2022 Volume 7
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Changes in Pleurotus ostreatus nutritional value and heavy metal profile as a result of supplementation with nano-additive

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  • The present study evaluated the effect of Lithovit-Amino25 on the nutrient profile and heavy metal composition of Pleurotus ostreatus. The product was tested in two doses applied at three different timings: T2: 3 g kg−1/spawning, T3: 3 g kg−1/after first harvest, T4: 3 g kg−1/spawning and after first harvest, T5: 5 g kg−1/spawning, T6: 5 g kg−1/after first harvest, and T7: 5 g kg−1/spawning and after first harvest. Compared to control (T1: non-treated substrate), mushrooms’ fibers and carbohydrates increased in all treatments, recording the highest values in T4 (4.16%) and T3 (18.42%), respectively. Protein content was higher in mushrooms of substrates treated at spawning, with a 0.33% improvement in T5. Fat content decreased in T3, T4, T6, and T5. Total sugars decreased in mushrooms of treated substrates, and glucose was the dominant sugar in mushrooms. Fructose increased in mushrooms of T3 and T4. Calcium, iron, and potassium decreased in mushrooms of treated substrates. Sodium decreased in T3, T5, and T7, magnesium increased in T2, and phosphorus increased only in T2 and T7. Copper content of all treated mushrooms was in the standard safe limit (< 40 ppm), and it decreased in T2, T4, and T5 by around 2.5, 6.6, and 5.1 ppm, compared to control. However, zinc content increased in mushrooms of all treated substrates, and nickel and lead, increased by respective ranges of 2.8−11.88 ppm and 9.1−21 ppm, higher than the safe limits. The product presented a risk of heavy metal bioaccumulation even with a low dose.
  • Aquaporins (AQPs) constitute a large family of transmembrane channel proteins that function as regulators of intracellular and intercellular water flow[1,2]. Since their first discovery in the 1990s, AQPs have been found not only in three domains of life, i.e., bacteria, eukaryotes, and archaea, but also in viruses[3,4]. Each AQP monomer is composed of an internal repeat of three transmembrane helices (i.e., TM1–TM6) as well as two half helixes that are formed by loop B (LB) and LE through dipping into the membrane[5]. The dual Asn-Pro-Ala (NPA) motifs that are located at the N-terminus of two half helixes act as a size barrier of the pore via creating an electrostatic repulsion of protons, whereas the so-called aromatic/arginine (ar/R) selectivity filter (i.e., H2, H5, LE1, and LE2) determines the substrate specificity by rendering the pore constriction site diverse in both size and hydrophobicity[59]. Based on sequence similarity, AQPs in higher plants could be divided into five subfamilies, i.e., plasma membrane intrinsic protein (PIP), tonoplast intrinsic protein (TIP), NOD26-like intrinsic protein (NIP), X intrinsic protein (XIP), and small basic intrinsic protein (SIP)[1017]. Among them, PIPs, which are typically localized in the cell membrane, are most conserved and play a central role in controlling plant water status[12,1822]. Among two phylogenetic groups present in the PIP subfamily, PIP1 possesses a relatively longer N-terminus and PIP2 features an extended C-terminus with one or more conserved S residues for phosphorylation modification[5,15,17].

    Tigernut (Cyperus esculentus L.), which belongs to the Cyperaceae family within Poales, is a novel and promising herbaceous C4 oil crop with wide adaptability, large biomass, and short life period[2327]. Tigernut is a unique species accumulating up to 35% oil in the underground tubers[2830], which are developed from stolons and the process includes three main stages, i.e., initiation, swelling, and maturation[3133]. Water is essential for tuber development and tuber moisture content maintains a relatively high level of approximately 85% until maturation when a significant drop to about 45% is observed[28,32]. Thereby, uncovering the mechanism of tuber water balance is of particular interest. Despite crucial roles of PIPs in the cell water balance, to date, their characterization in tigernut is still in the infancy[21]. The recently available genome and transcriptome datasets[31,33,34] provide an opportunity to address this issue.

    In this study, a global characterization of PIP genes was conducted in tigernut, including gene localizations, gene structures, sequence characteristics, and evolutionary patterns. Moreover, the correlation of CePIP mRNA/protein abundance with water content during tuber development as well as subcellular localizations were also investigated, which facilitated further elucidating the water balance mechanism in this special species.

