.W and funding from Queen's University to G.C.d.

**Institutional Review Board Statement:** This study did not involve humans or animals. *Brachypodium distachyon* plants were used in this study. Seeds of an inbred line, ecotype Bd21, were kindly provided by RIKEN, Wako, Japan. ¯

**Data Availability Statement:** All raw sequences were deposited in the National Centre for Biotechnology Information (NCBI) Sequence Read Archive (SRA) under BioProject ID: PRJNA782211, available at https://www.ncbi.nlm.nih.gov/bioproject/782211 (accessed on 21 November 2021).

**Acknowledgments:** We acknowledge Kristy Moniz for her technical support early in the project and MiGS and MR DNA for their preparation of libraries and sequencing, as well as Scot Dowd of MR DNA for his suggestions and advice.

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

#### **References**


### *Article* **Bonactin and Feigrisolide C Inhibit** *Magnaporthe oryzae Triticum* **Fungus and Control Wheat Blast Disease**

**S. M. Fajle Rabby 1,†, Moutoshi Chakraborty 1,†, Dipali Rani Gupta 1, Mahfuzur Rahman 2, Sanjoy Kumar Paul 1, Nur Uddin Mahmud 1, Abdullah Al Mahbub Rahat 1, Ljupcho Jankuloski <sup>3</sup> and Tofazzal Islam 1,\***

	- 1400 Vienna, Austria

**Abstract:** Wheat blast caused by the *Magnaporthe oryzae Triticum* (MoT) pathotype is one of the most damaging fungal diseases of wheat. During the screening of novel bioactive secondary metabolites, we observed two marine secondary metabolites, bonactin and feigrisolide C, extracted from the marine bacteria *Streptomyces* spp. (Act 8970 and ACT 7619), remarkably inhibited the hyphal growth of an MoT isolate BTJP 4 (5) in vitro. In a further study, we found that bonactin and feigrisolide C reduced the mycelial growth of this highly pathogenic isolate in a dose-dependent manner. Bonactin inhibited the mycelial development of BTJP 4 (5) more effectively than feigrisolide C, with minimal concentrations for inhibition being 0.005 and 0.025 μg/disk, respectively. In a potato dextrose agar (PDA) medium, these marine natural products greatly reduced conidia production in the mycelia. Further bioassays demonstrated that these secondary metabolites could inhibit the MoT conidia germination, triggered lysis, or conidia germinated with abnormally long branched germ tubes that formed atypical appressoria (low melanization) of BTJP 4 (5). Application of these natural products in a field experiment significantly protected wheat from blast disease and increased grain yield compared to the untreated control. As far as we are aware, this is the first report of bonactin and feigrisolide C that inhibited mycelial development, conidia production, conidial germination, and morphological modifications in the germinated conidia of an MoT isolate and suppressed wheat blast disease in vivo. To recommend these compounds as lead compounds or biopesticides for managing wheat blast, more research is needed with additional MoT isolates to identify their exact mode of action and efficacy of disease control in diverse field conditions.

**Keywords:** antifungal secondary metabolites; biocontrol; abnormal germ tube suppression of appressoria; *Streptomyces* sp.

### **1. Introduction**

Wheat is an essential staple dietary source for approximately 2.5 billion individuals in 89 different nations in the world. In low- and middle-income nations, it outperforms maize or rice as a source of protein. Wheat ranks second only to rice in the context of calorie supply. It is a primary food source in North Africa and West and Central Asia, accounting for up to half of the calories consumed (https://wheat.org/; accessed on 16 May 2022). Nonetheless, wheat is prone to various fungal diseases; the most notorious one is a wheat blast, caused by the pathogenic filamentous fungus *Magnaporthe oryzae Triticum* (MoT) pathotype. In 1985, the first case of the wheat blast was recorded in Brazil [1,2]. In 2016, Bangladesh experienced an alarming epidemic of the wheat blast that was the first incidence of the disease in Asia [3]. That epidemic destroyed 15,000 hectares of wheat fields with

**Citation:** Rabby, S.M.F.; Chakraborty, M.; Gupta, D.R.; Rahman, M.; Paul, S.K.; Mahmud, N.U.; Rahat, A.A.M.; Jankuloski, L.; Islam, T. Bonactin and Feigrisolide C Inhibit *Magnaporthe oryzae Triticum* Fungus and Control Wheat Blast Disease. *Plants* **2022**, *11*, 2108. https://doi.org/10.3390/ plants11162108

Academic Editor: Tika Adhikari

Received: 17 May 2022 Accepted: 18 July 2022 Published: 12 August 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

a yield loss of up to 100% [4]. Wheat blast is causing concerns among seed scientists as it has the potential to spread to important wheat-growing areas in South Asian and African countries [5]. Plant pathologists have warned that the disease might spread to India, Pakistan, and China, which are the second-, seventh-, and first-highest wheat producers in the world, respectively [4,6–8].

The MoT is a filamentous haploid ascomycete fungus. Its infection cycle has previously been described [9,10]. Briefly, MoT's three-celled hyaline airborne conidium lands on a wheat leaf and attaches to it using adhesive. It then begins to grow, developing into a slender germ tube with an appressorium at the tip. A tiny penetrating peg develops at the base of the appressorium, compressing the cuticle and allowing entry into the wheat epidermis. Wheat plasma membranes are penetrated by bulky, virulent mycelium, which then enters epidermal cells to complete tissue invasion [10–12]. It affects the aerial parts of the wheat plant, specifically the leaves, stems, nodes, and kernels encompassing all growth phases [7,13,14]. MoT usually affects spikes and bleaches the infected spikes, which results in malformed grains or producing no grain at all [4,15]. Wheat heads with severe infection may die, resulting in a considerable decrease in productivity. The early bleaching of spikelets above the infection point and the whole panicle is the most common symptom [4,7,16]. Contaminated seeds or grains as well as airborne conidia spread this disease, and the pathogen may persist in infected crop residues and seeds [17].

There is an ongoing demand for new plant chemotherapeutic agents that are unique from frequently used fungicides in their underlying mechanisms for advanced plant disease management. Another important reason for these needs is the occurrence of fungicideresistant pathogens, which results from the requirement of using many synthetic fungicides at high rates, with adverse environmental repercussions [18,19]. Several microorganisms have been authorized as biocontrol agents in many countries including the EU to date due to their relatively low toxic residues, environment-friendly properties, and low manufacturing cost [20]. However, scientific research suggests that these benefits are not always achieved as biological pesticides are mostly living organisms, and their performance varies owing to the influence of numerous biotic (nutritional requirements, host species, and pathogenic microbes) and abiotic (moisture, temperature, relative humidity) factors, which limit their fitness under field conditions [21,22]. In addition, some biological control microbes, such as *Bacillus cereus*, are known to cause human diseases, precluding their release in the environment. In this regard, microbial metabolites can be another suitable alternative to live microbes or synthetic fungicides that are also capable of controlling plant diseases with low detrimental effects on human health and the environment [23]. The versatility of biological activity and chemical structure of microbial metabolites as a pesticide is worth considering due to the potential benefits [24]. The second aspect of microbial metabolites as agricultural fungicides is the requirement of a relatively short period for biodegradation. According to Tanaka and Omura [25], they often decay within a month or even a few days, leaving low residue that should be less harmful to the environment. Metabolites derived from diverse microorganisms have been utilized extensively to address commercially important diseases of several plants [26].

Secondary metabolites extracted from *Streptomyces* species have shown a broad range of biological functions by blocking particular enzymes or proteins in signaling cascades [27–30]. Wheat blast management research of our working group took a comprehensive strategy including biologicals and biorational approaches. During the screening of new bioactive natural products against MoT, we discovered that a few metabolites of *Streptomyces* spp. inhibited the growth of MoT mycelia [31]. Two natural secondary metabolites, bonactin and feigrisolide C, extracted from marine *Streptomyces* spp., Act 8970 and ACT 7619, respectively, exhibited substantial growth inhibitory effects against a MoT isolate among many different compounds tested. The first acyclic ester of nonactic acid is bonactin, whereas feigrisolide C is a non-symmetric lactone associated with the nactic acid group [32,33]. Bonactin has shown antimicrobial properties against both bacteria and fungi. Many different microbes including *Bacillus megaterium*, *Klebsiella pneumoniea*, *Escherichia coli*, *Micrococcus luteus*, *Staphylococcus aureus*, *Saccharomyces cerevisiae*, and *Alicagenes faecalis* are sensitive to bonactin [33]. In antiviral, antibacterial, and enzyme inhibition tests, feigrisolides were found to have varying degrees of effectiveness. Synthesis of feigrisolide C has been achieved [34]. Nonactic acid esters are in general environmentally benign since soil microbes convert them to H2O and CO2 [35]. Inhibitory effects of bonactin and feigrisolide C on zoosporogenesis and motility of phytopathogenic Peronosporomycete zoospores have been reported [29]. A few studies have documented the toxicity level of these compounds to date. Bonactin is reported as non-carcinogenic and non-toxic to aquatic model organisms. It has been reported as a suitable natural compound for schizophrenia disorder, suggesting little or no toxicity to humans [36]. However, further research is needed to ascertain their safety for humans and the environment before using them as a potential lead component for the synthesis of agricultural fungicides for controlling wheat blast. There is currently no information available about the use of nonactic acid esters' antimicrobial activities to control wheat blast disease. To our best knowledge, this is the first report of marine natural antibiotics bonactin and feigrisolide C from *Streptomyces* spp. inhibiting a destructive wheat blast causing a MoT isolate and suppressing the disease in field conditions. The major targets of the current study were to: (i) assess the inhibitory effects of bonactin and feigrisolide C on the mycelia growth of BTJP 4 (5); (ii) evaluate the influences of these marine natural products on conidia production, germination, and the developmental transitions of conidia of BTJP 4 (5); (iii) assess the effect of these compounds on the suppression of wheat blast disease development caused by BTJP 4 (5) on leaves and spikes; and (iv) compare the disease inhibition efficiencies of these natural compounds with a commercialized fungicide Nativo®75WG.

#### **2. Results**

#### *2.1. Mycelial Growth Inhibition and Morphological Alteration of Hyphae*

Both bonactin and feigrisolide C considerably inhibited MoT mycelium development in the PDA medium in a dose-dependent manner (Figure 1). Bonactin inhibited mycelium development of the MoT isolate BTJP 4 (5) more efficiently than feigrisolide C. When bonactin and feigrisolide C were applied separately at 2 μg/disk, mycelial growth inhibition was 70.8 ± 0.8% and 68.1 ± 1.0%, respectively (Figure 2). Both bonactin and feigrisolide C demonstrated slightly lower inhibitory capacity than Nativo® WG 75 (82.7 ± 0.6% at 2 μg/disk). The inhibitory effects of these natural compounds enhanced as concentrations were raised from 0.005 to 2 μg/disk, reaching up to 71% for bonactin (Figure 2). Bonactin had more inhibitory efficacy than feigrisolide C but was slightly less effective than Nativo®WG 75 against the BTJP 4 (5) isolate. Both substances were ineffective against MoT at quantities lower than 0.005 μg.

