The protein coding sequence (CDS) of an orthologs GPCR gene from rice (LOC_Os04g51180.1) was retrieved and used to perform a Basic Local Alignment Tool (BLAST) search (E-value<1e-05) against the sugarcane expressed sequence tag (SUCEST) database (Vettore et al., 2003). SUCEST is a comprehensive collection of sugarcane RNA-sequencing (RNA-seq) transcript assemblies, with 237,954 high-quality ESTs prepared from 26 diverse tissue-specific cDNA libraries from 13 commercial sugarcane varieties (Vettore et al., 2001). Additionally, we mined publicly available sugarcane RNA-seq datasets through the National Center for Biotechnology Information (NCBI) Short Read Archive BLAST tool (Misra et al., 2007). We used all recovered reads (NCBI-SRA) and transcript hits (SUCEST) to assemble a consensus in silico sequence sharing highest similarity to the rice GPCR. The consensus transcript assembly defined a CDS of 1,407 nucleotides and encoded a GPCR-like protein. During the preparation stages of this manuscript, an allele-defined sugarcane draft genome was released (Garsmeur et al., 2018; Zhang et al., 2018). We compared our predicted ShGPCR1 sequence to the draft sugarcane genome, and found a single, perfectly matching locus.

We isolated total RNA from 100-mg sugarcane (variety CP72-1210) and energy cane (variety TCP10-4928) leaves using the Direct-zol RNA MicroPrep kit (Zymo Research, Irvine, CA, United States). We synthesized first-strand cDNAs for reverse transcription PCR (RT-PCR) using 1-μg total RNA and SuperScript III Reverse Transcriptase (Thermo Fisher Scientific, Waltham, MA, United States) with oligo(dT)₂₀ primer according to the manufacturer’s instructions. We amplified the GPCR-like sequence ShGPCR1 from sugarcane and energy cane using primers that recognize the start and stop codons of the gene, ShGPCR1-F (5'-GCGAGGAATACAGCAAGGGA-3') and ShGPCR1-R (5'-TGGGTCACCAAAGAAACATC-3'). We performed PCR reactions on a ProFlex™ PCR System (Applied Biosystems by Life Technologies, Carlsbad, CA, United States) in a total reaction volume of 50μl using 1μl of cDNA, 0.5μM of each target-specific primer, and 1.0U of Phusion DNA polymerase (New England BioLabs, Ipswich, MA, United States). PCR conditions were as: one denaturing cycle at 98°C for 30s, 30cycles each at 98°C for 15s, 62°C for 15s, and 72°C for 2min, and a final extension cycle at 72°C for 5min. We separated PCR amplicons from sugarcane and energy cane by electrophoresis on a 1% (w/v) agarose gel, before cloning into the pTEM73 vector and transformation into Escherichia coli strain DH5α. We isolated plasmid DNA from 10 randomly selected recombinant colonies using the Plasmid MiniPrep Kit (Zymo Research, Irvine, CA, United States) and by Sanger DNA-sequencing determined the identify of ShGPCR1 alleles present in sugarcane and energy cane (Table 1; Supplementary Dataset).

We used the DNA sequence of GPCR genes to deduce their amino acid sequences using the translate tool at the ExPaSy Bioinformatics Resource Portal.¹ We performed a homology search for the deduced amino acid sequences of ShGPCR1 from sugarcane and energy cane, using NCBI BLAST.² We then compared the amino acid sequence of the selected ShGPCR1 with GPCR-like proteins from Arabidopsis, rice, sorghum, maize, and cotton by multiple amino acid sequence alignment using ClustalW2.0.³ We performed phylogenetic analyses using the Neighbor-Joining method with pair-wise deletion of alignment gaps and Poisson correction for amino acid substitutions in MEGAX (Kumar et al., 2018). We used the ExPaSy PROSITE database of protein families and domains (Sigrist et al., 2013) to identify functional motifs and biologically significant sites in ShGPCR1. We predicted the presence of transmembrane domains using the Trans Membrane Hidden Markov Model 2 (TMHMM2) program (Krogh et al., 2001).