    PIP genes reported in Arabidopsis (Arabidopsis thaliana)[10] and rice (Oryza sativa)[11] were respectively obtained from TAIR11 (www.arabidopsis.org) and RGAP7 (http://rice.uga.edu), and detailed information is shown in Supplemental Table S1. Their protein sequences were used as queries for tBLASTn[35] (E-value, 1e–10) search of the full-length tigernut transcriptome and genome sequences that were accessed from CNGBdb (https://db.cngb.org/search/assembly/CNA0051961)[31,34]. RNA sequencing (RNA-seq) reads that are available in NCBI (www.ncbi.nlm.nih.gov/sra) were also adopted for gene structure revision as described before[13], and presence of the conserved MIP (major intrinsic protein, Pfam accession number PF00230) domain in candidates was confirmed using MOTIF Search (www.genome.jp/tools/motif). To uncover the origin and evolution of CePIP genes, a similar approach was also employed to identify homologs from representative plant species, i.e., Carex cristatella (v1, Cyperaceae)[36], Rhynchospora breviuscula (v1, Cyperaceae)[37], and Juncus effusus (v1, Juncaceae)[37], whose genome sequences were accessed from NCBI (www.ncbi.nlm.nih.gov). Gene structure of candidates were displayed using GSDS 2.0 (http://gsds.gao-lab.org), whereas physiochemical parameters of deduced proteins were calculated using ProtParam (http://web.expasy.org/protparam). Subcellular localization prediction was conducted using WoLF PSORT (www.genscript.com/wolf-psort.html).

    Nucleotide and protein multiple sequence alignments were respectively conducted using ClustalW and MUSCLE implemented in MEGA6[38] with default parameters, and phylogenetic tree construction was carried out using MEGA6 with the maximum likelihood method and bootstrap of 1,000 replicates. Systematic names of PIP genes were assigned with two italic letters denoting the source organism and a progressive number based on sequence similarity. Conserved motifs were identified using MEME Suite 5.5.3 (https://meme-suite.org/tools/meme) with optimized parameters as follows: Any number of repetitions, maximum number of 15 motifs, and a width of 6 and 250 residues for each motif. TMs and conserved residues were identified using homology modeling and sequence alignment with the structure resolved spinach (Spinacia oleracea) SoPIP2;1[5].

    Synteny analysis was conducted using TBtools-II[39] as described previously[40], where the parameters were set as E-value of 1e-10 and BLAST hits of 5. Duplication modes were identified using the DupGen_finder pipeline[41], and Ks (synonymous substitution rate) and Ka (nonsynonymous substitution rate) of duplicate pairs were calculated using codeml in the PAML package[42]. Orthologs between different species were identified using InParanoid[43] and information from synteny analysis, and orthogroups (OGs) were assigned only when they were present in at least two species examined.

    Plant materials used for gene cloning, qRT-PCR analysis, and 4D-parallel reaction monitoring (4D-PRM)-based protein quantification were derived from a tigernut variety Reyan3[31], and plants were grown in a greenhouse as described previously[25]. For expression profiling during leaf development, three representative stages, i.e., young, mature, and senescing, were selected and the chlorophyll content was checked using SPAD-502Plus (Konica Minolta, Shanghai, China) as previously described[44]. Young and senescing leaves are yellow in appearance, and their chlorophyll contents are just half of that of mature leaves that are dark green. For diurnal fluctuation regulation, mature leaves were sampled every 4 h from the onset of light at 8 a.m. For gene regulation during tuber development, fresh tubers at 1, 5, 10, 15, 20, 25, and 35 d after tuber initiation (DAI) were collected as described previously[32]. All samples with three biological replicates were quickly frozen with liquid nitrogen and stored at −80 °C for further use. For subcellular localization analysis, tobacco (Nicotiana benthamiana) plants were grown as previously described[20].