Bonactin extensively impeded BTJP 4 (5) hyphal growth at 2 μg/disk (70.8 ± 0.8%), 1.5 μg/disk (65.4 ± 1.0%), and 1 μg/disk (58.6 ± 1.3%), showing that inhibition and accelerated concentrations had a positive correlation. At 2, 1.5, and 1 μg/disk, feigrisolide C inhibited 68.1 ± 1.0%, 63.4 ± 1.3%, and 54.2 ± 1.0% hyphal growth of BTJP 4 (5). Bonactin and feigrisolide C had minimum suppressive concentrations of 0.005 and 0.025 μg/disk, respectively, and these compounds suppressed mycelial growth by 9.81 ± 1.3% and 13.3 ± 0.8% at 0.005 and 0.025 μg/disk, respectively. However, the minimal inhibitory concentration of Nativo® WG 75 was 0.05 μg/disk, but at higher doses starting at 0.1 μg/disk it outperformed the suppression percentage of the two other test compounds at equal concentrations. It is worth noting that at less than 0.1 μg/disk concentration, bonactin and feigrisolide C inhibited mycelial development more efficiently than Nativo® WG 75, and bonactin inhibited BTJP 4 (5) at a 10-fold lower dose.

**Figure 1.** Mycelial growth suppression and morphological changes of hyphae of a wheat blast fungus, *Magnaporthe oryzae Triticum* (MoT) isolate BTJP 4 (5) approaching the paper disks containing two marine natural products, bonatin and feigrisolide C, and Nativo® WG75 (20 μg/disk), a commercial fungicide known to growers as local standard in Bangladesh. Normal mycelial growth (**a**) of BTJP 4 (5) on PDA plate (10 days) and microscopic view of the growing typical tubular hyphal tips (**a***-* ) in the untreated control. Mycelial growth inhibition (**b**) and abnormal hyphal tips (**b***-* ) closer to the paper disk containing bonactin. Inhibited mycelia (**c**) and curly and irregular growth of hyphal tips (**c***-* ) by feigrisolide C. Mycelial growth inhibition (**d**) and severely damaged hyphal tips (**d***-* ) by the Nativo® WG75. Bar = 50 μm. The micrographs shown in panels A and B were captured with a digital camera (CAMEDIA C-3040 zoom; Olympus Optical Co. Ltd., Tokyo, Japan), and those in panels C and D were taken from a light microscope (IX70-S1F2; Olympus) by using the same digital camera connected to it.

**Figure 2.** Suppression effects of bonactin, feigrisolide C, and Nativo® WG75 on mycelial growth of *Magnaporthe oryzae Triticum* (MoT) isolate BTJP 4 (5) in PDA media. The data represents the mean ± standard errors of three replications for each rate of the test compound based on the Tukey HSD (honest significance difference) test at a 5% level.

Microscopic examinations of untreated BTJP 4 (5) revealed polar, cylindrical growth with smooth, hyaline, branching, plump, septate, and unbroken hyphae (Figure 1a,a ). Hyphae treated with bonactin and feigrisolide C grew irregularly and exhibited a higher frequency of branching per unit of the hyphal length. Cell walls of the hyphae were not smooth but had ridges that gave them a crinkled look as well as causing irregular cell swelling (Figure 1b,b ,c,c ). Similar effects of the fungicide Nativo®WG75 on hyphal growth were observed. Mycelia closer to the filter disk of Nativo®WG75 showed a comparable modification of MoT hyphae (Figure 1d,d ). However, compared to Nativo®WG75, the two natural products generated slightly different morphological aberrations in MoT, suggesting a possibly different mode of action.

#### *2.2. Conidiogenesis Inhibition*

Bonactin, feigrisolide C, and Nativo®WG75 considerably decreased the conidia production of BTJP 4 (5) at concentrations of 1, 5, and 10 μg/mL, respectively, and suppression increased with increasing concentrations from 1 to 5 to 10 g/mL (Figure 3). Almost no or only a few conidia were produced at 10 μg/mL in media amended separately with all three compounds. Microscopic examination also revealed broken hyphal tips and complete suppression of conidiophore formation in fungal colonies in Petri plates that were treated with these three compounds at 10 μg/mL.

**Figure 3.** Effects of bonactin, feigrisolide C, and Nativo® WG75 on suppression of conidiogenesis of *M. oryzae Triticum* isolate BTJP 4 (5) in the 96-multiwell plates at 1 μg/mL, 5 μg/mL, and 10 μg/mL. Image (**a**) control. Images in panels (**b**–**d**) are bonactin, feigrisolide C, and Nativo® WG75, respectively. Bar = 50 μm.

#### *2.3. Inhibition of Conidia Germination and Morphological Aberrations in Germinated Conidia*

To determine the MoT isolate BTJP 4(5)'s conidial germination inhibition capacity of test products, bonactin, feigrisolide C, and Nativo® WG75 were added to the multi-well plates at concentrations of 0.5 μg/mL. The rate of conidial germination was recorded after 6, 12, and 24 h of incubation at 25 ◦C (Table 1). Bonactin and Nativo® WG75 treatments dramatically inhibited conidia germination compared to the control, while no conidia germinated in feigrisolide C-treated plates after 6 h of incubation. All (100%) conidia germinated in the water while it was 49.7 ± 0.6% in Nativo® WG75-treated plates. The BTJP 4 (5)' conidia germination rates with bonactin and feigrisolide C were 79.1 ± 0.6% and 0 ± 0%, respectively, at 0.5 μg/mL.


**Table 1.** In vitro effects of bonactin and feigrisolide C on conidia germination and the developmental transitions of *M. oryzae Triticum* (MoT) isolate BTJP 4 (5) at 0.5 μg/mL.

<sup>a</sup> Data are mean value ± SE of three replications in each natural compound. Means within a column followed by a different letter(s) are significantly different according to Tukey's HSD (honest significance difference) post-hoc (*p* ≤ 0.05).

In the dark at 25 ◦C, 100% of conidia germination occurred in water during all incubation periods (6 h, 12 h, and 24 h), with normal germ tube and mycelial growth (Table 1, Figure 4a). At 0.5 μg/mL, both bonactin (panel b) and feigrisolide C reduced on the germination of the conidia and the post-germination developmental processes, resulting in abnormal transitions from one stage to another. During 6 h of incubation in the presence of bonactin, the conidia germination rate was 79.1 ± 0.6%, which had short germ tubes. After 12 h, 12.7 ± 0.4% of normal germ tubes were observed, whereas 66.5 ± 0.5% had abnormally long branched germ tubes. After 24 h, there were 9.5 ± 0.2% normal appressoria and 60.1 ± 0.3% atypical appressoria (low melanization), without any hyphal development (Table 1, Figure 4b).

In the case of feigrisolide C, 7.4 ± 0.5% of the conidia lysed after 6 h, and no germination occurred between 6 h and 24 h (Table 1, Figure 5c). In the presence of Nativo® WG75, 49.7 ± 0.6% of conidia germinated with normal germ tubes after 6 and 12 h, but no appressorial development took place. Nativo® WG75 also inhibited sporulation similar to feigrisolide C to prevent further mycelial growth after 24 h (Table 1, Figure 5A). It is worth mentioning that these compounds resulted in excessively long branching in germ tubes and conidia lysis, whereas Nativo® WG75 had no such effect.

**Figure 4.** Micrographs showing the changes in germination and developmental transitions of MoT conidia with time-course in the untreated control (panel (**a**)) and the presence of bonactin (panel (**b**)), feigrisolide C (panel (**c**)), and a commercial fungicide Nativo® WG75 (panel (**d**)) at 0.5 μg/mL. Bar = 10 μm.

**Figure 5.** (**A**). Representative images showing wheat blast disease (symptoms) suppression by varying doses (1–10 μg/mL) of bonactin, feigrisolide C, and Nativo® WG75. The compounds were liquefied in 1% DMSO and applied on the detached leaves of wheat (cv. BARI Gom 26) 24 h before artificial point inoculation with 20 <sup>μ</sup>L/point of suspension of conidia containing 1 <sup>×</sup> 105 conidia/mL. (a) Control, 1% DMSO, (b) bonatin, (c) feigrisolide C, (d) Nativo® WG75, and (e) uninoculated and untreated leaf. (**B**) Average lengths of blast lesions on detached wheat leaves pretreated with bonatin, feigrisolide C, and Nativo®WG75 compared to 1% DMSO treatment control. The data are the means ± standard errors of at least five replications for each dosage of the compounds at *p* ≤ 0.05. Vertical bars represent ± standard error.

#### *2.4. Wheat Blast Progression on Excised Wheat Leaves*

The two metabolites applied at 1, 5, and 10 μg/mL considerably decreased the wheat blast disease symptoms in excised leaves of wheat infected with BTJP 4 (5). The lesion lengths in the leaves pretreated with bonactin were 5.3 ± 0.2 mm at 1 μg/mL and 1.2 ± 0.2 mm at 5 μg/mL, respectively (Figure 5A,B). The blast lesion lengths with feigrisolide C were 6.1 ± 0.2 mm and 2.5 ± 0.1 mm at 1 μg/mL and 5 μg/mL, respectively (Figure 5A,B). Leaves of wheat treated with bonactin, feigrisolide C, and Nativo®WG75 at 10 μg/mL did not show any blast symptoms (Figure 5A,B). Normal blast lesions were visible on the water-treated leaves with average lengths of 9.3 ± 0.2 mm (Figure 5A,B). In comparison to both compounds, the fungicide effectively reduced lesion progression at 1 and 5 μg/mL.

#### *2.5. Wheat Blast Disease Suppression in the Field at the Heading Stage*

To determine the efficacy of these compounds in suppressing blast disease in artificially infected wheat spikes by BTJP 4 (5), a field experiment was conducted by using a commercial fungicide Nativo®75WG at 50 μg/mL as a local standard. In the field, bonactin and feigrisolide C considerably reduced wheat blast disease incidences (41% and 51.3%, respectively) (Figure 6c,d, Table 2), compared to 87.3% disease incidence in the untreated control (Figure 6b, Table 2).

**Figure 6.** Inhibition of wheat blast disease with bonactin, feigrisolide C at 5 μg/mL, and Nativo® 75WG at 50 μg/mL; (**a**) Uninoculated, untreated spike, (**b**) BTJP 4 (5) inoculation + water control + (**c**) bonactin + BTJP 4 (5) inoculation, (**d**) feigrisolide C + BTJP 4 (5) inoculation, (**e**) Nativo®75WG + BTJP 4 (5) inoculation.

**Table 2.** Effect of bonactin and feigrisolide C on wheat (variety-BARI Gom-26) yield and yield components in field conditions following the artificial inoculation with BTJP 4 (5).


\* Yield data are the mean ± SE collected from five replications of each treatment of the test compounds. Data followed by the same letter in a column are not significantly different according to Tukey HSD (honest significance difference) post-hoc statistic at the 5% level.

Furthermore, 32.3 ± 2.40% and 38.6 ± 1.20% blast severities were recorded in wheat plants pretreated with these compounds in comparison to 82.6% in the untreated control. Bonactin (112.9 ± 2.26 gm), feigrisolide C (106.4 ± 2.58 gm), and Nativo® 75WG (126.1 ± 2.70 gm) had significantly increased grain yields compared to the untreated control (64.6 ± 1.71 gm). Grain yields in the Nativo® 75WG were statistically similar to the healthy control (133.1 ± 2.33 gm). Nevertheless, both bioactive natural compounds' treatments had statistically lower but similar grain yields compared to the Nativo® 75WG fungicide and healthy control (Table 2).

Thousand-grain weights for Nativo® 75WG, feigrisolide C, bonactin, and the negative control were 43.2 ± 2.52, 38.7 ± 3.16, 40.1 ± 1.72, and 46.6 ± 1.57 gm, respectively. Grain yields in treated plots were considerably greater than the yield of the untreated control plot (31.7 ± 1.29 gm) (Table 2).