We generated embryogenic leaf rolls of sugarcane (variety CP72-1210) and subjected them to DNA bombardment as described above, with the following modifications. Embryogenic leaf rolls were cultured on MS0.6 medium (MS medium supplemented with 0.6mg/l of 2,4-D) for 8days in the dark at 28°C. Leaf roll disks (1mm diameter; 2–3 disks per bombardment) were preconditioned on MS0.6-osmoticum (MS6 with 0.2M D-mannitol and 0.2M D-sorbitol) for 4h before DNA particle bombardment. Gold particles were coated separately with plasmid DNA (5.0μg) of pTEM73-ShGPCR1-mGFP or pTEM73-mGFP. Twenty four hours after bombardment and incubation on MS0.6-osmoticum, leaf roll disks were transferred into MS3 medium and kept in the dark at 28°C for 10days prior to imaging. We rinsed the disks three times with water and sectioned them into 1mm thick pieces for microscopy; the pieces were stained with the cell membrane-specific lipophilic dye FM4-64 (Invitrogen, Thermo Fisher Scientific) for plasma membrane visualization (Lam et al., 2008), or with DAPI (Invitrogen, Thermo Fisher Scientific) for staining of nuclei. Stained pieces were rinsed three times with water and imaged on a Fluoview FV10i-LIV confocal laser scanning microscope (Olympus Life Sciences, Waltham, MA, United States) at 488, 546, and 385-nm excitation wavelengths for GFP, FM4-64, and DAPI, respectively. Images were acquired using the Z-stack scan mode (acquisition of images in different focus positions). Ten image planes were scanned with the optimal pixel size of 0.1μm.

The ShGPCR1 coding region (1,407bp) was PCR amplified from the cloned ShGPCR1 cDNA using gene-specific primers (Supplementary Table S1) and cloned at the BamHI and PmlI sites, replacing the custom synthesized codon-optimized (GenScript, Piscataway, NJ, United States) bialaphos-resistance (bar) gene into the minimal plant expression vector pTEM73 (Beyene et al., 2011) under control of the maize Ubi promoter (including first exon and intron) and the Cauliflower mosaic virus (CaMV) 35STTnos double terminator to generate pTEM73-ShGPCR1. The pTEM73-ShGPCR1 and pTEM73 (containing the bar selectable marker) were further used to perform biolistic-based co-transformation of sugarcane (Bower et al., 1996; Ramasamy et al., 2018). For subcellular localization of ShGPCR1, we PCR amplified the open reading frame of the mutant Green Fluorescent Protein mGFP (Haseloff et al., 1997) using high fidelity Phusion DNA polymerase (New England BioLabs, Ipswich, MA, United States). We cloned the mGFP PCR product by Gibson assembly (Gibson et al., 2009) into the pTEM73 vector downstream of the ShGPCR1 CDS, producing the pTEM73-ShGPCR1-mGFP construct, in which the chimeric gene is driven by the Ubi promoter and 35STTnos double terminator. Similarly, we cloned mGFP into pTEM73 to generate the control vector pTEM73-mGFP.