    Tissue-specific expression profiles of CePIP genes were investigated using Illumina RNA-seq samples (150 bp paired-end reads) with three biological replicates for young leaf, mature leaf, sheath of mature leaf, shoot apex, root, rhizome, and three stages of developmental tuber (40, 85, and 120 d after sowing (DAS)), which are under the NCBI accession number of PRJNA703731. Raw sequence reads in the FASTQ format were obtained using fastq-dump, and quality control was performed using fastQC (www.bioinformatics.babraham.ac.uk/projects/fastqc). Read mapping was performed using HISAT2 (v2.2.1, https://daehwankimlab.github.io/hisat2), and relative gene expression level was presented as FPKM (fragments per kilobase of exon per million fragments mapped)[45].

    For qRT-PCR analysis, total RNA extraction and synthesis of the first-strand cDNA were conducted as previously described[24]. Primers used in this study are shown in Supplemental Table S2, where CeUCE2 and CeTIP41[25,33] were employed as two reference genes. PCR reaction in triplicate for each biological sample was carried out using the SYBR-green Mix (Takara) on a Real-time Thermal Cycler Type 5100 (Thermal Fisher Scientific Oy). Relative gene abundance was estimated with the 2−ΔΔCᴛ method and statistical analysis was performed using SPSS Statistics 20 as described previously[13].

    Raw proteomic data for tigernut roots, leaves, freshly harvested, dried, rehydrated for 48 h, and sprouted tubers were downloaded from ProteomeXchange/PRIDE (www.proteomexchange.org, PXD021894, PXD031123, and PXD035931), which were further analyzed using Maxquant (v1.6.15.0, www.maxquant.org). Three dominant members, i.e., CePIP1;1, -2;1, and -2;8, were selected for 4D-PRM quantification analysis, and related unique peptides are shown in Supplemental Table S3. Protein extraction, trypsin digestion, and LC-MS/MS analysis were conducted as described previously[46].

    For subcellular localization analysis, the coding region (CDS) of CePIP1;1, -2;1, and -2;8 were cloned into pNC-Cam1304-SubN via Nimble Cloning as described before[30]. Then, recombinant plasmids were introduced into Agrobacterium tumefaciens GV3101 with the helper plasmid pSoup-P19 and infiltration of 4-week-old tobacco leaves were performed as previously described[20]. For subcellular localization analysis, the plasma membrane marker HbPIP2;3-RFP[22] was co-transformed as a positive control. Fluorescence observation was conducted using confocal laser scanning microscopy imaging (Zeiss LMS880, Germany): The wavelength of laser-1 was set as 730 nm for RFP observation, where the fluorescence was excited at 561 nm; the wavelength of laser-2 was set as 750 nm for EGFP observation, where the fluorescence was excited at 488 nm; and the wavelength of laser-3 was set as 470 nm for chlorophyll autofluorescence observation, where the fluorescence was excited at 633 nm.

    As shown in Table 1, a total of 14 PIP genes were identified from eight tigernut scaffolds (Scfs). The CDS length varies from 831 to 882 bp, putatively encoding 276–293 amino acids (AA) with a molecular weight (MW) of 29.16–31.59 kilodalton (kDa). The theoretical isoelectric point (pI) varies from 7.04 to 9.46, implying that they are all alkaline. The grand average of hydropathicity (GRAVY) is between 0.344 and 0.577, and the aliphatic index (II) ranges from 94.57 to 106.90, which are consistent with the hydrophobic characteristic of AQPs[47]. As expected, like SoPIP2;1, all CePIPs include six TMs, two typical NPA motifs, the invariable ar/R filter F-H-T-R, five conserved Froger's positions Q/M-S-A-F-W, and two highly conserved residues corresponding to H193 and L197 in SoPIP2;1 that were proven to be involved in gating[5,48], though the H→F variation was found in CePIP2;9, -2;10, and -2;11 (Supplemental Fig. S1). Moreover, two S residues, corresponding to S115 and S274 in SoPIP2;1[5], respectively, were also found in the majority of CePIPs (Supplemental Fig. S1), implying their posttranslational regulation by phosphorylation.

