Introduction

Aluminum (Al) is an abundant metal on the earth’s surface, constituting approximately 7% metal content, and its toxicity is a major restrictive factor for crop growth in acidic soils, especially in southern China. Once the soil pH < 5.0, Al3+ is easily dissolved, resulting in the inhibition of plant root elongation and induced appearance of programmed cell death (PCD) (Huang et al. 2014a; Zhan et al. 2013). PCD is genetically controlled cell death that is activated in response to environmental stresses in plants. The important role of caspases in regulating apoptosis in animals strongly influenced research on PCD in plants, resulting in many caspase-like activities being detected in plants (Acosta-Maspons et al. 2014; Coll et al. 2010; Lam and Zhang 2012). However, instead of caspases, metacaspases (caspase-like proteins, MCs) are involved in PCD events in plants (Fagundes et al. 2015; Minina et al. 2014).

Plant MCs are divided into two types, I and II, on the basis of their structure, with the former having an N-terminal prodomain that is not present in the latter (Choi and Berges 2013; Fagundes et al. 2015; Lam and Zhang 2012; Tsiatsiani et al. 2011). Evidence suggests that MCs are involved in signaling, developmental regulation and stress-induced PCD of plants (Fagundes et al. 2015; Huang et al. 2015; Lam and Zhang 2012). Several MC genes have been identified and play crucial roles in plant PCD. For example, in Arabidopsis thaliana, three type I (AtMC1–3) and six type II (AtMC4–9) genes have been identified (Tsiatsiani et al. 2011). AtMC1 and AtMC2 antagonistically control hypersensitive response-triggered PCD (Coll et al. 2010). AtMC1 is a positive regulator of cell death and depends on the conserved catalytic residues, whereas AtMC2 negatively regulates cell death and is independent of the catalytic residues (Coll et al. 2010). AtSERPIN1 is an inhibitor of AtMC1-mediated cell death and autocatalytic processing in planta (Lema Asqui et al. 2017).

Among the type II MCs, AtMC4 is a positive regulator of the biotic and abiotic stress-induced PCD in Arabidopsis (Watanabe and Lam 2011). AtMC5 mediates apoptosis triggered by oxidative stress or aging in yeast (Watanabe and Lam 2005). AtMC8’s expression is strongly upregulated by the oxidative stresses of UVC and H2O2 (He et al. 2008). AtMC9 is required for the efficient progression of autolysis in xylem cell death (Bollhoner et al. 2013; Escamez et al. 2016) and PCD in leaves (Wrzaczek et al. 2015). Additionally, the mcII gene in Norway spruce (Bozhkov et al. 2005), MC4 in wheat (Wang et al. 2012), MC9 in pepper (Kim et al. 2013), NbMCA1 in tobacco (Hao et al. 2007) and MCII-1 in litchi (Wang et al. 2017a) also appear to function in PCD and defense responses.

Recently, MC genes have been systematically predicted and identified through genome-wide screening, including in rubber tree (Liu et al. 2016a), grape (Zhang et al. 2013), rice (Wang and Zhang 2014), maize (Ahmad et al. 2012), and tomato (Hoeberichts et al. 2003; Liu et al. 2016b). Thus, MCs are vital regulators of plant PCD. However, to date, no MC genes have been reported in peanut.

Peanut, as an important oil crop worldwide, is mostly planted in acidic soils in China, especially in southern China. A negative correlation between Al-triggered PCD and tolerance to Al exists in peanut (Zhan et al. 2013). Different types of caspase-like proteases are activated in root tips of peanut during Al-induced PCD, and caspase-3-like protease is a crucial executioner (Huang et al. 2014b; Yao et al. 2016). MC proteases having caspase-3-like abilities have also been identified in Papaver rhoeas, Arabidopsis and poppy (Chai et al. 2017; Tan et al. 2016; Wilkins et al. 2015). In this study, we report that AhMC1, a type I MC, enhances Al-induced PCD in peanut root tips.

Materials and methods

Plant material and treatment

Plant preparation and treatment were performed as described by Yao et al. (2016) with slight modifications. Seeds of peanut cultivars Zhonghua 2 (ZH2, Al-sensitive) and 99–1507 (Al-tolerant) were incubated in wet perlite under dark conditions at 26 ± 2 °C for approximately 3 d to germinate. Seedlings were cultured in Hoagland’s nutrient solution (pH 4.2) under a 12- h light period with a photon flux density of 30–50 μmol m−2 s−1 from daylight fluorescent tubes (Philips Chinese Ltd., Shanghai, China) at 26 ± 2 °C when the seedlings had 2- cm long roots (1 d after germination). When the third leaves emerged, the seedlings were pretreated in Hoagland’s nutrient solution supplemented with 0.1 mM CaCl2 (pH 4.2) for 24 h.