We collected tops of field-grown sugarcane (Saccharum spp. hybrids), commercial variety CP72-1210, during the growing season, and prepared leaf rolls for transformation, as previously described (Gao et al., 2013; Ramasamy et al., 2018). Briefly, we surface-sterilized immature leaf rolls close to the apical meristem in 70% (v/v) ethanol for 20min, sliced them transversely into 1mm thick sections, and cultured them on MS (Murashige and Skoog, 1962) supplemented with 3mg/l of 2,4-dichlorophenoxyacetic acid [2,4-D] (MS3 medium) for 30–35days at 28°C. Embryogenic calli were preconditioned on MS3-osmoticum (MS3 with 0.2M D-mannitol and 0.2M D-sorbitol) for 4h before and after DNA particle bombardment, which was performed according to Ramasamy et al. (2018). Briefly, for DNA particle coating, we added 5.0μg of each of pTEM73-ShGPCR1 and pTEM73 (bar selectable marker) plasmids sequentially to 5.0-μg gold particles (0.3μm, Crescent chemical Co, NY, United States) suspension using 1M-calcium chloride and 14-mM spermidine. Next, we placed 4μl of this DNA particle suspension (0.5μg DNA/bombardment) at the center of a syringe filter and delivered into tissue with a particle inflow gun using a 26-in Hg vacuum and 7-cm-target distances. We incubated bombarded embryogenic calli on MS3 medium for 10–12days in the dark for recovery. Shoot regeneration was performed under selection on MS medium with 1.5mg/l of benzylaminopurine and 3mg/l of bialaphos, followed by root initiation on MS medium with 4mg/l of indolebutyric acid and 3mg/l of bialaphos. After 6–8weeks, once root formation was well established, we transplanted transgenic seedlings into potting soil (Sunshine Mix #1; Sun Gro Horticulture, Belleview, WA, United States) and moved them to a controlled-environment greenhouse.

We verified the presence of the ShGPCR1 and bar (selectable marker) genes in co-transformed sugarcane plants by PCR using the forward primer (5'-GATGCTCACCCTGTTGTTTG-3') from the Ubi promoter and the reverse primer (5'-GACAGATCGAGCTCTGACTAGG-3') from the Tnos terminator. We performed PCR on a ProFlex™ PCR System in a total reaction volume of 25μl using 100ng of genomic DNA (isolated from 0.5–1g of young leaves of 3–4-month-old plants using the protocol of Chiong et al., 2017), 0.1μM of each target-specific primer, and 1.0U of Taq DNA polymerase and ThermoPol™ buffer (New England BioLabs, Ipswich, MA, United States). PCR conditions were as: one denaturing cycle at 95°C for 30s, 30cycles each at 95°C for 15s, 52°C for 15s, and 68°C for 1min, and a final extension cycle at 68°C for 5min. The amplified PCR products were ~1,640 and 500bp in size, respectively. Plants with the expected size amplicons were selected for further molecular analysis.

Integration of the ShGPCR1 expression cassette into the sugarcane genome was determined by Southern blot hybridization. We digested genomic DNA (10μg/reaction) overnight with HindIII, electrophoresed on 0.8% (w/v) agarose gels, and transferred into nylon membranes (Amersham Hybond-XL, GE Healthcare Bio-Sciences Corp., Piscataway, NJ) in 0.4-M sodium hydroxide (Koetsier et al., 1993). Pre-hybridization, hybridization, washing, and detection of DNA gel blots were performed, using Church’s buffer as described by Sambrook et al. (1989) and Mangwende et al. (2009). The ShGPCR1-specific probe (1,407bp) was released from pTEM73-ShGPCR1 with HindIII digest and labeled with [α-³²P]dCTP using the Random Primers DNA Labeling kit (Invitrogen, ThermoFisher Scientific, Waltham, MA, United States).

We propagated sugarcane seedlings in vitro on half-strength MS medium, transplanted them later to Sunshine Mix #1 (Sun Gro Horticulture, Agawam, MA, United States) in plastic pots, and moved them to a controlled-environment greenhouse (25–30°C during the day and 15–24°C at night; 1,200–1,600μmol/m²/s at midday). Plants were fertilized once a week with soluble Peters® Professional 20-20-20 (The Scotts Company, Marysville, OH; 1.6g/l). For stress-inducible expression of endogenous ShGPCR1 in sugarcane (variety CP72-1210), we moved 10-week-old seedlings grown in the greenhouse to a 28°C growth chamber for a 1-week acclimation period before subjecting them to cold, drought, or salinity. For cold treatment, seedlings were moved from a 28°C growth chamber to one maintained at 0°C. For drought stress, seedlings were carefully pulled out of potting medium and left to wilt on a tray for 2, 6, and 28h. For salinity treatment, we drenched the soil with 200-mM NaCl. For ABA treatment, detached leaves (4cm in length) were treated with 10, 25, 50, and 100nM of ABA for 10h. Samples were collected from each of the stress-treated and untreated control seedlings at 2, 6, and 28h post-treatment, flash-frozen in liquid nitrogen, and stored at −80°C. For ShGPCR1 expression analysis, we pooled equal amounts of total RNA extracted from tissue at each of the three time points.