#### **3. Discussion**

In this study, we demonstrated for the first time that two nonactic acid esters extracted from marine *Streptomyces* spp. and named bonactin and feigrisolide C inhibited the growth and development of a destructive wheat blast pathogen *M. oryzae Triticum* (MoT) isolate BTJP 4 (5). Additionally, we discovered that these natural compounds were comparable to the commercial fungicide Nativo® WG75 in their efficacy in successfully reducing wheat blast disease in wheat leaves and spikes that had been artificially inoculated by BTJP 4 (5). These treatments also resulted in a modest increase in grain yield although the highest yield was obtained from fungicide treatment followed by two test compounds. Formation of conidia asexually in hyphal conidiophore and germination of conidia are critical for plant infection by the blast fungus [37–40]. Suppression of hyphal growth, conidia formation, and germination of many fungi, such as rice and wheat blast fungi, by various natural products, have been reported [29,31,33,41–45]. The nonactic acid esters are precursors of macrotetrolide antibiotics which have a broad spectrum of antimicrobial, anticancer, acaricidal, insecticidal, immunosuppressive, antiprotozoan (coccidiostatic), and antiparasitic properties [35,46–48]. In the current study, we did not focus on unraveling the underlying molecular mechanism associated with in vitro growth inhibition of wheat blast causing fungal pathogen and suppression of the disease in vivo. However, from a similar study, Islam et al. [29] found that the hydrolysis of mitochondrial ATP via increased ATPase function was likely associated with the mode of action of antimicrobial activities of macrotetrolides against phytopathogenic Peronosporomycete zoospores. Despite having outstanding biological properties, macrotetrolides have received extremely less attention in plant protection studies. To the best of our knowledge, it is the first report of two natural bioactive nonactic acid esters and precursors of macrotetrolides (bonactin and feigrisolide C) originated from marine *Streptomyces* spp. suppressing the highly aggressive wheat blast pathogen MoT isolate BTJP 4 (5) in vitro and in vivo. Additional study is needed to test the efficacy of these compounds against other strains of MoT as well as whether their antiblast activities are linked with the induction of increased mitochondrial ATPase activity in the asexual spores and hyphae of MoT.

One of the key discoveries of this study is that at almost equal concentrations of Nativo® WG75, both bonactin and feigrisolide C dramatically reduced hyphal growth, conidia production, and germination, and also caused morphological changes in germinated conidia. Our findings indicate that these natural substances inhibited conidial germination and mycelium growth, which consequently suppressed wheat blast disease in vivo.

The swelling phenomenon by these compounds on BTJP 4 (5) hyphae is another remarkable observation from our study (Figure 1b'–d'). We utilized doses ranging from 0.005 to 2 μg/disk in our experiment. Swelling increased with increased concentrations, showing a positive correlation of swelling with concentrations. Tensin [49], fengycin [50], gageopeptides, gageotetrin [44], and oligomycins [31] have all been reported to induce developmental aberrations in the tubular growth of the fungal hyphae. Developmental transitions in *Aphanomyces cochlioides* hyphae, such as increased swelling and excessive

branching, have been observed in response to xanthobaccin A from *Lysobacter* sp. SB-K88 or m *Pseudomonas fluorescence* phloroglucinols [51–54]. According to Schumacher et al. [33], bonactin from *Streptomyces* sp. greatly suppressed the hyphal development of *Saccharomyces cerevisiae*, but no data on the mycelial growth inhibitory activity of feigrisolide C has been documented to date. So far, this is known to be the first report of some nonactic acid and nonactic acid ester exhibiting swollen-like abnormal hyphae against a destructive wheat pathogen.

Conidiogenesis is the process of producing conidia, which are fungal spores that are grown asexually on the conidiophore [39]. The majority of fungal plant pathogens attack plants by these asexual spores. Inhibiting or preventing conidiogenesis and conidia germination can reduce the likelihood of host infection by fungal pathogens [55,56]. Future plant protection strategies should explore and rely on similar natural compounds that interfere with these processes. Therefore, another noteworthy finding from this study was that these compounds greatly decreased conidiogenesis (Figure 3), and conidial germination, and also triggered morphological alterations of BTJP 4 (5)'s conidia (Table 1, Figure 4).

Lysis of conidia and uneven branching of germ tube tips as well as unusually long hypha-like germ tubes were among the other distinct and interrelated phenomena found in this work (Figure 4B,C). Dame and co-workers [57] discovered a similar occurrence when they found that oligomycins derived from a marine *Streptomyces* sp. triggered lysis of phytopathogenic *Plasmopara viticola* zoospores that causes grapevine downy mildew disease. Homma and colleagues reported that lecithin induced abnormal branching in germ tube tips of rice blast fungus, and prevented the development of appressoria [58]. Similarly, *A. cochlioides*' cystospores germinated with hyperbranched germ tubes by the effects of diacetylphloroglucinol (DAPG) [54]. Bonactin caused atypical appressoria (low melanization), which restricted MoT fungal infection since appressorium melanization is essential for *M. oryzae* pathogenicity [11]. This compound may affect the gene expression related to the synthesis of melanin. This is also the first study to show that two esters impeded conidiogenesis, germination, and the development of appressoria of BTJP 4 (5) conidia. Future research should concentrate on the mechanisms by which these compounds suppress conidia formation, germination, and appressorium formation of MoT, as well as the impact of these natural bioactive compounds on the expression of genes associated with conidia germination and appressorium formation of BTJP 4 (5) or similar MoT isolates.

Nonactic acid esters are relatively safe for the environment since soil microorganisms can quickly convert them to H2O and CO2 [59]. Plant growth stimulation and specific insecticidal actions of nonactin antibiotic precursors have been documented [35,60]. Bonactin was reported to have antibacterial action and also antifungal action [33]. In a lab investigation, we noticed that nonactin had remarkable antifungal properties against MoT both in vivo and in vitro (our unpublished data). According to Schumacher et al. [33], antimicrobial activity can be achieved without the requirement for a macrotetrolide ring structure, such as the non-asymmetric lactone feigrisolide C, which has antibacterial and antiviral properties [61]. Islam and his colleagues [29] discovered that bonactin and feigrisolide C with other known macroletrolides suppress zoosporogenesis, hamper motility, as well as trigger lysis of *Plasmopara viticola* zoospores. The findings of the current work do not elucidate the detailed mechanism of action, but they do suggest that stimulation of ATPase activities in mitochondria or/and imbalance/translocation of cell cations could inhibit hyphal development and impede conidia germination. Identifying the role of ATPase in inhibiting hyphal growth, conidiogenesis, conidial germination, and appressoria formation may aid in our understanding of the biology and pathogenesis of filamentous plant pathogens. This naturally occurring ATPase inducer may thus be a promising pioneer ingredient for developing novel, efficient agrochemicals to fight this aggressive fungal pathogen.

In this study, wheat leaves pretreated with the test compounds showed shorter lesions than untreated checks (Figure 5). The majority of those lesions were small, and appeared as brown patches with spots of a pinhead size (scale 1) to roundish and fairly expanded grey dots that ranged in size from 1–2 mm in diameter (scale 3). The untreated control leaves had typical blast lesions covering 26–50% of the leaf surface (scale 7), according to the 9-scale blast disease assessment system developed by the IRRI SES (standard evaluation system) [55,56]. However, in the Nativo® WG75 treatment, no visible blast lesions were present. When disease control studies were conducted at the wheat heading stage, similar results were obtained. In artificially infected wheat spikes, blast disease progression was dramatically inhibited by bonactin and feigrisolide C (Figure 6). A popular systemic fungicide, Nativo® WG75, was used in this study as a local standard and positive control. In terms of suppression efficacy of the MoT fungus, the two marine natural compounds evaluated in the current study were comparable to that of the commercial fungicide. The active components of Nativo® WG75 are tebuconazole and trifloxystrobin. Belonging to the systemic triazole fungicide group, tebuconazole's mode of action is known as demethylase inhibitor (DMI). The development of the fungus is slowed down and can eventually be killed as DMI fungicides interfere with the production of sterol in fungal cell walls [62]. Trifloxystrobin, a fungicide in the strobilurin group, suppresses the spore germination of phytopathogenic fungi by disrupting energy production through blocking mitochondrial electron transport [62]. The modes of action of bonactin and feigrisolide C are possibly distinct from Nativo®WG75, despite the observation of a similar disease inhibition response. More research is needed to determine the fundamental mechanism through which these compounds suppress wheat blast. Before acknowledging these compounds as prospective fungicides for wheat blast, a large-scale field evaluation of their efficiency in preventing wheat blast infection is necessary. Recently, it has been found that secondary metabolites from both marine and terrestrial species can biologically suppress the wheat blast disease [55,56].

In today's agriculture, the development of fungicide resistance across pathogenic microorganisms is a major concern. Due to the inappropriate use of fungicides with a single-site active mode of action such as triazole and strobilurin (QoI), some resistant MoT mutant species have been found widely distributed [19]. Investigators are actively searching for new, effective antifungal chemicals possessing alternative modes of action to protect wheat plants against this lethal pathogenic fungus due to the risk of resistance development in conventional fungicides. The marine natural products, bonactin and feigrisolide C, exhibited almost equivalent bioactivity to the commercialized fungicide Nativo®WG75. The effectiveness of these compounds as inhibitors of the MoT isolate BTJP 4 (5) has suggested using them as candidates for agrochemical with a novel mode of action towards this wheat pathogenic fungus provided they are equally effective on other MoT strains under various agro-ecological regions. Further studies are needed with structurally diverse nactic acids, their esters, and macrotetrolides for understanding the structureactivity relationship of bonactin and feigrisolide C. However, very few reports have been published yet regarding their impacts on humans and the environment, and more research is also required to assess their toxicity level before using them to produce fungicides.

#### **4. Materials and Methods**

#### *4.1. Fungal Isolate, the Revival of a Synthetic Medium, and Host Plant Materials*

In 2016, during the first wheat blast epidemic in Jhenaidah, Bangladesh, we collected many MoT strains including BTJP 4 (5) from wheat cv. Prodip (BARI Gom-24) that showed blast infection on spikelets These isolates were preserved at 4 ◦C on dried filter paper for later use. We revived five isolates on a potato dextrose agar (PDA) medium and tested them in the lab for their normal colony characters and aggressiveness to select a representative one (BTJP 4) for this work (4). We also tested isolates collected from the field-infected wheat from 2016 to 2022 and found that they were equally sensitive to the commercial fungicide, Nativo® WG 75 (Figure 7). It appeared the clonal population introduced in Bangladesh from South America had not been mutated [37]. Therefore, we chose BTJP 4(5) for the whole study. On a potato dextrose agar (PDA) medium, the selected isolate was grown for seven days at 25 ◦C. Ten-days-old PDA-grown fungi fungal colonies were washed in an aseptic environment in a laminar flow hood with 500 mL of deionized water to remove aerial mycelia, and then kept at ambient temperature (25–30 ◦C) for 2–3 days to induce abundant conidia production [4,40,55,56]. The conidia were scraped out from each plate using a glass slide after adding 15 mL of water to each plate. Two-layer cheesecloth was used to filter out the hyphal mass, and the dilution was conducted to achieve 1 × <sup>10</sup><sup>5</sup> conidia/mL. Conidial germination was examined under a compound microscope by counting the number. Seedlings of blast disease-susceptible wheat variety Prodip (BARI Gom-24) at the five-leaf stage were used for the bioassay on leaves [31,55,56]. For assessing the wheat blast disease suppression efficacy of bonactin and feigrisolide C, these compounds were sprayed on field-grown wheat spikes at the flowering stage one day before inoculation of the plants with MoT conidia. The detailed method of artificial inoculation of wheat plants by MoT conidia was described recently by Paul et al. [40].

**Figure 7.** Sensitivity of different strains (**1**–**5**) of wheat blast fungus *Mahnaporthe oryzae Triticum* obtained from the field-infected spikes to various doses of a commercial fungicide Nativo. **1**, BTKP 22(3) collected in 2022; **2**, BTJP 194-2 collected in 2019; **3**, BTJP 1910-3 collected in 2019; **4**, BTJP 2 g collected in 2017; and **5**, BTJP 4 (5) collected in 2016. The PDA plates were cultured at 25 ◦C for 3 days after inoculation of the plates by various wheat blast strains.