For gene cloning, ZH2 seedlings were transferred to fresh Hoagland’s nutrient solution supplemented with 100 μM AlCl3 for 12 h, and approximately 1 cm root tips were removed and placed on filter paper. For real-time quantitative (RT-q) PCR and western blot analyses, leaves, stems, and root tips of the two cultivars were sampled at 0, 4, 8, 12, and 24 h during the 100- μM AlCl3 treatment, and plants not receiving the Al treatment were used as the controls. Samples from all the experiments were immediately frozen in liquid nitrogen and stored at −80 °C until used for RNA and protein extractions. Leaves of tobacco (Nicotiana tabacum cv. K326) were decontaminated in 0.1% HgCl2 (w/v) for 8 min, followed by 5% (v/v) detergent for 15–20 min, and running water for 30 min. Subsequently, a sample of 10 cm × 20 cm was taken after four or five washes in sterile distilled water and cultured on Murashige and Skoog (MS) medium supplemented with 3% (w/v) sucrose, 0.5% (w/v) agar powder (gel strength: >1400 g cm−2), and 1.0 mg L−1 6-Benzylaminopurine (6-BA) to induce cluster-buds. To obtain the seedlings, the cluster-buds were cultured in MS solid medium supplemented with 2.5 mg L−1 6-BA and 0.5 mg L−1 1-naphthylacetic acid (NAA), and subcultured for 20 d. The cultures were incubated at 26 ± 2 °C with a 16- h/8- h light/dark photocycle. Leaves were used as explants for genetic transformations.

Cloning and structural analysis of AhMC1

Total RNA from ZH2 root tips was extracted using the TRIzol method (Catalog No. 15596–018; Invitrogen Life Technologies, Carlsbad, California, USA) as described in the manufacturer’s instructions. A first-strand cDNA template for RACE-PCR amplification was synthesized according to the manufacturer’s instructions (Catalog No. 1621, TaKaRa Bio, Dalian, China). All the primers for PCR amplification were designed using Primer Premier 5.0 software (Table S1) and amplified full-length cDNAs. Each PCR reaction was carried out in a 25- μL final volume, containing 12.5 μL Taq polymerase Master Mix, 1 μL first-strand cDNA (approximately 10 ng), 1 μL PCR primers and 9.5 μL ddH2O. The reaction procedure is shown in Table S1. The 3′- and 5′-RACE reactions were performed using SMARTer® RACE 5′/3′ Kit. The RACE-PCR products were purified using a Gel Extraction Kit (Catalog No. DP209, Tiangen Biotech, Beijing, China), and the amplified products were cloned into vector pMD19-T (Catalog No. D102A, TaKaRa Bio, Dalian, China). The resultant recombinant plasmid was transformed into Escherichia coli strain DH5α and then sent to BGI Genomics Co., ltd. (Shenzhen, China) for sequencing. The 3′- and 5′- termination fragments were spliced together using DNAman (Lynnon Biosoft, San Ramon, USA) to obtain the full-length AhMC1 gene. All the sequences were analyzed based on non-redundant databases at NCBI (http://www.ncbi.nlm.nih.gov/) using BLAST. The theoretical molecular weight and isoelectric point of AhMC1 protein were predicted by ExPASy (http: //web.expasy.org/compute_pi/). Conserved domains were analyzed using the conserved domain search service at the NCBI web (http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi). The subcellular localization was predicted using PSORT prediction software (http://psort.hgc.jp/form.html). Transmembrane structures were analyzed using the TMHMM Server v.2.0 (http://www.cbs.dtu.dk/services/TMHMM/). Protein signal peptides were analyzed using the SignalP 4.1 server (http://www.cbs.dtu.dk/services/SignalP). The sequence of AhMC1 was uploaded to the NCBI GenBank database under accession number MN400565. The genomic DNA sequences of AhMC1 were obtained through a BLASTN search of the genomic databases (https://www.peanutbase.org/blast/) of Arachis duranensis and Arachis ipaensis, which are the diploid ancestors of cultivated peanut (Bertioli et al. 2016), and verified by PCR. The exon–intron structure of AhMC1 was identified using coding sequence alignments to corresponding genomic sequences.

Multiple sequence alignments and phylogenetic analysis

MEGA 6.0 (http://www.megasoftware.net) was used to conduct the phylogenetic analyses of the MCs and AhMC1. MC members used for the phylogenetic analysis were obtained from the GenBank (https://www.ncbi.nlm. nih.gov/genbank/) and TAIR (http://www.arabidopsis.org/) databases. The amino acid sequences of 21 members were aligned using ClustalX 1.83 software, and the phylogenetic trees were constructed with the neighbor-joining approach. The bootstrap values, and gaps and missing data treatments, were set at 1000 replicates and complete deletion options, respectively.

The MC genes were named using the first letter of the genus followed by the first letter of the species. Members of MCs used for phylogenetic analysis were indicated as following: AtMC1, AT1G02170.1 (A. thaliana); AtMC2, AT4G25110; AtMC3, AT5G64240.2; AtMC4, AT1G79340.1; AtMC5, AT1G79330.1; AtMC6, AT1G79320.1; AtMC7, AT1G79310.1; AtMC8, AT1G16420.1; AtMC9, AT5G04200.1; VaMC3, XP_017408034.1 (Vigna angularis); AiMC1, XP_016179981.1 (A. ipaensis); AiMC3, XP_016180860.1; AdMC1, XP_015943473.1 (A. duranensis); LaMC3, XP_019426387.1 (Lupinus angustifolius); CaMC3, XP_004485587.1 (Cicer arietinum); GmMC1, KRH43894.1 (Glycine max); MtMC1, XP_003593211.1 (Medicago truncatula); VrMC3, XP_014519171.1 (Vigna radiata var. radiata); CcMC1, XP_020222169.1 (Cajanus cajan); and GsMC1, A0A0B2R4L3 (Glycine soja).