    Table 1.  Fourteen PIP genes identified in C. esculentus.
    Gene name Locus Position Intron no. AA MW (kDa) pI GRAVY AI TM MIP
    CePIP1;1 CESC_15147 Scf9:2757378..2759502(–) 3 288 30.76 8.82 0.384 95.28 6 47..276
    CePIP1;2 CESC_04128 Scf4:3806361..3807726(–) 3 291 31.11 8.81 0.344 95.95 6 46..274
    CePIP1;3 CESC_15950 Scf54:5022493..5023820(+) 3 289 31.06 8.80 0.363 94.57 6 49..278
    CePIP2;1 CESC_15350 Scf9:879960..884243(+) 3 288 30.34 8.60 0.529 103.02 6 33..269
    CePIP2;2 CESC_00011 Scf30:4234620..4236549(+) 3 293 31.59 9.27 0.394 101.57 6 35..268
    CePIP2;3 CESC_00010 Scf30:4239406..4241658(+) 3 291 30.88 9.44 0.432 98.97 6 31..266
    CePIP2;4 CESC_05080 Scf46:307799..309544(+) 3 285 30.44 7.04 0.453 100.32 6 28..265
    CePIP2;5 CESC_05079 Scf46:312254..314388(+) 3 286 30.49 7.04 0.512 101.68 6 31..268
    CePIP2;6 CESC_05078 Scf46:316024..317780(+) 3 288 30.65 7.68 0.475 103.06 6 31..268
    CePIP2;7 CESC_05077 Scf46:320439..322184(+) 3 284 30.12 8.55 0.500 100.00 6 29..266
    CePIP2;8 CESC_14470 Scf2:4446409..4448999(+) 3 284 30.37 8.30 0.490 106.90 6 33..263
    CePIP2;9 CESC_02223 Scf1:2543928..2545778(–) 3 283 30.09 9.46 0.533 106.47 6 31..262
    CePIP2;10 CESC_10007 Scf27:1686032..1688010(–) 3 276 29.16 9.23 0.560 106.05 6 26..256
    CePIP2;11 CESC_10009 Scf27:1694196..1696175(–) 3 284 29.71 9.10 0.577 105.49 6 33..263
    AA: amino acid; AI: aliphatic index; GRAVY: grand average of hydropathicity; kDa: kilodalton; MIP: major intrinsic protein; MW: molecular weight; pI: isoelectric point; PIP: plasma membrane intrinsic protein; Scf: scaffold; TM: transmembrane helix.
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    To uncover the evolutionary relationships, an unrooted phylogenetic tree was constructed using the full-length protein sequences of CePIPs together with 11 OsPIPs and 13 AtPIPs. As shown in Fig. 1a, these proteins were clustered into two main groups, corresponding to PIP1 and PIP2 as previously defined[10,49], and each appears to have evolved into several subgroups. Compared with PIP1s, PIP2s possess a relatively shorter N-terminal but an extended C-terminal with one conserved S residue (Supplemental Fig. S1). Interestingly, a high number of gene repeats were detected, most of which seem to be species-specific, i.e., AtPIP1;1/-1;2/-1;3/-1;4/-1;5, AtPIP2;1/-2;2/-2;3/-2;4/-2;5/-2;6, AtPIP2;7/-2;8, OsPIP1;1/-1;2/-1;3, OsPIP2;1/-2;4/-2;5, OsPIP2;2/-2;3, CePIP1;1/-1;2, CePIP2;2/-2;3, CePIP2;4/-2;5/-2;6/-2;7, and CePIP2;9/-2;10/-2;11, reflecting the occurrence of more than one lineage-specific whole-genome duplications (WGDs) after their divergence[50,51]. In Arabidopsis that experienced three WGDs (i.e. γ, β, and α) after the split with the monocot clade[52], AtPIP1;5 in the PIP1 group first gave rise to AtPIP1;1 via the γ WGD shared by all core eudicots[50], which latter resulted in AtPIP1;3, -1;4, and -1;2 via β and α WGDs; AtPIP2;1 in the PIP2 group first gave rise to AtPIP2;6 via the γ WGD, and they latter generated AtPIP2;2, and -2;5 via the α WGD (Supplemental Table S1). In rice, which also experienced three WGDs (i.e. τ, σ, and ρ) after the split with the eudicot clade[51], OsPIP1;2 and -2;3 generated OsPIP1;1 and -2;2 via the Poaceae-specific ρ WGD, respectively. Additionally, tandem, proximal, transposed and dispersed duplications also played a role on the gene expansion in these two species (Supplemental Table S1).