#### *4.2. Chemicals*

Bonactin and feigrisolide C (Figure 8) were derived from the marine bacteria *Streptomyces* spp. Act 8970 and ACT 7619. Dr. Hartmut Laatsch, a Professor of Georg-August-Universitaet Goettingen in Germany, generously provided these pure chemicals as gifts [29]. The fungicide Nativo® WG 75 (50:50 mixtures of trifloxystrobin and tebuconazole) was purchased in Dhaka, Bangladesh from Bayer Crop Science Ltd. Stock solutions of test compounds were prepared using small amounts of DMSO (dimethyl sulfoxide), and then the solutions were diluted with water. The final mixture included a maximum of 1% (*v*/*v*) DMSO, which had no impact on the development or sporulation of BTJP 4 (5) mycelium [55,56].

**Figure 8.** Structures of bonactin (**A**) and feigrisolide C (**B**).

#### *4.3. Suppression of Mycelial Growth and Hyphal Morphological Alteration*

Using a modified disk diffusion technique as reported by Chakraborty et al. [31], the mycelial growth inhibition of MoT isolate BTJP 4(5) was determined by the application of bonactin, feigrisolide C, and the commercial fungicide Nativo®WG75 on filter paper disks. To prepare a range of concentrations from 0.005 to 2 μg/disk, the required amounts of natural compounds and the fungicide Nativo® WG75 were dissolved in ethyl acetate and water. Nine-millimeter-diameter filter paper disks (Sigma-Aldrich Co., St. Louis, MO, USA) were used to absorb the test compound solutions. In 9 cm-diameter Petri dishes with 10 mL of PDA, the treated disks were placed 2 cm apart from one side. The filter paper disks containing the test chemicals were placed on the opposite side of the actively growing 5 mm-diameter, 7-days-old mycelial plugs of BTJP 4(5). Petri dishes with fungal hyphal plugs against filter paper disks with Nativo®WG75 were used as a control. As a negative control, filter paper disks were coated with ethyl acetate and then allowed to evaporate at ambient room temperature. A fungal hyphal development reduction was observed after 10 days of culture. The Petri plates used as untreated controls were incubated at 25 ◦C until the fungus had colonized and covered the whole surface of the agar. The test was conducted five times with five replications for each concentration. Using a ruler and two perpendicular lines drawn on the lower side of each plate, the radial growth of the fungal culture was measured in centimeters. Measurements were also recorded for the inhibition zone and associated fungal colony diameter influenced by the test compounds and the fungicide. Inhibition percentage radial growth (RGIP) [55,56] was calculated as:

$$\text{RGIP} \%= \frac{\text{Control plate radial growth} - \text{Treated plate radial growth}}{\text{Control plate radial growth}} \times 100$$

Results including radial growth suppression from the disk diffusion test were captured using a digital camera of CAMEDIA C-3040 zoom. At 40× and 100× magnification, an Olympus IX70-S1F2 microscope was used to study the mycelial morphology at the sharp end of the cultures approaching the control and treated disks. The mycelial growth including aberration was photographed using the same digital camera attached to the microscope.

#### *4.4. Suppression of Conidiogenesis*

The stock solutions of each compound were prepared in 10 μL of DMSO and then diluted with distillate water to obtain concentrations of 1, 5, and 10 μg/ml. The final mixtures had a maximum of 1% (*v*/*v*) of DMSO, which had no impact on BTJP 4 (5) sporulation or hyphal development. A 5 mL solution of Nativo®WG75 was prepared to achieve each 1, 5, and 10 μg/mL concentrations by dissolving the required amount of formulation in distilled water that was used as a positive control. A conidiogenesis inhibition test of a MoT isolate was established in our lab and used for this work [31,39,56]. Briefly, to deplete nutrients and promote conidiogenesis, the mycelium of a 10-day-old BTJP 4 (5) Petri plate culture was rinsed [4,39]. After being treated, ten mm BTJP 4 (5) hyphal agar blocks were treated with 50 μL of each test compound and Nativo®WG75 at the aforementioned doses and then placed on Nunc multi-well plates. The mycelial agar block of MoT with 1% DMSO in the same amount of sterile water was used as a negative control. Treated BTJP 4 (5) mycelial plugs were incubated at 28 ◦C and >90% RH under alternating light and dark cycles for 14 and 10 h, respectively. After 24 h, conidiogenesis was observed under a 40× Zeiss Primo Star microscope for analysis, and pictures were taken with a Zeiss Axiocam ERc 5s. With five replications for each treatment, the test was repeated five times.

#### *4.5. Suppression of Conidial Germination and Morphological Changes in Germinated Conidia*

Each natural compound was first liquefied in 10 μL of DMSO before being diluted with distilled water to a concentration of 0.1 μg/mL. As a positive control, a 0.1 μg/mL solution of Nativo®WG75 was prepared in distilled water. We used the methodology developed previously by us for MoT isolate conidial germination investigations [31,55,56]. Briefly, a

<sup>100</sup> <sup>μ</sup>L solution containing 1 × <sup>10</sup><sup>5</sup> conidia/mL of BTJP 4 (5) was directly mixed with a 100 μL solution containing 0.1 μg/mL of product to obtain a 200 μL final solution in the well of a 96-multiwell plate containing test compounds comprising 0.5 μg/mL. Immediately after blending with a glass rod, the suspension was incubated for 6, 12, and 24 h at 25 ◦C in a Ziploc plastic bag with layers of moist paper towel. Sterile water that contained 1% DMSO was employed as a control. A total of 100 conidia from each of the five replications were examined with a Zeiss Primo Star microscope at a 100× magnification. The photographs were acquired with a Zeiss Axiocam ERc 5s, and the percentage of conidia germination, and the morphological alterations of spore germ tubes and appressoria, were determined. The experiment was repeated five times, with at least five replications for each treatment. The conidia germination percentage was calculated as: CG% = (C − T)/C × 100; where %CG = conidia germination, C = average conidia germination percentage in control, and T = average conidia germination percentage in treated samples.

#### *4.6. Wheat Blast Progression on Detached Wheat Leaves*

Bonactin and feigrisolide C stock solutions were made using a small quantity of DMSO. The final DMSO content never exceeded 1% when the natural substances were dissolved in sterile distilled water to obtain concentrations of 1, 5, and 10 μg/mL. Nativo®WG75 was prepared in concentrations of 1, 5, and 10 μg/mL as well. As a negative control, sterilized water that contained 1% DMSO was utilized. This experiment was carried out according to the procedures outlined by Chakraborty et al. [31,55,56]. The first step was to separate wheat leaves from seedlings at the five-leaf stage and place them on plates covered with wet paper towels. Each leaf was treated with three 20 μL drops of the appropriately prepared test compound at the aforementioned concentrations, and the leaves were left to dry for 15 min. Following that, inoculation was conducted on each spot with 1 μL conidial solution containing 1 × <sup>10</sup><sup>5</sup> BTJP 4 (5) conidia/mL, and the plates were incubated at 28 ◦<sup>C</sup> in the darkness for the first 30 h, then under constant lighting for the following two days. The experiment was repeated five times with five different samples each time. For each treatment and compound concentration, the diameter of blast lesions induced by MoT was measured on three leaves per experiment.

#### *4.7. Determination of Wheat Blast Control Efficacy of Bonactin and Feigrisolide C under Field Conditions*

#### 4.7.1. Soil Preparation and Seed Sowing

The experiment was carried out in the research field of the Bangabandhu Sheikh Mujibur Rahman Agricultural University (BSMRAU) in Gazipur, Bangladesh. The trial site was situated 8.4 m above sea level at a latitude of 24.09◦ north, and a longitude of 90.26◦ east. Weeds and stubbles were pulled out of the soil after it had been gently plowed. During soil preparation, adequate quantities of well-decomposed cow dung were applied. Gypsum, muriate of potash, triple super phosphate, and urea were applied as chemical fertilizers at a rate of 11-50-28-70 kg/ha. [63]. 3 to 4 days before seed sowing, the final soil preparation included the application of additional fertilizers along with two-thirds of the urea as a baseline dose. 20 days after the first irrigation, the final one-third of the urea was applied. In the first week of December, BARI Gom-26 wheat seeds were sown. Before sowing, the seeds were treated with Vitavex 200 (3 g/kg seed). There were three replications per treatment and the size of the experimental plot was 1 m2. All of the plots were properly labeled. The required irrigation work was performed, along with additional cross-cultural tasks. The experiment was conducted using a randomized complete block design (RCBD).

#### 4.7.2. Infection Assay in the Wheat Reproductive Phase

The test compounds were applied at a concentration of 5 μg/mL in each plot and allowed to dry overnight, whereas sterile water containing 1% DMSO served as a negative control. BTJP 4 (5) spore suspension was sprayed to wheat fields immediately after

flowering. The positive control was the fungicide Nativo® 75WG, whereas the negative control was deionized distillate water. To establish a humid atmosphere suitable for spore germination, polyethylene sheets were placed over plots before inoculation.

#### 4.7.3. Data Collection and Analysis for Disease Severity

During the reproductive phase, data were recorded on the total number of tillers, productive tillers, infected tillers per hill, the full length of the spikes, diseased area of the spikes, seeds per spike, 1000-grain weight, and grain production per hill. During the vegetative phase, data were collected on the total number of seedlings, the number of infected seedlings per pot, the overall length of the leaves, and the infected area of the leaves. The disease intensity (DI) was calculated using the formula:

$$\text{DI} = \frac{\text{Total infected plants}}{\text{Total plants observed}} \times 100$$

A 5-point scale was used to assess the severity of blast disease, with % infection accounting for the length of the spike that was infected by blast. The scales were 0 for the absence of lesions, 1 for infection rates between 1% and 25%, 2 for infection rates between 26% and 50%, 3 for infection rates between 51% and 75%, and 4 for infection rates between 76% and 100% on the length of damaged leaves. Blast severity was measured by the following formula:

$$\text{DS} = \frac{n \times \text{v}}{\text{N} \times \text{V}} \times 100\%$$

where DS = disease severity *n* = number of blast-infected leaves v = value score for blast severity N = number of observed leaves V = value of highest score.

#### *4.8. Statistical Analysis, Experimental Design, and Replications*

The efficacy of the pure compounds was examined in the laboratory and the field, respectively, using completely randomized design (CRD) and randomized complete block design (RCBD). All statistical analyses were performed using Microsoft Office Excel 2015 and IBM SPSS Statistics 25. Tukey's HSD (honest significance difference) test was used to compare the treatment means. The tables and figures utilized the mean value ± standard error and there were five replications per treatment.

#### **5. Conclusions**

In this study, we demonstrated for the first time that marine natural products, bonactin and feigrisolide C, from *Streptomyces* species, suppressed the mycelial growth and asexual development of an isolate of MoT fungus and inhibited the progression of wheat blast disease caused by that isolate in vivo. Large-scale in vitro and field testing of these compounds with multiple isolates is necessary to determine whether they are potential candidates or lead compounds for developing an effective fungicide against wheat blast disease. More investigation is also needed to determine their level of toxicity towards humans and the environment, as well as their specific method of action and the structure– activity association between these bioactive natural compounds and the wheat blast fungus *M. oryzae Triticum*.