Real-time quantitative PCR analysis

RT-qPCR analyses of AhMC1 expression in different tissues (leaves, stems, and roots) of peanut were performed with SYBR® Premix Ex Taq™ (Bio-Rad Laboratories, Hercules, CA, USA) using the CFX96™ Real-Time System, according to the supplier’s manuals. The UBQ10R gene (accession number EG030441) (Table S2) was used as the internal control to quantify the relative transcript levels (Yao et al. 2019). The primers of AhMC1 and UBQ10R are shown in Table S1. The reactions were prepared in a total volume of 20 μL containing 10 μL SYBR® Premix Ex Taq™, 2 μL cDNA, 1 μL each 10- μM primer, and 6 μL ddH2O. The relative level of gene expression was calculated using the 2−ΔΔCt formula. Three technical replicates were run for each biological sample.

Western blot analysis

Approximately 200 mg of peanut root tips were ground in liquid nitrogen and then transferred to 500 μL protein lysis buffer (Catalog No. P0013B, Beyotime Institute of Biotechnology, Shanghai, China) containing 50 mM Tris–HCl (pH 7.4), 0.15 M NaCl, 10% glycerol (w/v), 0.1% sodium dodecyl sulfate (SDS) (w/v), 1% TritonX-100 (w/v), 1% sodium deoxycholate (w/v) and 1 mM phenylmethanesulfonyl fluoride. The extracts were placed on ice for 20 min, followed by centrifugation at 12,000×g for 15 min at 4 °C. The supernatants were transferred to a fresh 1.5 mL tube and 6–10 times the volume of 1:1 acetone:alcohol (v/v) was added. Samples were centrifuged at 15,000×g for 10 min at 4 °C and then maintained at −20 °C for 1 h or overnight. The pellets were redissolved in 100 μL lysis buffer after being air dried for 10 min. The lysates were either used immediately or stored at −80 °C. The protein concentrations were determined using an enhanced BCA protein assay kit (Catalog No. P0010S, Beyotime Institute of Biotechnology, Shanghai, China) according to the manufacturer’s instructions.

For western blotting, equal amounts of lysates were mixed with 4× SDS-loading dyes treated for 10 min at 99 °C and electrophoresed in 12% SDS-polyacrylamide gels. The separated proteins were transferred to polyvinylidene fluoride membranes (Catalog No. IPVH00010, Millipore, Burlington, MA, USA) using a trans-blot cell (Bio-Rad) after electrophoresis. The membranes were blocked with 5% Difco™ Skim milk (Catalog No. D8340, Solarbio Science & Technology Co, Ltd., Beijing, China) at least 1.5 h. After washing three times with 1× TBS-T (Catalog No. T1081–500, Solarbio Science & Technology Co, Ltd., Beijing, China) for 5 min each, membranes were incubated with a polyclonal primary antibody to the whole AhMC1 peptide produced by Ab-mart Company (Shanghai, China) at a 1:500 dilution overnight. After washing three times in the same washing buffer, for a total of 15 min, each membrane was incubated with a secondary rabbit anti-goat antibody conjugated with horseradish peroxidase conjugate (Catalog No. ZB-2306, ZSGB-Bio, Beijing, China) at a 1:5000 dilution at room temperature for 2 h. After several washes with the same washing buffer, 5 min each time, an ECL western blotting analysis system was used to detect the substrates using an ECL chemiluminescence kit (Catalog No. CW0049B, CWBio, Beijing, China) according to the manufacturer’s instructions. The integrated optical densities (IODs) of the bands produced by western blotting were analyzed using Gel-pro Analyzer 4.0 software (Media Cybernetics, Bethesda, MD, USA) (Cheng et al. 2013). The relative IODs of the bands were calculated using the ratios of the IOD of AhMC1 protease to the IOD of β-Actin protease.

Subcellular localization of AhMC1 determined by immune-electron microscopy

To investigate the ultrastructural localization and changes after Al treatments, the root tips of ZH2 were treated with 100 μM AlCl3 for 0, 4, 8, 12, and 24 h. The root tips (approximately 2 mm) were fixed in 100 mM phosphate buffer (pH 7.2) containing 4% methanol, and 0.5% glutaraldehyde for 2 h at 4 °C. The tissues were dehydrated in an ethanol series, permeated in a K4M series at −20 °C and embedded in K4M araldite resin. Ultrathin sections were cut with an LKB ultra microtome (8800, LKB, Bromma, Sweden) and collected on nickel grids for the immune colloidal gold technique.

Grid-mounted tissue sections were processed for colloidal gold immunocytochemistry by incubating the sections with primary antibody (1:100 dilution), and subsequently the immunolabeled patterns were visualized with a protein A-colloidal gold complex (14- nm gold particles). This was followed by conventional staining with 2% uranyl acetate in 50% methanol for 10 min and then lead citrate for 7 min. The sections were examined under a transmission electron microscope (JEM-1200EX, JEOL, Tokyo, Japan) operated at 80 KV.