We also analyzed the expression of ShGPCR1 in three ShGPCR1:OE lines under drought, salinity, and cold stresses (Begcy et al., 2012; Belintani et al., 2012; Da Silva, 2017). We performed drought stress-tolerance assays with 4-month-old ShGPCR1-OE and NT plants derived from tissue culture, acclimated in well-watered conditions in 15-L plastic pots for 1week in a controlled-environment greenhouse and later subjected to a progressive drought by withholding water for 40days until soil moisture reached ~10% for NT and~15% for transgenic plants. For salinity stress-tolerance assays, we watered 1-month-old ShGPCR1-OE and NT plants with 200-mM NaCl (200ml per 1-L pot) in reverse-osmosis water by soil drenching once a week for a period of 2weeks. For chilling stress-tolerance assays, we treated 2-month-old ShGPCR1:OE and NT plants grown in the greenhouse at 4°C for 3weeks, followed by −5°C for 4h in a temperature-controlled growth chamber. We also performed chilling treatments with 2-week-old seedlings in MS medium in Magenta boxes at 4°C for 24h, followed by −20°C for 4h. Foliar symptoms of stress were evaluated, and severity of symptoms was rated based on a scale of 1–3, where 1=mild (few leaf curling and wilting), 2=moderate (<25–50% of leaves showing curling and wilting with concomitant necrosis), and 3=severe (>75% of leaves showing leaf curling and wilting with concomitant necrosis). Leaf tissues of control and stressed ShGPCR1-OE and NT plants were harvested, flash-frozen in liquid nitrogen, and kept at −80°C for expression analysis of ShGPCR1 and stress-related marker genes (LEA, DHY, SCDR4, and GOLS for drought, ERF3, SOS1, and ShNHX1 for salinity, and SsNAC23, CBF2, and ScADH3 for cold; Supplementary Table S1). All stress experiments were carried out in a randomized block design using 3–4 biological replications, i.e., independent plants per line per treatment.

We evaluated the ShGPCR1-OE lines and NT plants for leaf relative water content (RWC) and agronomic parameters every 7days for a period of 40days of progressive drought stress. We measured RWC using the detached leaf method at the end of the drought stress treatment (Dhanda and Sethi, 1998). We cut fully expanded leaves (1cm×4cm), weighed them to obtain fresh weight (FW), floated them on de-ionized water in a Petri dish, and kept at 10°C for 4h. We then patted the resulting turgid leaves dry on filter paper to remove excess water and weighed them again to obtain their turgid weight. After weighing, we dried the leaf segments at 80°C in a hot air oven for 24h before measuring their dry weight. We calculated RWC using the formula: RWC (%)=[(FW−DW)/(TW−DW)]×100. The agronomic parameters considered and measured here were culm diameter (cm), culm height (cm), flag leaf length (cm), and dry root weight (g). We also estimated total biomass (aboveground parts, such as culms and leaves) by calculating the fresh biomass weight and DW after drying of samples at 80°C until they reached a constant weight, using an analytical balance. We calculated the total biomass using the formula: Total biomass (%)=(DW of total biomass/FW of total biomass)×100. Significant differences were determined using two-sample Student’s t-test (p<0.05).