**Author Contributions:** Conceptualization and methodology, T.I.; methodology, S.M.F.R., A.A.M.R. and M.C.; formal analysis, M.C., S.M.F.R., S.K.P., A.A.M.R. and N.U.M.; software, M.C., S.M.F.R., S.K.P. and N.U.M.; validation, T.I.; writing—original draft preparation, M.C., S.M.F.R. and S.K.P.; writing—review and editing, T.I., D.R.G., M.R. and L.J.; supervision, T.I. and D.R.G.; project administration, T.I.; funding acquisition, T.I., M.R. and L.J. All authors have read and agreed to the published version of the manuscript.

**Funding:** The Krishi Gobeshona Foundation (KGF) of Bangladesh provided funding for this study to Tofazzal Islam of the Institute of Biotechnology and Genetic Engineering (IBGE) at BSMRAU, Bangladesh, under project Nos. KGF TF50-C/17 and TF 92-FNS/21.

**Institutional Review Board Statement:** This research content included no human participants and/or animals.

**Informed Consent Statement:** This manuscript has not been published or presented elsewhere in part or entirety and is not under consideration by another journal. We have read and understood your journal policies and we believe that neither the manuscript nor the study violates any of these. All the authors have been personally and actively involved in substantive work leading to the manuscript and will hold themselves jointly and individually responsible for its content. All co-authors agreed to this submission.

**Data Availability Statement:** The manuscript includes all the data.

**Conflicts of Interest:** The authors disclose that they have no competing interests.

#### **References**


### *Review* **An Insight into the Abiotic Stress Responses of Cultivated Beets (***Beta vulgaris* **L.)**

**Seher Yolcu 1,\*,†, Hemasundar Alavilli 2,\*,†, Pushpalatha Ganesh 3, Muhammad Asif 1, Manu Kumar <sup>4</sup> and Kihwan Song 2,\***


**Abstract:** Cultivated beets (sugar beets, fodder beets, leaf beets, and garden beets) belonging to the species *Beta vulgaris* L. are important sources for many products such as sugar, bioethanol, animal feed, human nutrition, pulp residue, pectin extract, and molasses. *Beta maritima* L. (sea beet or wild beet) is a halophytic wild ancestor of all cultivated beets. With a requirement of less water and having shorter growth period than sugarcane, cultivated beets are preferentially spreading from temperate regions to subtropical countries. The beet cultivars display tolerance to several abiotic stresses such as salt, drought, cold, heat, and heavy metals. However, many environmental factors adversely influence growth, yield, and quality of beets. Hence, selection of stress-tolerant beet varieties and knowledge on the response mechanisms of beet cultivars to different abiotic stress factors are most required. The present review discusses morpho-physiological, biochemical, and molecular responses of cultivated beets (*B. vulgaris* L.) to different abiotic stresses including alkaline, cold, heat, heavy metals, and UV radiation. Additionally, we describe the beet genes reported for their involvement in response to these stress conditions.

**Keywords:** beet cultivation; abiotic stress; alkaline; cold; heat; heavy metals; stress tolerance; ultraviolet radiation

#### **1. Introduction**

Economically important cultivated beets such as fodder beets, sugar beets, garden beets (e.g., red beet), and leaf beets (e.g., Swiss chard) belong to the sub-species *Beta vulgaris* L. ssp. *vulgaris* [1,2]. All beets originate from a halophytic plant, *Beta vulgaris* L. ssp. *maritima* (sea beet or wild beet), also known as *Beta maritima* L. [3]. Among them, leaf beets and garden beets are used as vegetables [2,4], fodder beets as animal feed [1,2], and sugar beets serve as the source of sucrose, bioethanol, biodegradable polymers, and biofertilizers [5–8]. In addition to these advantages, beets such as Swiss chard and red beet are a rich source of pigments, termed betalains [9–12]. Cultivation of beets is widely distributed throughout Turkey and Mediterranean and European countries [13]. Fodder beet plants, which grow at a temperature between 8 ◦C and 25 ◦C [1], are cultivated in coastal areas of many countries [14] as well as continental habitats [15]. Wild beet (*Beta maritima* L.) is especially distributed along the coasts of Mediterranean Sea and the European North Atlantic Ocean [3], and it shows significantly higher salt tolerance during germination and seedling stages when compared to other beet varieties [15–19]. Although previous reports have shown genetic diversity in beet species, due to insufficient genetic

**Citation:** Yolcu, S.; Alavilli, H.; Ganesh, P.; Asif, M.; Kumar, M.; Song, K. An Insight into the Abiotic Stress Responses of Cultivated Beets (*Beta vulgaris* L.). *Plants* **2022**, *11*, 12. https://doi.org/10.3390/ plants11010012

Academic Editors: Ewa Muszy ´nska, Kinga Dziurka and Mateusz Labudda

Received: 6 November 2021 Accepted: 14 December 2021 Published: 21 December 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

variation in cultivated beets [15,20,21], the use of wild beet can provide a remarkable source of genetic variability for crop improvement under stressful conditions [20].

Crop plants are subjected to various abiotic stresses, resulting in loss of yield or decreased productivity. Plants have different adaptive and protective strategies at morphological, physiological and molecular levels to cope with environmental stress conditions [21]. Although stress conditions negatively affect beet growth, yield, and quality, the beet cultivars are able to tolerate abiotic stress conditions such as salinity, drought, cold, heat, and heavy metals [18,22–28]. Sugar beets exhibit tolerance to cadmium (Cd) and are capable of accumulating heavy metals such as Cd and nickel (Ni) [27]. The improvement of beet varieties with better heat tolerance is also an important task due to climate change and global warming [29]. Therefore, we need breeding techniques and agronomic practices for better tolerance to biotic and abiotic stresses in beets [30]. Thus, cultivated beets and their wild ancestor are important genetic sources for crop breeding programs and studying abiotic stress tolerance [15,31]. In the present review, we summarize the morpho-physiological, biochemical, and molecular alterations in cultivated beets (*B. vulgaris* L.) under alkaline, cold, heat, heavy metal, and UV stresses.

#### **2. Responses of Cultivated Beets (***B. vulgaris* **L.) to Different Abiotic Stresses Including Alkaline, Temperature, Heavy Metal, and UV**

Although several studies report different responses of beet cultivars to environmental stresses, research articles and reviews mostly focus on salt and drought response mechanisms in beets [22,23,32–34]. However, a comprehensive review describing the responses of cultivated beets to several abiotic stress factors including cold, heat, alkaline, heavy metal, and UV is lacking. Therefore, this review focuses on the responses of cultivated beets (*B. vulgaris* L.) to alkaline, cold, heat, heavy metal, and UV stresses at morpho-physiological, biochemical, and molecular levels. In Table 1, we demonstrate the list of beet genes known for their involvement in response to alkaline, cold, and heavy metal stress.


**Table 1.** Beet genes known to be involved in response to alkaline, cold, and heavy metal stresses.

#### *2.1. Alkaline Stress*

Alkaline stress (high pH) is one of the abiotic constraints of plants, which co-exists with salt stress and elicits severe detrimental damages to global agricultural production [44]. Over 954 million hectares of land on the globe is affected by salinity [45]. Salt stress results

from a neutral salt such as NaCl. Although alkaline salt stress is a type of salt stress, it is caused by alkaline salts such as NaHCO3 and Na2CO3, which is shortly called alkaline stress and causes more damage than neutral salt [46,47]. Numerous research groups across the globe have been perusing tolerance mechanisms to understand the salt stress responses in various crops and model land plants [48,49]. However, the studies focused on high salinity together with alkaline stress are minuscule [44,46]. Apparently, when the plants simultaneously encounter high salinity and high pH, their cumulative damage is more severe than their single occurrence [44]. Several previous reports determined that sugar beet can sustain moderate exposure to saline and alkaline conditions [35,37,48]. However, only a few reports investigated the responses of beets under alkaline stress conditions [47,50]. Hence, to alleviate the alkaline stress-induced damages in commercially important crops such as beets, we need to build a comprehensive knowledge repository that helps devise better strategies for generating stress-tolerant cultivars to attain sustainable agriculture [35,51,52]. Furthermore, developing high salinity-resistant cultivars will efficiently and rationally utilize salinity-affected areas in cultivated lands [45].

Although alkaline stress and salt stress share many common features, such as osmotic stress and ion toxicity, the alkaline condition has unique differences to consider as a different stress form [51]. The alkaline stress includes three principle factors that negatively impact plant growth and development: high soil pH, Na+ toxicity, and water deficiency [51]. For example, it has been shown that alkaline stress-induced Na+ toxicity and oxidative stress decreased photosynthesis and growth in tomato plants. Moreover, alkaline stress led to higher Na+/K+ ratio and lower K<sup>+</sup> content in tomato seedlings [51], and the expression of genes encoding Na+ transporters such as *SlNHX1*, *SlNHX2*, *SlSOS1*, *SlHKT1,1*, and *SlHKT1,2* were found to increase in tomato roots exposed to NaHCO3 [52,53]. However, we still do not know how sugar beet plants maintain Na+-K+ homeostasis under alkaline stress conditions and whether Na+ transporters contribute to the alkaline stress response in beets. High alkaline pH causes the occurrence of oxidative stress through reactive oxygen species (ROS) and the production of malondialdehyde (MDA), which damage the membrane integrity and intracellular components in plants [47]. To decrease the ROSinduced oxidative stress, plants use several enzymatic and non-enzymatic antioxidants [54]. Enzymatic antioxidants including superoxide dismutase (SOD), catalase (CAT), peroxidase (POX) and ascorbate peroxidase (APX) are involved in scavenging of superoxide radicals and hydrogen peroxide (H2O2) [54–56]. Under salt stress, cultivated beets and wild beet show higher antioxidant enzyme activities [57–59]. Similarly, Zou et al. [30] reported that the alkaline stress-tolerant beet cultivar KWS0143 displayed higher antioxidant enzyme activities such as CAT and APX than the sensitive cultivar Beta464 under the same growth conditions [30]. This implies that the tolerant plants are bestowed with durable antioxidant defense equipped with APX, CAT and SOD enzymes to circumvent the cellular damages under salt-alkaline stress [30]. Hence, we need to identify genetic resources with a strong innate antioxidant defense system to fortify beet cultivars with alkaline stress tolerance. In addition to oxidative stress, soils with high pH perturb the macro and micronutrient balance in the soil, which drives the plant to a physiological depression [50]. Previously, Oster et al. [60] classified the alkaline stress into three categories based on the alkaline salt percentage in soil. According to this classification, the alkalinity is considered as mild (3% salt content and pH 7.1–8.5), moderate (3–6% salt and the pH is 8.5–9.5), and severe (>3–6% salt and the pH over 9.5) [60]. In contrast to the detrimental effects of alkaline stress, mild alkaline stress can help the plants to grow bigger and healthier [50,61]. Likewise, in a recent report, Geng et al. [50] examined the differential proteomic responses of sugar beet seedlings by treating them with pH 5, pH 7.5, and pH 9.5 (acidic, neutral, and alkaline) conditions. In the study, they found that the acidic pH caused more growth retardation and enzymatic aberrations than that of neutral and alkaline pH conditions [50]. In contrast to other reports, the alkaline conditions (pH 9.5) significantly improved plant height, fresh weight, total leaf and root area, net photosynthetic rate, stomatal conductance, intercellular CO2 concentration, and chlorophyll contents compared to neutral and acidic

soils [50]. Moreover, a few more reports found that mild alkaline stress caused better growth, leaf chlorophyll contents, photosynthetic index, and antioxidant activities in sugar beet seedlings [30,61]. Geng et al. [61] found that neutral salt (NaCl:Na2SO4, 1:1, Na+ 100 mM) remarkably decreased growth and photosynthesis when compared with mild neutral salt (NaCl:Na2SO4, 1:1, Na+ 25 mM) and alkaline conditions (Na2CO3, Na<sup>+</sup> 25 mM) in sugar beet plants. In contrast, plants displayed a significant increase in total biomass, leaf area, and photosynthesis under mild neutral salt and alkaline conditions [61]. Interestingly, sugar beet plant growth was not impacted by high alkaline salt (Na2CO3, Na+ 100 mM) as compared to control [61]. We speculate that by virtue of being tolerant to mild saline–alkaline stress, the sugar beet cultivars might display better growth, and we need further experimental evidence to learn the growth patterns of different beet cultivars altered under mild alkalinity. Nevertheless, the growth retardation of plants is found to be proportionately elevating along with the increase in alkaline stress severity [62]. Additionally, alkaline stress responses in plants are usually governed by a multigenic effect, but not by a single gene expression, which implies the intricate stress signaling mechanism [36,63,64].