Protein expression, purification and activity assay

Recombinant pET-15b-AhMC1 was transformed into E. coli strain Rosetta (DE3) (Sangon Biotech Co., Ltd., Shanghai, China). The strain cells were cultured in LB liquid media supplemented with ampicillin (50 μg mL−1) and chloramphenicol (34 μg mL−1) at 37 °C with a shaking at 220 rpm for 2–3 h until the OD600 value reached 0.4–0.6. They were subsequently induced using 0.5 mM isopropyl-β-D-thiogalactoside at 25 °C overnight and then at 37 °C for 4 h with a shaking at 220 rpm. Subsequently, the cells were pelleted by centrifugation at 4000×g for 10 min and resuspended in 500 μL Tris-buffered saline (pH 7.4) containing 8 M urea, 50 mM Tris-HCl, and 300 mM NaCl. SDS-PAGE was performed, using 40 μL supernatant mixed with 10 μL 5× protein loading buffer treated for 10 min at 99 °C. After ultrasonication on ice for 20 min, the supernatant was collected by centrifugation at 12,000×g for 20 min at 4 °C and then purified using Ni-NTA affinity chromatography as described by Wang et al. (2017b). Purified AhMC1 was confirmed through western blotting experiments. Fluorogenic Ac-DEVD-pNA, as a substrate of caspase-3, was used to measure enzymatic activities as described previously (Yao et al. 2016).

Tobacco AhMC1 transformation and transgenic plant confirmation

To investigate the roles of AhMC1 in plants, transgenic tobacco plants were generated. The sense (ST) and transgenic genes were cloned using cDNA of ZH2 as the template and the primer pairs shown in Table S1. The genes were independently inserted into a pBI121-eGFP vector that contained green fluorescent protein (GFP), cauliflower mosaic virus 35S (CaMV35S) promoter and nopaline synthase terminator using XhoI and SpeI restriction sites to generate the recombination plasmids pBI121-AhMC1-eGFP. Subsequently, the recombinant plasmids were independently transformed into N. tabacum cv. K326 using an Agrobacterium-mediated transformation method. From 60 putative transgenic tobacco seedlings, 30 ST plants were propagated in selective MS culture medium supplemented with 0.5 mg L−1 6-BA, 0.2 mg L−1 NAA, and 100 mg L−1 kanamycin (Kan), and rooted in 1/2 MS medium supplemented with 0.5 mg L−1 NAA and 100 mg L−1 Kan. The transgenic plants were produced using the leaf-disk infection method in a 100 mg L−1 Kan-selection culture. T1 seedlings were used for transgenic plant confirmation. The green fluorescence of GFP was detected using a confocal laser scanning microscope with excitation at 488 nm and emission capture at 500–550 nm. Simultaneously, genomic DNA was extracted from roots of transgenic T1 generations and wild-type (WT) tobacco plants using the cetyl trimethyl ammonium bromide method with a slight modification (Gilio et al. 2017). The specific primers used for PCR confirmation are shown in Table S1, and each PCR reaction was carried out in a final volume of 25 μL, containing 12.5 μL Taq polymerase Master Mix, 1 μL first-strand cDNA (approximately 10 ng), 1 μL PCR primers, and 9.5 μL ddH2O.

Assays of relative root growth, cell death and Al content in transgenic tobacco

The effects of different treatments on root growth, cell death and Al content were determined as described by Yao et al. (2016) in two peanut cultivars. The relative root growth was determined by measuring the main root length, and the relative root elongation was calculated using the following formula: (L24h − L0h) × 100% L0h−1, where L24h represents the root length after the 24- h Al treatment and L0h represents the initial root length before the Al treatment.

Assay of caspase-3-like activities in transgenic tobacco

The caspase-3-like activity was determined as described by Yao et al. (2016). In brief, root tips (1.0 ± 0.1 cm) were homogenized in lysis buffer (approximately 100 μL/3–10 mg sample). The homogenate was immediately collected by centrifugation at 12,000×g for 10 min at 4 °C. The supernatant was used for the reactions or stored at −80 °C. Caspase-3-like activities were detected using a 100- μL reaction mixture containing 25 μL extract, 5 μL 2 mM caspase substrate, and 70 μL caspase assay buffer, at 405 nm with a microplate reader (Tecan Infinite RF200 pro, Swiss) after incubation under continuous dark conditions at 37 °C for 2 h. The protein concentration of the extract was determined quantitatively using the Bradford assay.

RT-qPCR analysis of AhMC1 in transgenic tobacco

The expression of AhMC1 in the T1 generation of transgenic tobacco plants under Al stress was detected using RT-qPCR. NtActin transcripts (Table S1) were used as an internal control in tobacco K326 (Pena et al. 2017) to quantify AhMC1 expression. The 35- cycle reaction was carried out in a total volume of 20 μL, containing 10 μL SYBR® Premix Ex Taq™, 2 μL cDNA, 1 μL of each 10 μM primer, and 6 μL ddH2O. The RT-qPCR analysis method was the same as for AhMC1 expression in peanut.

Assays of maleic dialdehyde (MDA), superoxide dismutase (SOD), peroxidase (POD), and catalase (CAT) activities in transgenic tobacco

MDA was measured using the thiobarbituric acid (TBA)-based colorimetric method as described by Phan et al. (2016). Superoxide dismutase (SOD), peroxidase (POD), and catalase (CAT) activities were determined as described by Pan et al. (2017). In brief, SOD activity was determined by measuring the inhibition of photochemical reduction of nitro-blue-tetrazolium at 560 nm, and POD activity was determined by measuring the rate of guaiacol oxidation and monitoring the absorbance at 470 nm. CAT activity was determined by detecting the decomposition of H2O2 and monitoring the decrease in the absorbance at 240 nm over 3 min.