    Figure 1.  Structural and phylogenetic analysis of PIPs in C. esculentus, O. sativa, and A. thaliana. (a) Shown is an unrooted phylogenetic tree resulting from full-length PIPs with MEGA6 (maximum likelihood method and bootstrap of 1,000 replicates), where the distance scale denotes the number of amino acid substitutions per site. (b) Shown are the exon-intron structures. (c) Shown is the distribution of conserved motifs among PIPs, where different motifs are represented by different color blocks as indicated and the same color block in different proteins indicates a certain motif. (At: A. thaliana; Ce: C. esculentus; PIP: plasma membrane intrinsic protein; Os: O. sativa).

    Analysis of gene structures revealed that all CePIP and AtPIP genes possess three introns and four exons in the CDS, in contrast to the frequent loss of certain introns in rice, including OsPIP1;2, -1;3, -2;1, -2;3, -2;4, -2;5, -2;6, -2;7, and -2;8 (Fig. 1b). The positions of three introns are highly conserved, which are located in sequences encoding LB (three residues before the first NPA), LD (one residue before the conserved L involved in gating), and LE (18 residues after the second NPA), respectively (Supplemental Fig. S1). The intron length of CePIP genes is highly variable, i.e., 109–993 bp, 115–1745 bp, and 95–866 bp for three introns, respectively. By contrast, the exon length is relatively less variable: Exons 2 and 3 are invariable with 296 bp and 141 bp, respectively, whereas Exons 1 and 4 are of 277–343 bp and 93–132 bp, determining the length of N- and C-terminus of PIP1 and PIP2, respectively (Fig. 1b). Correspondingly, their protein structures were shown to be highly conserved, and six (i.e., Motifs 1–6) out of 15 motifs identified are broadly present. Among them, Motif 3, -2, -6, -1, and -4 constitute the conserved MIP domain. In contrast to a single Motif 5 present in most PIP2s, all PIP1s possess two sequential copies of Motif 5, where the first one is located at the extended N-terminal. In CePIP2;3 and OsPIP2;7, Motif 5 is replaced by Motif 13; in CePIP2;2, it is replaced by two copies of Motif 15; and no significant motif was detected in this region of CePIP2;10. PIP1s and PIP2s usually feature Motif 9 and -7 at the C-terminal, respectively, though it is replaced by Motif 12 in CePIP2;6 and OsPIP2;8. PIP2s usually feature Motif 8 at the N-terminal, though it is replaced by Motif 14 in CePIP2;2 and -2;3 or replaced by Motif 11 in CePIP2;10 and -2;11 (Fig. 1c).

    As shown in Fig. 2a, gene localization of CePIPs revealed three gene clusters, i.e., CePIP2;2/-2;3 on Scf30, CePIP2;4/-2;5/-2;6/-2;7 on Scf46, and CePIP2;10/-2;11 on Scf27, which were defined as tandem repeats for their high sequence similarities and neighboring locations. The nucleotide identities of these duplicate pairs vary from 70.5% to 91.2%, and the Ks values range from 0.0971 to 1.2778 (Table 2), implying different time of their birth. According to intra-species synteny analysis, two duplicate pairs, i.e., CePIP1;1/-1;2 and CePIP2;2/-2;4, were shown to be located within syntenic blocks (Fig. 2b) and thus were defined as WGD repeats. Among them, CePIP1;1/-1;2 possess a comparable Ks value to CePIP2;2/-2;3, CePIP1;1/-1;3, and CePIP2;4/-2;8 (1.2522 vs 1.2287–1.2778), whereas CePIP2;2/-2;4 harbor a relatively higher Ks value of 1.5474 (Table 2), implying early origin or fast evolution of the latter. While CePIP1;1/-1;3 and CePIP2;1/-2;8 were characterized as transposed repeats, CePIP2;1/-2;2, CePIP2;9/-2;10, and CePIP2;8/-2;10 were characterized as dispersed repeats (Fig. 2a). The Ks values of three dispersed repeats vary from 0.8591 to 3.0117 (Table 2), implying distinct times of origin.