To measure ShGPCR1 transcript levels in sugarcane, we isolated total RNA from 0.5g young leaves of stress-treated wild type and corresponding untreated controls, using the Direct-zol™ RNA MiniPrep kit (Zymo Research, Irvine, CA, United States). We synthesized first-strand cDNAs using 1-μg total RNA and the SuperScript™ IV Reverse transcription kit (Invitrogen, ThermoFisher Scientific). We performed quantitative RT-PCR (RT-qPCR) on a CFX384 Real-Time System (Bio-Rad Laboratories, Inc., Hercules, CA, United States) with the iTaq™ Universal SYBR Green Supermix (Bio-Rad Laboratories, Inc.) using 3–4 biological replicate samples and two technical replicates for qPCR analysis. Primers were designed with Primer 3.0.⁴ Results were analyzed and recorded as C_T (threshold cycle) values. We quantified each transcript relative to the sugarcane reference gene, ANTHRANILATE PHOSPHORIBOSYLTRANSFERASE (APRT, GenBank CA089592.1; Casu et al., 2012), using the comparative C_T method (2^−ΔΔC_T). Primer pairs used for RT-qPCR were as: ShGPCR1-F2 (5''-AAGTCCAGGCACTGGAAGAG-3'') and ShGPCR1-R2 (5''-AACACAATACACCGACAAAGCA-3''), and APRT2-F (5'-CGGTCGTTTCTGGTTTTGTT-3') and APRT2-R (5'-CGCCAAGAATGTGGTATGTG-3'). Significant differences were determined using two-sample Student’s t-test (p<0.05).

To examine the accumulation of ShGPCR1 transcripts in the sugarcane ShGPCR1-OE lines, we performed RT-PCR on a ProFlex PCR System in a total reaction volume of 25μl using 1μl of cDNA (synthesized from 1μg of total RNA extracted from leaves), 0.1μM of each target-specific primer, and 1.0U of Taq DNA polymerase and ThermoPol™ buffer (New England BioLabs). PCR conditions were as: one denaturing cycle at 95°C for 30s, 30cycles each at 95°C for 15s, 52°C for 15s, and 68°C for 1min, and a final extension cycle at 68°C for 5min. Primer pairs used in the RT-PCR analysis were as: ShGPCR1-F and ShGPCR1-R for ShGPCR1, and APRT2-F and APRT2-R for APRT2. Primer sequences for the sugarcane stress-responsive marker genes LEA, DHY, SCDR4, GOLS, ERF3, SOS1, ShNHX1, SsNAC23, CBF2, and ScADH3 used in the stress-tolerance assays of the ShGPCR1-OE lines are provided in Supplementary Table S1 (Nogueira et al., 2005; Iskandar et al., 2011; Reis et al., 2014; Su et al., 2014, 2020; Li et al., 2018; Devi et al., 2019; Theerawitaya et al., 2020; Brindha et al., 2021).

Ca²⁺ measurements of leaf cells were performed as previously described (Li et al., 2014) with the following minor modifications: transverse sections of (5–10mm) sugarcane ShGPCR1:OE and NT leaves were prepared using razor blades in Tyrode solutions (145-mM NaCl, 5-mM KCl, 2-mM CaCl₂, 1-mM MgCl₂, 10-mM Glucose, and 20-mM HEPES, pH 7.4) with 23μM of the Ca²⁺-sensing dye Fluo-4AM (Invitrogen™ Molecular Probes™, Eugene, OR, United States) and 2.5μl of power concentrate (100x; Invitrogen™ Molecular Probes™) for 1h at room temperature. Subsequently, leaf pieces were placed on a glass coverslip and visualized under an Olympus IX71 inverted microscope attached with PTI EasyRatioPro system (HORIBA Scientific, Piscataway, NJ, United States). Changes in fluorescence of single cells were recorded with EasyRatioPro v3.4 software (HORIBA Scientific) with an excitation wavelength of 488nm and an emission wavelength of 525nm. GTP (100mM) and ionomycin (1mM; 1–3μl) were added to the leaf samples during live measurements to test their effects on Ca²⁺ release. All Ca²⁺ imaging data were analyzed with EasyRatioPro (PTI, HORIBA Scientific) software and further processed with Excel (Microsoft, Redmond, WA, United States), Igor Pro v8.0 (Wavemetrics, Lake Oswego, OR, United States) software, and graphs were plotted with Origin Pro v2020 (Originlab, Northampton, MA, United States) software. The data were presented as means±SE (n=number of plant cells). For all analyses, data were pooled to attain a sample size of 33–99 plant cells. Significant differences were determined using two-sample Student’s t-test (p<0.05).