Statistical analyses

Three repetitions were performed to determine each value and their standard deviations. Data were analyzed using IBM SPSS Statistics 19.0 software (Ehningen, Germany) and presented as the means ± standard deviations. The statistical significance was determined using Student’s t test. The values were plotted using the means ± standard errors (SEs) of the replications (n = 3). Values in figures marked with different lowercase and capital letters are significantly different at 0.05 and 0.01 probability levels, respectively.

Results

Cloning and structural features of AhMC1

AhMC1 had a 1068-bp open reading frame, encoding a peptide of 355 amino acids, with a molecular weight and isoelectric point of 39,669.01 Da and 6.40, respectively. It contained five exons interrupted by four introns (Figs. S1 and 1), but no results were generated using PSORT software online to predict AhMC1’s subcellular localization. Transmembrane structures and a signal peptide analysis showed that AhMC1 had no transmembrane helices nor a signal peptide, indicating that it should localize on the cytoplasmic matrix or organelle substrate. A comparison of the deduced AhMC1 amino acid sequence with GenBank and EMBL databases revealed a high degree of conservation (Fig. S2).

Fig. 1
figure 1

Exon-intron structures of AhMC1 gene. The exons are showed with orange boxes and introns by black lines. Numbers show the length of gene’s exons and introns in bp

Full size image

As seen in Fig. S2, the AhMC1 protein contained a putative conserved caspase-like catalytic domain peptidase C_14 p20 subunit (InterPro accession No. IPR001309) at the N-terminus and a non-catalytic p10 subunit (IPR002138) in the C-terminal domain. A region of 42 aa, which was shorter than that of AtMC4–9 in A. thaliana, linked the p20 and p10 subunits. Additionally, a lesion-simulating disease-1 (LSD1)-type domain was also found at the N-terminus, containing two CXXC-type zinc finger structures (IPR005735). LSD1, which encodes C2H2 zinc finger protein GATA-type transcription factors, negatively regulated the hypersensitive response (Fig. S2). These features indicated that AhMC1 belonged to the type I MC family defined by Fagundes et al. (2015).

Data obtained from the phylogenetic analysis of AhMC1 using MEGA 6.0 software (Fig. 2) revealed two different types (I and II) in the phylogenetic tree. Type I consisted of 15 members, including AhMC1, AtMC1–3 and AdMC1, while type II contained 6 proteins, AtMC4–9.

Fig. 2
figure 2

Phylogenetic relationships of the metacaspase family proteins from peanut and other plant species. The phylogenetic tree was constructed using the Neighbor-Joining method using MEGA 6.0 software. The caspase gene Casp1 (NM_009807.2) from Mus musculus was used as an outgroup to root the tree. Bootstrap values shown at the nodes were set to 1000 replicates, and branch lengths corresponding to the divergence of sequences are indicated by the relative scale (0.2 weighted sequence divergence). Two groups were indicated. The acronym of metacaspase genes were named using the first letter of the genus followed by the first letter of the species: AdMC(Arachis duranensis) Metacaspase; AhMC(Arachis hypogaea); AiMC(Arachis ipaensis); AtMC (Arabidopsis thaliana); CaMC (Cicer arietinum); CcMC(Cajanus cajan); GmMC (Glycine max); GsMC(Glycine soja); LaMC (Lupinus angustifolius); MtMC(Medicago truncatula); VaMC (Vigna angularis); VrMC(Vigna radiata var. radiate)

Full size image

AhMC1 expression was induced by Al stress in peanut

The AhMC1 protein has three forms: native form of approximately 39 KD and two activated forms of 19 KD and 17 KD, which could be detected at the same time (Martinon and Tschopp 2004). The expression of different forms of AhMC1 protein occurred in the root tips of ZH2 and 99–1507 (Fig. 3). For ZH2, the expression levels of the activated forms were greater than the native form (Fig. 3a and b). In contrast, the expression levels of the activated forms in 99–1507 were lower than the native form (Fig. 3c and d). As treatment time increased, the expression levels of all AhMC1-protein forms first increased and then decreased. In addition, the highest expression levels, which were 0.19, 3.57, and 3.15, respectively, occurred at 12 h after treatment (Fig. 3b). For 99–1507, the expression of the AhMC1 protein showed an overall upward trend under Al-stress conditions (Fig. 3c). Significant differences were observed between Al-treated and control samples, with the native form reaching a maximum 0.36 at 24 h (Fig. 3d). Thus, the activated protein expression level in ZH2 at 12 h was significantly higher than in 99–1507 and the native form of the AhMC1 protein in ZH2 was more easily cleaved and activated, thereby inducing PCD, compared with than in 99–1507 under Al-stress conditions.