Numerous reports suggest that under alkaline stress, several physiological parameters, including stomatal conductance (Gs), transpiration rate (Tr), relative water content (RWC), water use efficiency (WUE), accumulation of photosynthetic pigments, and the net photosynthetic rate (Pn), were dropped [47,62]. Specifically, the photosystem-II (PSII) quantum efficiency (Fv/Fm) ratios are negatively affected by alkaline stress, which reduce the electron transport rate [65]. Furthermore, high alkaline conditions dampen the leaf area (LA) and chlorophyll contents (Chl a and b), specifically Chl b, which lowers the photosynthetic rate and WUE [66]. All these physiological parameters will eventually curtail the seedling growth and seedling emergence under alkaline stress [30,62]. In another study, Liu et al. [66] assessed the physiological responses of white Swiss chard under saline and alkaline conditions. Their study identified that although Swiss chard retains higher RWC under alkaline stress, the seedlings suffered from alkaline stress in terms of plant growth. The growth retardation was likely caused by high pH, CO3 <sup>2</sup>−, and HCO3 – toxicity [66]. Additionally, the physiological indicators such as chlorophyll contents, WUE, and the ionic balance were also perturbed in Swiss chard under 50–100 mM alkaline stress [66]. While comparing the glycine betaine (GB) and proline levels, they found that the GB levels in sugar beet were lower in 50 mM alkaline stress than that of 50 mM salt stress, whereas they did not find any significant alterations in proline levels [66]. This bolsters the notion that the GB plays a more critical role in mediating the alkaline stress tolerance than proline for Swiss chard [66]. It is a well-known fact that compatible solutes including GB, proline, and soluble sugars are remarkably increased under salt stress conditions to maintain photosynthesis and stomatal conductance in beets [67–69].

In addition to physiological and biochemical responses of beets under alkaline stress, only few genes have been reported to be involved in alkaline stress response in beets. For example, Wu et al. [35] identified 58 putative *WRKY* genes in the sugar beet genome, and among them, nine genes were found to be responsive to the alkaline stress stimulus (~15 mM to 100 mM NaCHO3) in both root and shoot tissues [35]. In the study, they found augmented expression of the *BvWRKY10* gene in shoots and *BvWRKY16* expression in root tissues under alkaline stress [35]. The differential expression of *BvWRKY* genes in different tissues implies their functional roles in mediating the alkaline stress responses in different tissues and needs further experimental attention. The WRKY family of transcription factors is plant-specific and plays many critical roles in diverse aspects of plant physiological processes, including abiotic stress responses [70]. Through a transcriptomic approach, some of the differentially expressed genes (DEGs) were shown in alkaline stress-treated beets. Recently, Zou et al. [36] identified differential expression of 1270 genes in alkaline stress-tolerant cultivar KWS0143 in response to alkaline stress. They irrigated the plants with 75 mM alkaline solution (Na2CO3:NaHCO3, 1:2, pH 9.67) and harvested the leaf tissues three (short-term) and seven days (long-term) after the treatments [36]. Compared

to the control groups, the short-term and long-term treatments induced the expression of '*Ethylene-insensitive protein 2*' (*LOC104884677*) and '*Metal tolerance protein 11*' (*LOC104886952*) genes, respectively [36]. The results suggest that some of these DEGs would be useful for developing alkaline-tolerant beet cultivars. In another report, Zou et al. [47] assessed the roles of long non-coding RNAs (lncRNAs) in sugar beets under different alkaline stress conditions as previously described in Zou et al. [36] by high-throughput RNA sequencing [47]. In this study, they identified 93 differentially expressed alkaline stressresponsive IncRNAs. Furthermore, additional functional attribution of candidate target genes revealed their association with diverse biological processes, including kinase activity, ribosomal and ribonucleoprotein constituents, and protein metabolic activity, and denotes the association of specific target genes with lncRNAs [47]. In addition, Zou et al. [71] treated the sugar beet seedlings with an alkaline solution and performed small RNA sequencing [71]. They found 53 novel microRNAs (miRNAs) responsive to long-term and short-term alkaline stresses [71]. Similarly, the gene ontology (GO) analysis uncovered enrichment of miRNAs related to the "redox process" and they reported the involvement of '*polyphenol oxidase*' (LOC04900758) gene as the target of alkali-responsive miRNAs. In addition to this, the other 29 miRNAs responsive to long-term alkaline stress can be useful as potential targets to fortify crops with alkaline stress resistance. In Table 2, we summarize the alkaline stress responses in sugar beet varieties.



#### *2.2. Cold and Heat Stresses*

Because plants are sessile organisms, the ambient temperature has a profound impetus on their entire life cycle, reflecting on their spatial distribution and seasonal behaviors [72]. Their surrounding temperatures also influence the plant growth rate and development, and each plant system has its own set of minimum, optimum, and maximum range of temperatures for survival [73]. Crop production varies depending on the severity of temperatures [74]. Furthermore, plants differentially respond to cold or heat stress according to their developmental stage. Hence, to circumvent the yield damages associated with capricious climates, we need to accumulate the morpho-physiological responses for individual crop varieties. Furthermore, more studies should be performed in order to characterize stress-responsive genes and determine the molecular mechanisms under low and high-temperature stresses in beets, as we have limited knowledge on beet responses to temperature changes.

#### 2.2.1. Cold Stress

Low temperature is one of the most important constraints, impeding plant growth, distribution, biological activity, production, and, ultimately, economic yield [75]. The sensitivity and responses of sugar beet to cold temperatures depend on its developmental stage. Cold is known to drive several developmental events in sugar beet in early and later stages, such as germination, growth, bolting, and accumulation of molassigenic products in the roots [76]. In sugar beets, exposure to cold temperatures at the early seedling stages causes severe root growth retardation and reduced sugar yield [75,77]. Although cold temperatures (i.e.,−2 ◦C) result in loss of cotyledon viability, the seedlings at 3–4 leaf stage can withstand freezing temperatures up to −10 ◦C [78,79]. Furthermore, sugar beet roots and shoots show differential responses to cold stress. For instance, in three sugar beet genotypes (GT1, GT2, and GT3), cold temperatures impacted taproot growth more than the shoot growth [80]. It has been reported that there are variations in cold stress tolerance and sensitivity among *Beta* germplasms [81]. Hence, to generate cold-tolerant varieties in commercially essential crops such as beets, knowledge pertaining to their responses to cold conditions is the most important prerequisite [82]. In some geographical sections, sugar beet seeds are sown in early autumn to expose them to shallow winter temperatures (below 0 ◦C). This practice helps protect the sugar beets from pathogen *Cercospora* attacks and drought stress [75]. Such an early seed sowing in fall, also known as "autumn sowing", was reported to produce sugar beets with better field emergence than the spring-sown beets [76]. Nevertheless, prolonged exposure of sugar beets at the young seedling stage to extreme cold temperatures seriously limits the yield [75]. Cold-treated sugar beet plants displayed a decrease in photosynthetic efficiency, quantum yield of PSII, leaf CO2 concentration, CO2 assimilation rate, and leaf transpiration rate [40,80]. Moreover, compatible solutes such as glucose, fructose, and raffinose in leaves were increased by 0 ◦C and 4 ◦C cold treatments [40,80], but decreased in taproots in response to freezing temperature [40]. Consistently, under freezing conditions, the sucrose content decreased in roots, followed by leakage of the root sap due to cell alteration in membrane permeability and infection with microbes. Water infiltration due to rapid freezing/thawing can also lead to softening of the root tissue and gradual rotting [83]. Rodrigues et al. [80] reported an interesting finding for the first time. Vernalization (long-term cold treatment at 4–15 ◦C) leads to a reversal of phloem translocation from taproots (sink tissue) to shoots (source tissue). Redirection of sugar flux is required for induction of flowering in sugar beet. This process might be the reason for the sugar beet sensitivity to freezing temperatures [80]. In a very recent work, three sugar beet genotypes (GT1, GT2, and GT3) were evaluated for freezing tolerance. Freezing temperatures caused the production of ROS, raffinose accumulation, and transcription of genes involved in raffinose metabolism in leaves and taproots [40]. These results suggest that raffinose metabolism has a protective role against freezing injury in sugar beet. Moreover, ROS-scavenging enzymes including SOD and CAT significantly enhanced in response to 4 ◦C [40]. Consistently, the maximum expression levels of genes

encoding antioxidant enzymes such as CAT, APX, ascorbate reductase, and glutathione peroxidase (GPX) were seen at 4 ◦C, but the expression was reduced at 0 ◦C. The findings indicate the temperature-dependent ROS production in sugar beet plants.

To date, very few sugar beet genes that function in cold stress response have been functionally characterized under cold stress conditions. In some reports, the transcript levels of genes involved in photosynthesis and compatible solute biosynthesis were investigated in cold-treated beets. For example, Rodrigues et al. [80] reported a sharp increase in the expression of photosynthesis-related genes encoding rubisco activase, rubisco small subunit, a chlorophyll a/b binding protein, and plastocyanin under cold stress. Kito et al. [39] isolated and characterized two sugar beet genes, *B. vulgaris RS1* and *RS2* (*BvRS1* and *BvRS2*), encoding raffinose synthase, which is involved in raffinose biosynthesis. The transcript levels of *BvRS1* and *BvRS2* genes were induced by cold stress in sugar beet leaves and roots [39]. Similarly, in a very recent study, the transcript abundances of galactinol synthase encoding genes, *GOLS2* and *GOLS3*, and two *RS* genes, *BvRS2* and *BvRS5*, were increased by freezing temperature [40]. Surprisingly, the expression of *BvRS5* gene and raffinose amounts remarkably induced in the taproots of freezing-tolerant beet cultivars, GT2 and GT3, but not in the sensitive one, GT1. As compared to other beet genotypes, the GT2 showed the maximum expression levels of *GOLS* and *RS* genes and raffinose levels in taproots, indicating the highest freezing tolerance in GT2 [40]. These findings suggest that the survival of taproot tissue under cold stress might depend on the accumulation of raffinose. As compatible solutes and antioxidants, raffinose family oligosaccharides have important roles in plant response to abiotic stress and stabilizing membranes and proteins [84,85]. In addition to genes involved in raffinose metabolism, the *B. vulgaris Integral Membrane Protein (BvIMP)* gene is the closest homolog of *A. thaliana early response to dehydration-like 6 (AtERDL6)*, which was previously reported for its cold stress-responsive function [86]. Cold stress may lead to elevations in the transcription of *BvIMP* gene and vacuolar sugar trafficking in sugar beet leaves, which is critical for cold stress response and seed germination [37]. Ectopic overexpression of *BvIMP* in *Arabidopsis* resulted in altered glucose concentration during cold conditions, lower accumulation of monosaccharides, and cold-sensitive phenotype compared to the wild-type [37]. In a recent study, Porcel et al. [38] uncovered and isolated a novel endoplasmic reticulum-located aquaporin gene, *B. vulgaris COLD1 (BvCOLD1)*, which is specific to the *Chenopodiaceae* subfamily. The *BvCOLD1* gene is ubiquitously expressed in all tissues of sugar beet [38]; however, its expression was not changed by cold stress [38,75]. In contrast to the wild-type plants, overexpression of *BvCOLD1* restored the membrane fluidity in transgenic *Arabidopsis* lines under cold temperatures and rendered tolerance to cold stress, suggesting that it could be a useful gene for developing biotechnological strategies in order to generate cold-tolerant beet cultivars [38].