We used a combination of comparative genomics and bioinformatics tools to identify ShGPCR1. First, we used the DNA sequence of a GPCR orthologs gene from rice (LOC_Os04g51180.1) to perform a BLAST search (with cutoff E-value<1e-05) against the SUCEST (Vettore et al., 2003) and the National Center for Biotechnological Information (NCBI) short reads archive (Misra et al., 2007) for sugarcane sequences. We assembled a 1,407-bp sequence, hereafter called ShGPCR1 (Saccharum spp. hybrids GPCR1), in silico from the retrieved hits. We compared our identified CDS to the draft monoploid sugarcane genome (Garsmeur et al., 2018; Zhang et al., 2018) that was recently released. We identified a single and perfect match (E-value=0) to a transcript (Sh_221E20_t000010), thus validating our targeted comparative genomics pipeline, which we applied before the release of the draft genome.

Next, we amplified the endogenous CDS from both sugarcane and energy cane using RT-PCR with primers specific to the predicted CDS. Because sugarcane and energy cane are hybrid genomes with complex ploidy, it is important to understand haplotype divergence of the homologs alleles of the corresponding gene. Hence, we isolated and sequenced several clones representing homeologs alleles of ShGPCR1 (Table 1). We identified several alleles from the analysis of 10 independent ShGPCR1 clones isolated from sugarcane and energy cane (Table 1; Supplementary Dataset). In general, the ShGPCR1 alleles showed a high degree of conservation (~98% nucleotide identity) among homologs isolated from energy cane and sugarcane. These results are consistent with the high collinearity and conservation in gene structure and nucleotide sequence (~95.7% identity) among several other homologs genes observed in sugarcane, despite the complex polyploidy of the sugarcane genome (Garsmeur et al., 2011). Together, these results suggest that gene coding sequences are under purifying selection pressure in sugarcane and that homologs alleles may have all retained biologically relevant functions.

The predominant ShGPCR1 allele corresponding to the 1,407-bp cDNA encoded a full-length protein with a deduced protein sequence of 468 amino acids (Supplementary Dataset), a predicted molecular mass of 53.5kDa, and an isoelectric point of 8.8. The ShGPCR1 protein showed ~99% similarity to other GPCR-like proteins from sorghum (Sobic.006G203300.1) and maize (Zm2g129169_T01) and~96% similarity to the rice COLD1 GPCR protein (LOC_Os04g51180.1; Supplementary Figure S1). To assess the evolutionary relationship between ShGPCR1 and other plant GPCRs, we performed a phylogenetic analysis with selected GPCRs from monocots and dicots. In general, ShGPCR1 clustered with GPCRs from closely related monocots, such as sorghum (Sorghum bicolor), foxtail millet (Setaria italica), maize, rice, and Brachypodium (Brachypodium distachyon), but it was more distant from dicots, such as Arabidopsis, cabbage (Brassica oleracea), cotton (Gossypium raimondii), citrus (Citrus sinensis), and potato (Solanum tuberosum; Figure 1A).