Fig. 3
figure 3

Expression of AhMC1 in ZH2 and 99–1507cultivar under 100 μM AlCl3 treatment for 0, 4, 8, 12, 24 h. a, Immunoblotting of ZH2; b, Relative IOD of the bands in ZH2; c, Immunoblotting of 99–1507; d, Relative IOD of the bands in 99–1507

Full size image

Subcellular localization of AhMC1

AhMC1’s subcellular localization changed as Al exposure time increased. Limited colloidal gold particles were found in the nuclear membranes of meristematic cells in peanut root tips not exposed to the Al stress, indicating that the synthesis of AhMC1 protein was very low (Fig. 4a–c). Greater amounts of colloidal gold particles were distributed in the nuclear membranes after 4 h of the 100 μM AlCl3 treatment (Fig. 4d). Likewise, limited colloidal gold particles were observed in both the cytoplasm and mitochondria under the same conditions, having disperse distributions (Fig. 4e). Moreover, a great amount of AhMC1 was also observed in the nucleus, cytoplasm, mitochondria, and cell wall, with clustered distributions after 8 h of the Al treatment (Fig. 4f–h). However, after 24 h of treatment, there was less AhMC1 protein in the mitochondria, but more AhMC1 protein in the cytoplasm, indicating that the AhMC1 protein transferred from mitochondria to cytoplasm and accumulated in the cytoplasm during Al stress (Fig. 4i and j). Limited gold particles appeared in nuclei and cell walls (Fig. 4i and j). The AhMC1 polyclonal antibody was replaced by normal rabbit serum in the negative control, revealing that no gold particles were located on mitochondria, cytoplasm, nucleus, cell wall, or other parts of meristematic cells (Fig. 4k and l).

Fig. 4
figure 4

Subcellular localization of AhMC1 by immune-electron microscopy in root tips of ZH2. a-c, 100 μmol/L AlCl3 treatment for 0 h (a: ×40,000; b-c: ×60,000); d-e, 100 μmol/L AlCl3 treatment for 4 h (×60,000); f-h, 100 μmol/L AlCl3 treatment for 8 h (×60,000); i, 100 μmol/L AlCl3 treatment for 12 h (×60,000); j, 100 μmol/L AlCl3 treatment for 24 h (×60,000); k-l, Negative control (k: ×60,000; l: ×40,000). Black arrows showing gold particles. N, Nucleus; NW, Nuclear membranes; CW, Cell wall; M, Mitochondria; G, Golgi apparatus; rER, Rough endoplasmic reticulum

Full size image

Protein expression, purification and activity assay

A large amount of soluble AhMC1 fusion protein was produced by inducing E. coli Rosetta (DE3) cells bearing pET-15b- AhMC1 in LB liquid media with 0.5 mM isopropyl-β-D-thiogalactoside for 8 h at 20 °C. The purified AhMC1 protein exhibited high caspase-3-like protease activity, which was much greater than that in the ZH2 root tip cells (Fig. 5).

Fig. 5
figure 5

The caspase-3-like protease activities in root tip cells of ZH2 and pET-15b-AhMC1 protein. Values are present as the mean ± SD of three biological replications

Full size image

Construction of transgenic tobacco plants

All the seedlings were produced following co-cultivation of WT calli with Agrobacterium tumefaciens carrying both AhMC1 and GFP genes. PCR analysis and green fluorescence protein detection confirmed that AhMC1 had been transformed into tobacco and stably inherited (Fig. 6). PCR indicated that 25 ST and 21 AT plants contained both the AhMC1 and GFP genes (Fig. 6a). The transformation efficiencies were 83.33% for ST plants and 70% for AT plants, respectively, based on Kan resistance (data not shown). The GFP gene in T1 transgenic and WT tobacco plants was observed under a confocal laser scanning microscope and revealed that only transgenic plants produced green fluorescence (A1 and A3, Fig. 6b).

Fig. 6
figure 6

Identification of transgenic tobacco plants. a. Identification of AhMC1 and GFP gene by PCR. Lines from left to right: lanes 1–4, sense transgenic plants; lanes 5–8, antisense transgenic plants; lane (−), negative control (untransformed plants); lane (+), positive control (the Pbi121-AhMC1-eGFP plasmid); lane M, (DL2 000 marker). b. Green fluorescence detected in root tips of transgenic plants and the control by laser scanning confocal microscope. Bar = 100 μm. A1: Fluorescent field of transgenic tobacco; A2: Bright field of transgenic tobacco; A3: Merge field of transgenic tobacco; A4: Fluorescent field of wild-type tobacco; A5: Bright field of wild-type tobacco; A6: Merge field of wild-type tobacco. Bar = 100 μm

Full size image

In addition, the phenotypic differences among WT, ST, and AT plants in T1 were analyzed. The plant types and florescence were almost similar among WT and ST, AT. Compared with WT, ST and AT plants had shorter petioles and smaller leaf blades, but there was not distinct difference between ST and AT (Fig. 7a and b).

Fig. 7
figure 7

The phenotype of transgenic (ST, sense transgenic, AT, antisense transgenic) and WT (wild type) tobacco in T1 generation. a. the whole plants; b. the leaves(from left to right: WT, ST, AT); c. Caspase-3-like activity under Al treatments; d. Relative expression level under Al treatments

Full size image

AhMC1 enhanced Al-induced PCD by increasing the caspase-3-like activity

Caspase-3-like activities were monitored in this study to elucidate the function of AhMC1 accumulation. As shown in Fig. 7c, a significant increase in the caspase-3-like protease activities in root tips of WT, AT, and ST tobacco was observed after Al treatments. The caspase-3-like activity increased along with the Al concentration, but there were no obvious differences between the 100- and 200 μM AlCl3 treatments. The caspase-3-like activity in ST (5.27 μmol·pNA·μg−1 protein) was significantly greater than those in AT (2.22 μmol·pNA·μg−1 protein) and WT (3.53 μmol·pNA·μg−1 protein) plants after 24- h 100 μM AlCl3 treatment, which represented 3.21-, 1.44-, and 2.39- fold increases compared with the control (P value <0.01), respectively.