    Figure 2.  Duplication events of CePIP genes and synteny analysis within and between C. esculentus, O. sativa, and A. thaliana. (a) Duplication events detected in tigernut. Serial numbers are indicated at the top of each scaffold, and the scale is in Mb. Duplicate pairs identified in this study are connected using lines in different colors, i.e., tandem (shown in green), transposed (shown in purple), dispersed (shown in gold), and WGD (shown in red). (b) Synteny analysis within and between C. esculentus, O. sativa, and A. thaliana. (c) Synteny analysis within and between C. esculentus, C. cristatella, R. breviuscula, and J. effusus. Shown are PIP-encoding chromosomes/scaffolds and only syntenic blocks that contain PIP genes are marked, i.e., red and purple for intra- and inter-species, respectively. (At: A. thaliana; Cc: C. cristatella; Ce: C. esculentus; Je: J. effusus; Mb: megabase; PIP: plasma membrane intrinsic protein; Os: O. sativa; Rb: R. breviuscula; Scf: scaffold; WGD: whole-genome duplication).
    Table 2.  Sequence identity and evolutionary rate of homologous PIP gene pairs identified in C. esculentus. Ks and Ka were calculated using PAML.
    Duplicate 1 Duplicate 2 Identity (%) Ka Ks Ka/Ks
    CePIP1;1 CePIP1;3 78.70 0.0750 1.2287 0.0610
    CePIP1;2 CePIP1;1 77.20 0.0894 1.2522 0.0714
    CePIP2;1 CePIP2;4 74.90 0.0965 1.7009 0.0567
    CePIP2;3 CePIP2;2 70.50 0.1819 1.2778 0.1424
    CePIP2;4 CePIP2;2 66.50 0.2094 1.5474 0.1353
    CePIP2;5 CePIP2;4 87.30 0.0225 0.4948 0.0455
    CePIP2;6 CePIP2;5 84.90 0.0545 0.5820 0.0937
    CePIP2;7 CePIP2;6 78.70 0.0894 1.0269 0.0871
    CePIP2;8 CePIP2;4 72.90 0.1401 1.2641 0.1109
    CePIP2;9 CePIP2;10 76.40 0.1290 0.8591 0.1502
    CePIP2;10 CePIP2;8 64.90 0.2432 3.0117 0.0807
    CePIP2;11 CePIP2;10 91.20 0.0562 0.0971 0.5783
    Ce: C. esculentus; Ka: nonsynonymous substitution rate; Ks: synonymous substitution rate; PIP: plasma membrane intrinsic protein.
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    According to inter-species syntenic analysis, six out of 14 CePIP genes were shown to have syntelogs in rice, including 1:1, 1:2, and 2:2 (i.e. CePIP1;1 vs OsPIP1;3, CePIP1;3 vs OsPIP1;2/-1;1, CePIP2;1 vs OsPIP2;4, CePIP2;2/-2;4 vs OsPIP2;3/-2;2, and CePIP2;8 vs OsPIP2;6), in striking contrast to a single one found in Arabidopsis (i.e. CePIP1;2 vs AtPIP1;2). Correspondingly, only OsPIP1;2 in rice was shown to have syntelogs in Arabidopsis, i.e., AtPIP1;3 and -1;4 (Fig. 2b). These results are consistent with their taxonomic relationships that tigernut and rice are closely related[50,51], and also imply lineage-specific evolution after their divergence.