One hallmark of GPCRs is their secondary structure, which consists of an N-terminal extracellular domain for membrane anchoring, 7–9 transmembrane (TM) helical-spanning domains connected by three extracellular N-terminal loops for ligand binding, three C-terminal loops in the cytosol for heterotrimeric G-protein binding, and an intracellular C-terminal tail for phosphorylation and desensitization (Wheatley et al., 2012; Ding et al., 2013). Amino acid sequence alignment of the ShGPCR1 protein to corresponding GPCRs from monocots and dicots showed a high degree of conservation (Supplementary Figure S1). We determined the presence of the TM region by TMHMM2 prediction (Krogh et al., 2001). ShGPCR1 possessed the nine TM helices linked by alternate intra- and extracellular loops; its N-terminal and C-terminal domains were also present on opposite sides of the membrane, as in other GPCRs (Ma et al., 2015; Figure 1B). Furthermore, we identified, by PROSITE motif analysis of ShGPCR1, a conserved Ras GTPase-activating protein domain and an ATP-/GTP-binding region, both important for GPCR function (Figure 1C). These domains were well characterized in the Arabidopsis GTG1, GTG2, and GCR2 proteins (Liu et al., 2007; Pandey et al., 2009; Yadav and Tuteja, 2011; Shi and Yang, 2015). The ShGPCR1 protein also contained six N-myristoylation sites (Figure 1C), known to be important for co-translational or post-translational modification of GPCRs and to help anchor the protein to the membrane (De Jonge et al., 2000; Utsumi et al., 2005). These myristoylation motifs were essential for regulating signal transduction (Sessa et al., 1993; De Jonge et al., 2000) and cellular responses to high salinity (Ishitani et al., 2000). GPCRs can be phosphorylated in response to ligand stimulation by GPCR kinases and protein kinases from a diverse range of kinase families, which determines specificity in signaling outcomes (Torrecilla et al., 2007). ShGPCR1 possessed five predicted protein kinase C and three casein kinase II phosphorylation sites (Figure 1C) that might be important for its biochemical regulation, and in relaying Ca²⁺-dependent signals (Torrecilla et al., 2007; Yadav and Tuteja, 2011). Together, these bioinformatics analyses uncover conserved and characteristic features of ShGPCR1.

To investigate the in planta function of ShGPCR1, we first determined whether ShGPCR1 transcript levels were altered in response to abiotic stress, as was reported for other plant GPCRs (Ma et al., 2015; Anunanthini et al., 2019). Quantitative RT-PCR (RT-qPCR) showed that ShGPCR1 transcript levels were significantly induced (p<0.05) upon exposure to drought and salinity stress: 5.2-fold in culms and 2.4-fold in leaves (drought), and 7.1-fold in culms and 2.2-fold in leaves (salinity), relative to control tissues (Figure 1D). Cold stress modestly enhanced ShGPCR1 expression in leaves and culms (~1.2–1.37-fold; Figure 1D) that was statistically significant (p<0.05) when compared to untreated tissues. ShGPCR1 expression was also significantly induced (p<0.05) by exogenous ABA treatment (Supplementary Figure S2), suggesting that ShGPCR1 responses may be ABA dependent. The increased ShGPCR1 transcript level in response to the various stresses may lead to changes in Ca²⁺ ion flux and reactive oxygen species (ROS; Sanders et al., 1999; Munns and Tester, 2008; Mohanta et al., 2018). These often act as secondary messengers to coordinate stress-mediated signal transduction with their cognate protein kinases for adaptation to adverse conditions. In addition to the upregulation of ShGPCR1, the induction of GPCR proteins, such as COLD1 from rice, maize, sorghum, and sweetcane (Erianthus arundinaceus) under drought, salt, or cold stress, underscores the regulatory role of GPCR proteins under abiotic stress (Ma et al., 2015; Anunanthini et al., 2019).

We next examined the subcellular localization of ShGPCR1. We cloned the full-length ShGPCR1 CDS in frame with the CDS of green fluorescent protein (mGFP) at the C-terminus. We then transiently delivered the construct into sugarcane by bombardment of embryogenic leaf rolls (8days old). Confocal microscopy of bombarded leaf rolls stained with the plasma membrane-specific dye FM4-64 showed that ShGPCR1::mGFP co-localized with FM4-64 primarily at the plasma membrane (Figure 1E; Supplementary Figure S3). Visualization of nuclei with DAPI staining showed no overlapping nuclear localization with ShGPCR1::mGFP (Figure 1E; Supplementary Figure S3). The mGFP alone was detected mostly in the cytosol (Supplementary Figure S3). Together, the results suggest that ShGPCR1 is predominantly a membrane-localized protein and supports the in silico predictions (Figure 1C) in a manner similar to other GPCRs (Ma et al., 2015), possibly functioning by maintaining cell membrane integrity under stress.