Likewise, the same change trend of AhMC1 expression in ST, AT, and WT plants occurred with the same Al treatment (Fig. 7d). The expression of AhMC1 increased more steadily and rapidly in root tips treated with greater Al concentrations in ST compared with AT and WT plants, and there was no remarkable difference between AT and WT plants receiving the same treatment (Fig. 7d). Thus, AhMC1 appeared to enhance Al-induced PCD in peanut root tips, which may be triggered by an increased caspase-3-like activity.

Overexpression of AhMC1 decreased resistance to Al stress

Al significantly inhibited root elongation in ST tobacco (Fig. 8a). Compared with the control, 50 μM Al could significantly inhibit the elongation of roots in ST plants, and the relative root elongation (RRE) was decreased by Al treatments. However, this Al treatment did not affect AT and WT plants. A 24- h treatment with 200 μM AlCl3 produced RRE values in ST, AT, and WT of 58.07%, 66.52% and 66.22%, respectively, compared with untreated plants (Fig. 8a). Thus, the overexpression of AhMC1 decreased root elongation.

Fig. 8
figure 8

Relative root length (a), root cell death (b), and Al content (c) in transgenic and WT tobacco plants under Al treatment

Full size image

The integrity of the cell membrane during Al stress was evaluated using the nonpermeable dye, Evans blue. Al significantly increased the cell death of the membrane to differing degrees. Compared with the control, 50 μM AlCl3 rapidly increased root tip cell death in ST plants, which was significantly greater than in AT and WT plants. When root tips were treated with 200 μM AlCl3 for 24 h, there were up to 1.19- and 1.15- fold increases in the cell death in ST plants overexpressing AhMC1 compared with AT and WT plants (Fig. 8b). Therefore, ST plants had greatly increased levels of cell death.

Al accumulation in root tips was detected using an inductively coupled plasma optical emission spectrometry technique. As shown in Fig. 8c, significant increases in Al accumulation in root tips were observed as Al concentrations increased, especially in ST plants. However, compared with AT and WT plants, the Al contents of ST plants were the greatest at the same concentrations. There were significant differences between ST and both AT and WT, but there was no significant difference between AT and WT plants.

AhMC1 enhanced Al-induced PCD by changing the ROS capacity

To investigate the alteration in the redox status after Al treatments, the lipid peroxidation levels in root tips were assessed by measuring the MDA contents. There were no significant differences in the MDA content among ST, AT and WT not receiving an Al treatment. Significant changes in the lipid peroxidation level were observed initially after a 24- h 50 μM AlCl3 treatment. The differences in the MDA contents in ST and WT plants was approximately 1.8- fold at 24 h after the 200 μM AlCl3 treatment. Likewise, there was no significant difference in the MDA contents of AT and WT under Al stress conditions (Fig. 9a). The activities of some antioxidant enzymes were monitored in this work to elucidate the function of AhMC1 accumulation in ST, AT and WT plants. The activities of SOD, POD, and CAT markedly and steadily increased along with the Al concentration compared with the control, but the degree of increase in ST plants was minimum (Fig. 9b–d). The transgenic ST plants showed significantly lower SOD, POD, and CAT activities than those of AT and WT plants after a 24-h 200 μM AlCl3 treatment.

Fig. 9
figure 9

Activities of MDA (a), SOD (b), POD (c) and CAT (d) in transgenic and WT tobacco plants under Al treatment

Full size image

Discussion

MCs are widespread in the Viridiplantae and are divided into two types (I and II) based on amino acid sequence, which have similarities to metazoan “initiator” and “executioner” caspases, respectively (Hill and Nystrom 2015; Uren et al. 2000). In this study, the full-length of AhMC1 in peanut was cloned and characterized. A sequence alignment analysis showed that the amino acid sequence of AhMC1 had high homology with those from other species, especially Leguminosae family caspase-like proteases, and this indicated that AhMC1 was a MC member. In Arabidopsis, both proline (AtMC1 and AtMC2) and glutamine (AtMC3) were enriched in predicted N-terminal prodomains, which represent upstream “initiator” caspases in mammalians and might be responsible for protein–protein interactions between MCs and/or other signaling complexes, leading to subsequent MC activation (Vercammen et al. 2004). Fagundes et al. (2015) hypothesized that the acquisition of the zinc finger domain occurred later during MC gene family’s evolution, and they reported a distribution of 259 type I and 113 type II MCs in 42 plant species. The exon-intron structures in tomato (Liu et al. 2016b), rubber tree (Liu et al. 2016a), and rice (Wang and Zhang 2014) also supported type I MCs having greater intron numbers and short final exons in comparison with type II MCs. Here, the prodomain of AhMC1 protein was enriched with proline, indicating that it might have similar function to those of AtMC1 and AtMC2 (Tsiatsiani et al. 2011). In conclusion, we hypothesize that AhMC1 is a typical member of the type I MCs.