    As described above, phylogenetic and syntenic analyses showed that the last common ancestor of tigernut and rice is more likely to possess only two PIP1s and three PIP2s. However, it is not clear whether the gene expansion observed in tigernut is species-specific or Cyperaceae-specific. To address this issue, recently available genomes were used to identify PIP subfamily genes from C. cristatella, R. breviuscula, and J. effuses, resulting in 15, 13, and nine members, respectively. Interestingly, in contrast to a high number of tandem repeats found in Cyperaceae species, only one pair of tandem repeats (i.e., JePIP2;3 and -2;4) were identified in J. effusus, a close outgroup species to Cyperaceae in the Juncaceae family[36,37]. According to homologous analysis, a total of 12 orthogroups were identified, where JePIP genes belong to PIP1A (JePIP1;1), PIP1B (JePIP1;2), PIP1C (JePIP1;3), PIP2A (JePIP2;1), PIP2B (JePIP2;2), PIP2F (JePIP2;3 and -2;4), PIP2G (JePIP2;5), and PIP2H (JePIP2;6) (Table 3). Further intra-species syntenic analysis revealed that JePIP1;1/-1;2 and JePIP2;2/-2;3 are located within syntenic blocks, which is consistent with CePIP1;1/-1;2, CePIP2;2/-2;4, CcPIP1;1/-1;2, CcPIP2;3/-2;4, RbPIP1;1/-1;2, and RbPIP2;2/-2;5 (Fig. 2c), implying that PIP1A/PIP1B and PIP2B/PIP2D were derived from WGDs occurred sometime before Cyperaceae-Juncaceae divergence. After the split with Juncaceae, tandem duplications frequently occurred in Cyperaceae, where PIP2B/PIP2C and PIP2D/PIP2E/PIP2F retain in most Cyperaceae plants examined in this study. By contrast, species-specific expansion was also observed, i.e., CePIP2;4/-2;5, CePIP2;10/-2;11, CcPIP1;2/-1;3, CcPIP2;4/-2;5, CcPIP2;8/-2;9, CcPIP2;10/-2;11, RbPIP2;3/-2;4, and RbPIP2;9/-2;10 (Table 3 & Fig. 2c).

    Table 3.  Twelve proposed orthogroups based on comparison of representative plant species.
    Orthogroup C. esculentus C. cristatella R. breviuscula J. effusus O. sativa A. thaliana
    PIP1A CePIP1;1 CcPIP1;1 RbPIP1;1 JePIP1;1 OsPIP1;3 AtPIP1;1, AtPIP1;2,
    AtPIP1;3, AtPIP1;4,
    AtPIP1;5
    PIP1B CePIP1;2 CcPIP1;2, CcPIP1;3 RbPIP1;2 JePIP1;2
    PIP1C CePIP1;3 CcPIP1;4 RbPIP1;3 JePIP1;3 OsPIP1;1, OsPIP1;2
    PIP2A CePIP2;1 CcPIP2;1 RbPIP2;1 JePIP2;1 OsPIP2;1, OsPIP2;4,
    OsPIP2;5
    AtPIP2;1, AtPIP2;2,
    AtPIP2;3, AtPIP2;4,
    AtPIP2;5, AtPIP2;6
    PIP2B CePIP2;2 CcPIP2;2 RbPIP2;2 JePIP2;2 OsPIP2;2, OsPIP2;3
    PIP2C CePIP2;3 CcPIP2;3 RbPIP2;3, RbPIP2;4
    PIP2D CePIP2;4, CePIP2;5 CcPIP2;4, CcPIP2;5 RbPIP2;5
    PIP2E CePIP2;5 CcPIP2;5 RbPIP2;6
    PIP2F CePIP2;6 CcPIP2;6
    PIP2G CePIP2;7 CcPIP2;7 RbPIP2;7 JePIP2;3, JePIP2;4
    PIP2H CePIP2;8 CcPIP2;8, CcPIP2;9 RbPIP2;8 JePIP2;5 OsPIP2;6 AtPIP2;7, AtPIP2;8
    PIP2I CePIP2;9, CePIP2;10,
    CePIP2;11
    CcPIP2;10, CcPIP2;11 RbPIP2;9, RbPIP2;10 JePIP2;6 OsPIP2;7, OsPIP2;8
    At: A. thaliana; Cc: C. cristatella; Ce: C. esculentus; Je: J. effuses; Os: O. sativa; Rb: R. breviuscula; PIP: plasma membrane intrinsic protein.
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    Tissue-specific expression profiles of CePIP genes were investigated using transcriptome data available for young leaf, mature leaf, sheath, root, rhizome, shoot apex, and tuber. As shown in Fig. 3a, CePIP genes were mostly expressed in roots, followed by sheaths, moderately in tubers, young leaves, rhizomes, and mature leaves, and lowly in shoot apexes. In most tissues, CePIP1;1, -2;1, and -2;8 represent three dominant members that contributed more than 90% of total transcripts. By contrast, in rhizome, these three members occupied about 80% of total transcripts, which together with CePIP1;3 and -2;4 contributed up to 96%; in root, CePIP1;1, -1;3, -2;4, and -2;7 occupied about 84% of total transcripts, which together with CePIP2;1 and -2;8 contributed up to 94%. According to their expression patterns, CePIP genes could be divided into five main clusters: Cluster I includes CePIP1;1, -2;1, and -2;8 that were constitutively and highly expressed in all tissues examined; Cluster II includes CePIP2;2, -2;9, and -2;10 that were lowly expressed in all tested tissues; Cluster III includes CePIP1;2 and -2;11 that were preferentially expressed in young leaf and sheath; Cluster IV includes CePIP1;3 and -2;4 that were predominantly expressed in root and rhizome; and Cluster V includes remains that were typically expressed in root (Fig. 3a). Collectively, these results imply expression divergence of most duplicate pairs and three members (i.e. CePIP1;1, -2;1, and -2;8) have evolved to be constitutively co-expressed in most tissues.