To further understand the in planta function of ShGPCR1 in sugarcane stress signaling, we constitutively expressed a representative full-length CDS of ShGPCR1 (EC2 allele, Table 1) under the control of the maize Ubiquitin 1 promoter (pUbi) and the double terminator from Cauliflower mosaic virus 35S (35ST) and Agrobacterium (Agrobacterium tumefaciens) nopaline synthase (Tnos; Figure 2A), using established sugarcane DNA bombardment transformation methods (Bower et al., 1996; Ramasamy et al., 2018). We determined the presence of the ShGPCR1 transgene and co-transformed bar selectable marker in transgenic sugarcane plants by PCR, with primers spanning the pUbi-bar-35STTnos (in pTEM73) cassette (Figure 2C; Supplementary Figure S8). We then identified several ShGPCR1 transgenic lines with simple (1–2 insertions) or complex (>2–7 insertions) integration events, as detected by Southern blot hybridization with a full-length ShGPCR1 CDS probe (Supplementary Figure S4). Multiple integration sites are a typical outcome of sugarcane biolistic transformation, which is largely considered preferable because of its applicability to diverse sugarcane genotypes, in contrast to Agrobacterium-mediated methods, which exhibit strong genotype specificity (Ramasamy et al., 2018). We confirmed overexpression of ShGPCR1 in the transgenic plants by RT-PCR using ShGPCR1-specific primers. The expression of sugarcane ANTHRANILATE PHOSPHORIBOSYLTRANSFERASE (APRT2; Casu et al., 2012) was used as housekeeping gene reference (Figures 2D,E; Supplementary Figure S8). Next, we micro-propagated three independent transgenic plants (1–3) in bioreactors (Da Silva et al., 2020) that showed stable expression of ShGPCR1 and no deleterious growth phenotypes relative to non-transgenic (NT) controls (Figure 2B) to scale up plant material for diverse stress-tolerance studies.

Ca²⁺ is a critical divalent cation for plant cells. The intracellular Ca²⁺ level is very low in cytosol (~100–200nM), while a high level of Ca²⁺ can be found in apoplasts (10μM–10mM), the vacuole (0.2mM to 1–5mM), and the endoplasmic reticulum (varies between 50and 500μM; Stael et al., 2012). Growing evidence indicates that changes in intracellular Ca²⁺ level or sensitivity play crucial roles in plants’ biotic and abiotic stress responses (Huda et al., 2013). Specifically, GPCRs have been known to trigger Ca²⁺ influx and signaling in plants (Ma et al., 2015). To test whether ShGPCR1 affects cellular Ca²⁺ fluxes, we measured Ca²⁺ levels in an ShGPCR1-OE sugarcane line using the potent Ca²⁺-sensitive fluorescent dye Fluo-4AM (Ma et al., 2015). We found a significant enhancement in the Ca²⁺ release in response to GTP between ShGPCR1:OE and NT leaf cells (ShGPCR1:OE: 0.048±0.002; NT: 0.013±0.002; p=2.5E⁻²²; Figures 5A–C). To further investigate whether there is any difference in the global Ca²⁺ levels between ShGPCR1:OE and NT leaf cells, we performed Ca²⁺ imaging in the presence of ionomycin, a bonafide calcium ionophore known to increase global Ca²⁺ levels in cells (Qiu et al., 2020). Our results showed that there was no significant difference in the overall Ca²⁺ level between ShGPCR1:OE and NT leaf cells (ShGPCR1:OE: 0.17±0.029; NT: 0.13±0.009; p=0.15; Figures 5D–F). Taken together, these results indicate that ShGPCR1 affects intracellular Ca²⁺ levels in response to GTP. Given that ShGPCR1 predominantly localized to the plasma membrane, the ShGPCR1-mediated Ca²⁺ increase in response to GTP could be due to an influx from an apoplast source to the cytosol. This increase in intracellular Ca²⁺ levels via ShGPCR1 could further trigger a signaling cascade to impart abiotic stress tolerance (Figure 6).