It is likely that MCs carry out various functions. In recent decades, some MC genes have been cloned and characterized, and PCD occurs in response to various pathogens and elicitors, as well as abiotic stresses, in different species (Cao et al. 2012; Coll et al. 2014; Hao et al. 2016; Watanabe and Lam 2011; Wang et al. 2017b). The expression specificity of MCs in cells undergoing PCD was various in different tissues and organs of plants, such as Arabidopsis, rice, rubber tree, and grape (Kwon and Hwang 2013; Liu et al. 2016a; Wang and Zhang 2014). MCs became more active after the generation of signals leading to PCD, such as BAX expression, fungal pathogen attack (Wang et al. 2012), and ultraviolet light and H2O2 exposure (He et al. 2008). Liu et al. (2016b) showed that type I MCs (MC1–6) were significantly expressed in the roots of tomato during abiotic stress- or ethylene-induced PCD. In this study, AhMC1 was highly expressed under Al-stress, and this was consistent with the expression levels of MCs in rice and tomato (Fig. 3a). Notably, the expression of AhMC1 was in accordance with the occurrence of Al-triggered PCD and the activity levels of caspase-like proteases in peanut. Caspase-like proteases were activated during Al-induced PCD in peanut root tips, and there was a positive correlation between PCD and exposure time (Yao et al. 2016; Zhan et al. 2014). Therefore, we focused on the role of caspase-like proteases in Al induced PCD. MCs exhibit a caspase-3-like protease activity (Chai et al. 2017; Li and Franklin-Tong 2008; Tan et al. 2016; Wilkins et al. 2015). Here, there was a change between caspase-3-like protease activity and AhMC1’s expression level during Al-induced PCD, which further verified that AhMC1 has a caspase-3-like protease activity. Additionally, the purified AhMC1 protein which was expressed in E. coli Rossetta (DE3) cells also exhibited a high caspase-3-like protease activity (Fig. 5). Moreover, it is striking that both MC expression and caspase-like activity, possibly acting in the same signaling cascade, are required for effective PCD. Zhao et al. (2013) reported that the downregulation of NtCYS, a cystatin-regulating NtCP14, or the overexpression of NtCP14, a papain-like Cys protease, increased the caspase-like activity and caused premature PCD in tobacco. This suggests that AhMC1 promotes PCD under Al-stress and is likely involved in the caspase-like protease signaling cascade.

Subcellular localization is critical in determining the mode of action for plant MCs. The subcellular localization of AhMC1 during Al-induced PCD was examined. AhMC1’s subcellular localization changed as Al exposure time increased. Thus, the AhMC1 protein may transfer from mitochondria to cytoplasm and accumulate in cytoplasm during Al-stress conditions. Arabidopsis MC9 was also present in the apoplast, nucleus, and cytoplasm, and its subcellular localization changed during the late autolysis process (Bollhoner et al. 2013; Tsiatsiani et al. 2013). However, it was previously reported that OsMC5, -6, and -8 were exclusively localized in the cytoplasm without any localization in the nucleus (Huang et al. 2015). Our studies indicated that a potential cascade of cellular events occurs during Al-induced PCD via mitochondria-dependent pathways. Additionally, ROS is involved in regulating Al-induced PCD (Huang et al. 2014b). In this mechanism, including ROS bursts and caspase 3-like protease activity, AhMC1 protein probably participated downstream of the signal transduction pathway. More studies need to be performed.

Overexpressing AhMC1 in tobacco resulted in plants being less tolerant to Al stress. The T1 generation of AhMC1 transgenic tobacco plants showed diverse resistance levels, which indirectly indicated a function for AhMC1 gene in Al-stress responses as assessed by PCD indexes and physiological traits. Wang et al. (2009) reported that Caenorhabditis elegans apoptotic suppressor Ced-9, a Bcl-2 homologue, inhibited both Al-induced PCD and Al-induced activity of caspase-like vacuolar processing enzyme, a crucial executioner of PCD, in tobacco. Ced-9 significantly alleviated Al’s inhibition of root elongation, decreased Al accumulation in the root tips, and greatly inhibited Al-induced gene expression in early responses to Al, leading to the enhanced Al tolerance of tobacco plants. There is also a negative correlation between Al-induced PCD and Al-resistance in peanut (Huang et al. 2014a; Zhan et al. 2013). Here, we showed that ST plants had shorter roots, and higher Al contents, as well as cell death, AhMC1 expression, and caspase-3-like activity levels, than WT and AT plants under Al stress (Figs. 7c and d, 8a–c). A critical level of activated MC was needed to trigger Al-induced PCD, while in MC-overexpression plants this level was reached at a higher dose, which could promote the occurrence of PCD. The results indicate that AhMC1 induces PCD as a trigger under Al stress.

MDA content was an indicator of the level of lipid peroxidation damage that resulted from oxidative stresses (Bailly et al. 1996). The MDA content of ST plants was significantly greater than those of AT and WT plants after Al treatments. This indicated that lipid peroxidation damage induced by Al was more serious in ST plants than in AT and WT plants. Al induces ROS bursts in mitochondria, which is an important component of the plant’s reaction to toxic Al effects (Matsumoto and Motoda 2013). POD, CAT and POD activities treated with high AlCl3 (>100 μM) were significantly lower in ST plants than in WT plants (Fig. 9b–d), which indicated that the overexpression of AhMC1 inhibited the antioxidant defensive reactions and affected the efficiency of the ROS scavenging system against ROS-mediated injury under Al stress. Additionally, the overexpression of AhMC1 might have played a key role in improving ROS accumulation by inhibiting the activities of the SOD, POD, and CAT enzymes.