#### 2.2.2. Heat Stress

Elevated temperatures and water deficit conditions tend to elicit similar impacts on plant water content where the evaporation exceeds the water intake, eventually leading to the plant wilting [87]. Across the globe, we face rapid climate changes and adverse weather problems; hence, developing heat-tolerant crops is the need of the hour. High temperatures impede many vital developmental events such as seed germination and impact seed vigor and viability and seedling emergence, and eventually challenge their survival [88,89]. Critical physiological processes, including photosynthesis and PSII activity, were also affected due to electron transport chain block under heat stress [90,91]. Of late, sugar beet cultivation is also expanding to the tropical and sub-tropical areas, and more people pay attention to cultivation of the sugar beets in summer [29,92]. Ironically, there are few studies aimed to select the heat-tolerant sugar beet cultivars. For the identification of the heat-tolerant beet genotypes, currently, there are no universally approved criteria. Different research groups used different parameters to evaluate the heat stress tolerance in different beet cultivars. For instance, Malmir et al. [92] considered the seed vigor index

and root length as evaluation parameters of heat stress tolerance in the early growth stage [92]. To investigate the effects of heat on early growth in sugar beet, they compared 31 sugar beet genotypes under heat stress conditions. Among all the variants tested, the tolerant genotype displayed relatively higher germination, seed vigor, plumule length, and seedling length compared to other genotypes, suggesting that the tolerant one is a prospective cultivar to expand the sugar beet cultivation to tropical areas [92]. Under high temperatures, the leaf temperature, which is associated with vapor pressure deficit (VPD) and stomatal conductance, is known to be enhanced [93]. Moreover, another recent study showed the stress tolerance index (STI) and average root and recoverable sugar yields as selection parameters to identify heat-tolerant lines among 18 sugar beet breeding lines [29]. Among them, six lines were found to have the highest yield, and two lines can sustain under heat stress [29]. In a previous work, two fodder beet cultivars (Ecdogelb and Ecdorot) were used to reveal the impacts of different light intensities and temperatures on fodder beet physiology [94]. High temperature affected root weight ratio (RWR), dry leaf weight (DLW), dry root weight (DRW), total dry weight (TDW), specific leaf area (SLA), net assimilation rate (NAR), and relative growth rate (RGR) in both cultivars at low light intensity [94]. For example, under high light intensity and temperature (20 ◦C), the cultivar Ecdorot exhibited enhancements of leaf weight ratio (LWR). The highest RGR, RWR, and DLW levels were recorded in response to high temperature and low light intensity in both cultivars. High temperatures result in increments of the growth in root crops, but adversely impact the final biomass [95]. When the temperature was increased from 14 ◦C to 19.6 ◦C, an increase in the SLA was also observed [94]. Leaf area, which is used as a selection parameter of drought-tolerant beet cultivars, determines the plant growth rate during initial phase of development [96] and is associated with root and sugar yield [32]. Thus, we assume that the leaf area could be an important parameter to enhance sucrose yield of beets under high temperature conditions.

Unfortunately, so far, no beet genes have been functionally characterized under high temperature conditions. Moreover, the knowledge on beet physiological and biochemical responses is very limited. Hence, comprehensive studies should be performed in different beet cultivars under heat conditions to gain a better understanding of heat tolerance mechanisms in beets at different developmental stages. In Table 3, we summarize the low and high temperature stress responses in cultivated beets.

#### *2.3. Heavy Metal Stress*

Generally, heavy metals are a group of metals and metalloids with atomic density more than 5 g cm<sup>−</sup>3, or five times or more, greater than water [97], including lead (Pb), cadmium (Cd), nickel (Ni), cobalt (Co), iron (Fe), zinc (Zn), chromium (Cr), arsenic (As), silver (Ag), and the platinum group elements. Mining and smelting operations and agriculture have caused heavy metal contamination of soils with Cd, copper (Cu), and Zn in many areas of the world [98]. Moreover, due to vigorous mining and industrial activities, the metal pollution in soils is becoming prevalent day by day and posing a severe threat to ecological balance [99,100]. For example, in 2002, 22,000 t of Cd, 93,900 t of Cu, 783,000 t of Pb, and 1,350,000 t of Zn were released into the environment on the global scale [101,102]. The buildup of heavy metals in arable lands results in contamination of soils, making them unsuitable for cultivation of plants, including beets. Therefore, the need for collecting scientific information regarding effects of various heavy metals on plants, response mechanisms of plants to heavy metal stress, and agronomic management of this stress can not be overemphasized.


#### **Table 3.** Cold and heat stress responses in cultivated beets.

Exposure of plants to toxic levels of heavy metals causes various metabolic and physiological alterations depending on the metal of concern, level of stress, plant species, cultivar, and other biotic and abiotic factors [103–105]. Most of the mineral ions such as Zn, Ni, manganese (Mn), etc., are required for all metabolic activities in plants at miniscule amounts. However, if the metal ion presence exceeds the threshold, they tend to exert detrimental effects on plant metabolism, resulting in leaf chlorosis, necrosis, turgor loss, a decrease in the rate of seed germination, and a crippled photosynthetic apparatus, which

could cause plant death [106–108]. Among the heavy metal ions, Cd, Zn, and Cu are reported as the most toxic metals, with serious health hazards to humans when they infiltrate the food chain [109]. Like other plants, heavy metals adversely affect the sugar beet as they proscribe various metabolic activities [27,110,111]. For example, heavy metals such as Pb damage the vacuolar membrane in red beet taproots [112]. Lead is one of the most toxic metals for plant cells, and it negatively affects plant growth, photosynthesis, respiration, and membrane transport [113]. Cd treatment in *B. vulgaris* caused growth retardation, leaf chlorosis, and increased root/whole plant ratio [114] with decreased roottip respiration and photosynthesis [110,114]. As compared to control plants, Cd-treated plants exhibited lower shoot dry weights, photosynthetic pigments, and reduction in water content of shoots and fine roots, dramatically [114]. Direct application of Cd on isolated leaves, protoplasts, and chloroplasts inhibited CO2 fixation without affecting the PSI or PSII and dark respiration rate, whereas indirect Cd application through the culture medium decreased the maximal quantum yield of CO2 assimilation [110]. Papazoglou and Fernando [27] tested the growth and heavy metal tolerance of sugar beet plants in Cdand Ni-contaminated soil [27]. They found that the highest Ni concentration (20 g) was lethal to the plants, and an interesting fact they found was that the single application of Ni caused higher toxic effects than the combination of Ni and Cd [27]. Nevertheless, the combination of Cd (5 g) and Ni (10 g) treatment resulted in a drastic reduction in fresh and dry biomass of aerial parts and beets, and a decrease in plant height [27]. Very recently, Haque et al. [43] found that toxic levels of Cd cause growth retardation of sugar beet plants because of low iron levels resulting in photosynthetic inefficiency, and cellular oxidative stress [43]. Cd-treated plants displayed sensitivity to oxidative stress, leading to an increase in levels of O2 − and H2O2 in roots and shoots. In addition, Haque et al. [43] examined the antioxidant defense system in sugar beet under heavy metal stress and found that Cd stress caused an enhancement of CAT enzyme activity in the shoots, whereas the activities of other antioxidant enzymes such as SOD, APX, and GR did not change in neither roots nor shoots. Furthermore, the results from a previous study indicated reduced uptake of N, P, Mg, K, Mn, Cu, and Zn upon Cd toxicity [114]. Similar to Cd stress, Zn toxicity decreased macronutrient concentrations (N, K, and Mg), whereas it enhanced the P level in shoots as well as roots [115]. In sugar beets, Cu and Zn treatments also significantly reduced plant growth, shoot and root lengths, and dry weight [116]. At high Cu concentrations, the shoots showed turgor loss, but lower Cu concentration did not affect plant growth [116]. Sagardoy et al. [115] reported that the toxic level of Zn reduced water content, leaf numbers, and root/shoot ratio, along with wrinkled and chlorotic leaves in sugar beet [115]. Root proteome analysis of sugar beet showed slight changes in metabolism under low and mild Zn levels, but higher levels of Zn led to cell death and cessation of metabolism through decreasing aerobic respiration and damaging defense systems required for oxidative stress response. Thus, the results showed that toxic Zn levels caused damages to the oxidative stress defense mechanisms due to Zn competition with divalent cations such as Fe, which might strengthen the symptoms of Zn toxicity in plants [117]. In summary, the results denote that the degree of toxicity of heavy metals on plant metabolism depends on plant species, the duration of stress, and type and concentration of heavy metals they were exposed to [111].

Several studies highlighted foliar uptake of heavy metals and their effects on the membrane permeability through the cuticle and percentage of open stomata in sugar beet [118,119]. A previous study demonstrated that sugar beet seedlings grown in nutrient solution containing high concentrations of CdCl2 showed an increased leaf transpiration rate and a decreased stomatal aperture area. Thus, higher Cd concentrations affected the permeability of the leaf cuticle [119]. Apart from seedlings, Cd stress was also shown to negatively influence sugar beet taproot growth. For instance, long-term Cd exposure caused decreased sucrose uptake and diminished dry weight in taproots, but the direct addition of Cd2+ to the medium enhanced the sucrose uptake at the tonoplast [120]. Increased accumulation of Cd lowered the contents of glucose, fructose, and sucrose in both shoots and roots of sugar beet [121] and inhibited the activity of plasma membrane H+-ATPase (PM H+-ATPase) [122]. Additionally, in several studies, changes in the activity of enzymes related to metal homeostasis and nitrate metabolism were investigated in heavy metaltreated sugar beets. For instance, the activity of ferric chelate reductase (FCR) involved in iron homeostasis was decreased under short-term exposure of Pb and Cd, but prolonged exposure increased the FCR activity in sugar beet roots [123]. Recently, Haque et al. [43] reported that the reduction in FCR activity and expression of *iron-regulated transporter 1 (BvIRT1)* gene suggested a negative impact of Cd in Fe acquisition. In another study, the Pb-treated sugar beet plants exhibited altered Cu deficiency levels and increased FCR activities [114]. When sugar beet plants were exposed to the highest concentrations of heavy metals (Ni and Cd), the nitrate content and nitrate reductase (NR) activity dramatically dropped in the leaves [111].

To cope with heavy metal stress, plants have developed certain strategies involving two type of mechanisms, i.e., avoidance and tolerance [124]. The avoidance mechanisms emphasize on limiting the uptake of heavy metals (e.g., Cd) into the plant, whereas tolerance refers to storing (e.g., in vacuoles) and accumulation of heavy metals by binding it to peptides, amino acids, and proteins [125,126]. To limit uptake of heavy metals and detoxify them, plants have developed certain mechanisms, including the development of morphological structures such as thick cuticle and cell walls, mycorrhizal symbiosis, and biologically active tissues such as trichomes [127–129]. Sugar beet, like canola, is a non-mycorrhizal plant species, and therefore has a limited ability to phytostabilize heavy metals and has been suggested as a source of phytoremediation of heavy metals [130,131] despite the negative effects of heavy metals on beet growth, physiology, and metabolism. For instance, among different crop plants tested, red beets have the capacity of removing Cd from soils [130]. It has been reported that sugar beet plants have the ability to accumulate Ni, Pb, and Cd [27,132]. Papazoglou and Fernando [27] suggested that sugar beet could be a suitable crop for phytoextraction of Cd as it can accumulate Cd and produce biomass. Similarly, Yadav et al. [132] compared several crops for their capacity to accumulate heavy metals and found that sugar beets accumulated the highest amount of Cd and Pb among the studied crops. These findings clearly suggest that sugar beet could be an efficient source for phytoremediation of heavy metal-contaminated soils. Since heavy metals such as Cd and Pb have serious effects on human and animal health, sugar beets grown on heavy metal-contaminated soils must not be used for food and feed purpose, but only for industrial purposes such as bioethanol production. Due to the hazardous nature of heavy metals, heavy metal-contaminated areas are of limited use, and removal strategies of excessive heavy metals from soils are required [133]. Phytoremediation is a promising approach to dampen the toxic effects of heavy metal pollution by utilizing the artificial hyperaccumulators. Transgenic plants, which can take up the persistent heavy metals, serve as artificial hyperaccumulators. For instance, Liu et al. [131] found an important role of glutathione (synthesized by *γ*-glutamylcysteine synthetase-glutathione synthetase) in cellular tolerance of heavy metal stress. Overexpression of *γ-glutamylcysteine synthetase-glutathione synthetase (StGCS-GS)* gene from *Streptococcus thermophilus* in sugar beet plants showed the explicit role of *StGCS-GS* in enhancing Cd, Zn, and Cu tolerance and accumulation of these metals in shoots of transgenic sugar beets [131]. Transgenic lines also displayed resistance to different heavy metal combinations, i.e., 50 μM Cd-Zn, Cd-Cu, Zn-Cu, and Cd-Zn-Cu, and had higher levels of glutathione (GSH) and phytochelatin (PC) compared to the WT [131]. Moreover, a study by Dronnet et al. [134] concluded that the sugar beet pulp is economical and highly selective in binding of divalent metal cations such as Cd2+, Cu2+, Ni2+, Pb2+ and Zn2+; thus, it could be useful as a substrate to entrap heavy metals in aqueous solution. Surprisingly, it was reported that the intake of juice extracted from red beet roots protects the chickens from Cd-induced oxidative stress with enhanced immune power [135]. However, it is unfortunate that the response mechanisms of cultivated beets and wild beet to heavy metal stress is yet to be investigated in detail. Further comprehensive studies are necessary to examine the influences of heavy metal contamination on

different beet cultivars, and yield and quality of bioethanol [27]. In addition, only few genes have been reported for their involvement in heavy metal response in beets. For instance, two MTP genes, *BmMTP10* and *BmMTP11* encoding metal-tolerant proteins from wild beet (*B. maritima*), were found to render tolerance to high concentrations of Mn2+ when expressed in yeast cells. Transcript level of *BmMTP10* gene was augmented by the presence of excessive Mn2+, but *BmMTP11* transcription was not altered, suggesting that BmMTP10 and BmMTP11 proteins have non-redundant functions in Mn detoxification [41]. Thus, the study demonstrated that the BmMTP10 protein, which is localized to the Golgi apparatus, is specific to Mn2+ transport and decreased Mn2+ levels in yeast cells [42]. Ni detoxification was regulated by a couple of genes in *B. maritima* named as toxic nickel concentration (NIC), i.e., *NIC3*, *NIC6*, and *NIC8* [42]. It was speculated that all three genes are involved in tolerance to Ni toxicity. Yeast cells expressing a cDNA clone (NIC6) from *B. maritima* showed substantially high tolerance to Ni but not to the other heavy metals such as Co, Cd, and Zn [42]. Even though the excess Ni accumulation is toxic to plants, *B. maritima* plants overcome the Ni-induced toxicity by internal sequestration, but not by effluxing Ni [42]. In a very recent study, under Cd stress, sugar beet roots displayed higher levels of putative *inactive Cd/Zn-transporting ATPase (BvHMA3)* and *natural resistance-associated macrophage protein 3* (*BvNRAMP3*) gene expression, suggesting that these genes might participate in Cd uptake [43]. Interestingly, in response to Cd application, no significant changes have been observed in the expression of *phytochelatin 3 (BvPC3)* gene encoding PCs [43], which are involved in the detoxification of Cd [136].

Further studies on sugar beet are needed to investigate the physiological, cellular, and molecular alterations induced by heavy metals to help plant biologists develop breeding strategies to improve sugar beet cultivars with efficient phytoremediation ability and ability to grow in heavy metal stress-affected fields [43]. In Table 4, we summarize the heavy metal stress responses in beets.


**Table 4.** Heavy metal stress responses in beets.


**Table 4.** *Cont*.

#### *2.4. Ultraviolet (UV) Stress*

Ultraviolet radiation (UV) causes various changes in metabolic activities of plants, imposing malfunctions and retarded overall growth. The key processes in plants affected by UV radiation include photosynthesis, biomass, respiration, transpiration, etc. UV-B (280–320 nm) radiation becomes a serious threat to the organisms because of the reduction in stratospheric ozone [137]. The stress triggers changes at molecular level by protein degradation, altering the double helical structure of DNA and antioxidant contents, etc. However, under UV stress conditions, plants adopt defensive tolerant mechanisms [138,139].

We have very limited information about the physiological and biochemical responses of beets to UV stress. Moreover, there are no reports on the molecular mechanisms and genes involved in UV stress response in beets. A report by Panagopoulos et al. [140] demonstrated that the leaves of sugar beets curled inwards and positioned towards light source with 68% growth reduction over control (ROC) under yellow light, whereas the plants were dead under a combination of yellow light and UV-B after three weeks [140]. They found that some parameters such as leaf area, fresh and dry weights, and total chlorophyll levels in sugar beet were decreased under UV radiation [140]. On the other hand, carotenoid concentrations showed different patterns upon imposition of UV radiation. For example, yellow light and a combination of white light + UV-B resulted in higher carotenoid contents, suggesting the protective role of these pigments against photo-oxidation [140]. The study also showed an increase in leaf peroxidase activity under the combination of white light and UV-B [140]. The increased peroxidase activity and ultraweak luminescence upon UV-B exposure and ascorbic acid incubated leaves represents a strong correlation in *Hibiscus* leaves [141] and sugar beet [140]. In a recent study, the most widely cultivated Iranian sugar beet variety, BR1, was used to analyze biochemical and physiological responses against different doses (3.042, 6.084, and 9.126 kJm−2d <sup>−</sup>1) of UV-B radiation [142]. The UV-B-treated sugar beet plants showed a drastic growth retardation with reduction in fresh weight, dry weight, and height. Moreover, total chlorophyll and carotenoid contents and photochemical efficiency of PSII were reduced in UV-treated plants. Interestingly, no significant raise in the proline levels was noticed. Betalain levels increased by 8%, 28%, and 34% with increased UV-B radiation of 3.042, 6.084, and 9.126 kJm−2d <sup>−</sup>1, respectively, indicating that these water-soluble pigments possess tolerant metabolic function in sugar beet varieties against UV-B radiation. Hence, it is likely that the BR1 variety is a suitable plant material for areas with UV-B irradiation [142].

Levall and Bornman [143] showed the establishment of a reproducible regeneration technique in sugar beet, wherein production of somaclonal variations was observed and UV-B-tolerant plants were selected. After additional UV-B treatment, unselected somaclones displayed significantly higher UV damage and lower carotenoid levels than the selected plants [143]. The UV irradiation exposure in in vitro conditions exhibited more tolerant callus parts than the protoplasts, paving the way for the selection of UV-tolerant sugar beet somaclones [143]. In another study, Levall and Bornman [137] showed differences between *Cercospora*-sensitive and -tolerant sugar beet plants upon the combined biotic (*Cercospora* fungal infection) and abiotic (UV radiation) stresses. The line tolerant to fungal infection was shown to be tolerant to UV-B alone and combined UV-B and biotic stresses; however, the photosynthetic yield significantly reduced in the sensitive line [137]. A report by Bornman et al. [144] showed that the UV-B radiation was not capable of penetrating organelles such as chloroplasts, resulting in intact thylakoids [144]. On the other hand, the ultrastructural image of sugar beet leaves showed prominent damages due to UV-B radiation (290–320 nm), whereas UV-C (254 nm)-treated sugar beet plants showed fewer structural changes, leading to a higher quantity of starch in chloroplasts, grana stacks fused to each other, and decreased damage to the leaf surface [145].

The results described above suggest that beet plants are adversely affected by UV stress conditions at the morpho-physiological level. However, molecular mechanisms and UV stress-responsive genes in beets are still elusive. Further studies are needed to better understand the UV stress response mechanisms at the morpho-physiological, biochemical, and molecular levels in different beet cultivars. In Table 5, we summarize the responses of cultivated beets to UV radiation.


**Table 5.** UV stress responses in cultivated beets.

#### **3. Concluding Remarks**

As an economically important crop plant, cultivated beets have multifarious industrial applications ranging from food and nutrition to sugar and bioethanol production. Despite beet tolerance to different abiotic stresses [16,24], the cultivation of beets is often challenged by various adverse environmental factors [34]. These climatic abnormalities are anticipated to be more aggravated due to human industrial activities as well as global warming effects. Hence, to meet the global food security demands, developing stress-resilient plant genotypes is one of the most important topics for crop production in stress-affected fields. However, selection of the suitable beet genotypes tolerant to environmental conditions is an arduous task for plant breeders [29] as there is no clear and comprehensive understanding about the stress signaling pathways and tolerance mechanisms in different climatic regions. Even though our understanding of the heavy metal accumulation ability of beets is limited, sugar beet plants have been suggested as a candidate for phytoremediation [28,126,135]. Sugar beets grown in contaminated soils pose a serious threat to human and animal health. Therefore, use of sugar beets grown for phytoremediation must be limited to industrial purposes, such as bioethanol production. Furthermore, we have limited experimental data showing the molecular mechanisms underlying the stress response of *B. vulgaris* genotypes under extreme temperatures (cold and heat), UV radiation, high pH, and heavy metals. Although the beet cultivars show some degree of stress resistance, persistent exposure to these abiotic constraints takes a toll of their development and growth potential. On the other hand, the wild beet (*B. maritima*) displays better stress tolerance compared to the modern beet cultivars as it is rich in allelic diversity [18,34]. Most likely, the modern cultivars lost some of their stress tolerance traits during progressive domestication. While utilizing the genetic variability in wild beet and stress-tolerant beets, we can ameliorate the allelic diversity, which further eases the improvement of tolerant varieties.

Since several beet cultivars were introduced and acclimated to tropical and sub-tropical climates, it would be thus essential to establish the pan-genomic studies of beet cultivars to uncover the precise genetic modifications responsible for the ecological adaptations. Establishing the phenotypic and genotypic diversity of various beet cultivars grown in

different climatic zones by utilizing the modern bioinformatic advents can enable us to generate stress-resistant crops. Consequently, further investigations are necessary to design breeding strategies under abiotic stress, and compare stress response mechanisms and signaling pathways between cultivated beets and wild beet. In Figure 1, we summarize morpho-physiological, biochemical, and molecular changes in beets under different abiotic stresses including alkaline, cold, heat, heavy metals, and UV radiation.

**Figure 1.** Schematic representation of morpho-physiological, biochemical, and molecular alterations in beets under alkaline, cold, heat, heavy metal, and UV conditions. This figure was created via BioRender.com (accessed on 12 December 2021).

**Author Contributions:** Conceptualization, S.Y.; writing—original draft preparation, S.Y., H.A., P.G. and M.A.; writing—review and editing, S.Y. and M.K.; preparation of Figure 1 and Tables, S.Y.; funding assistance, K.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no competing interests.

#### **References**


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