    Figure 3.  Expression profiles of CePIP genes in various tissues, different stages of leaf development, and mature leaves of diurnal fluctuation. (a) Tissue-specific expression profiles of 14 CePIP genes. The heatmap was generated using the R package implemented with a row-based standardization. Color scale represents FPKM normalized log2 transformed counts, where blue indicates low expression and red indicates high expression. (b) Expression profiles of CePIP1;1, -2;1, and -2;8 at different stages of leaf development. (c) Expression profiles of CePIP1;1, -2;1, and -2;8 in mature leaves of diurnal fluctuation. Bars indicate SD (N = 3) and uppercase letters indicate difference significance tested following Duncan's one-way multiple-range post hoc ANOVA (p< 0.01). (Ce: C. esculentus; FPKM: Fragments per kilobase of exon per million fragments mapped; PIP: plasma membrane intrinsic protein)

    As shown in Fig. 3a, compared with young leaves, transcriptome profiling showed that CePIP1;2, -2;3, -2;7, -2;8, and -2;11 were significantly down-regulated in mature leaves, whereas CePIP1;3 and -2;1 were up-regulated. To confirm the results, three dominant members, i.e., CePIP1;1, -2;1, and -2;8, were selected for qRT-PCR analysis, which includes three representative stages, i.e., young, mature, and senescing leaves. As shown in Fig. 3b, in contrast to CePIP2;1 that exhibited a bell-like expression pattern peaking in mature leaves, transcripts of both CePIP1;1 and -2;8 gradually decreased during leaf development. These results were largely consistent with transcriptome profiling, and the only difference is that CePIP1;1 was significantly down-regulated in mature leaves relative to young leaves. However, this may be due to different experiment conditions used, i.e., greenhouse vs natural conditions.

    Diurnal fluctuation expression patterns of CePIP1;1, -2;1, and -2;8 were also investigated in mature leaves and results are shown in Fig. 3c. Generally, transcripts of all three genes in the day (8, 12, 16, and 20 h) were higher than that in the night (24 and 4 h). During the day, both CePIP1;1 and -2;8 exhibited an unimodal expression pattern that peaked at 12 h, whereas CePIP2;1 possessed two peaks (8 and 16 h) and their difference was not significant. Nevertheless, transcripts of all three genes at 20 h (onset of night) were significantly lower than those at 8 h (onset of day) as well as 12 h. In the night, except for CePIP2;1, no significant difference was observed between the two stages for both CePIP1;1 and -2;8. Moreover, their transcripts were comparable to those at 20 h (Fig. 3c).

    To reveal the expression patterns of CePIP genes during tuber development, three representative stages, i.e., 40 DAS (early swelling stage), 85 DAS (late swelling stage), and 120 DAS (mature stage), were first profiled using transcriptome data. As shown in Fig. 4a, except for rare expression of CePIP1;2, -2;2, -2;9, and -2;10, most genes exhibited a bell-like expression pattern peaking at 85 DAS, in contrast to a gradual decrease of CePIP2;3 and -2;8. Notably, except for CePIP2;4, other genes were expressed considerably lower at 120 DAS than that at 40 DAS. For qRT-PCR confirmation of CePIP1;1, -2;1, and -2;8, seven stages were examined, i.e., 1, 5, 10, 15, 20, 25, and 35 DAI, which represent initiation, five stages of swelling, and maturation as described before[32]. As shown in Fig. 4b, two peaks were observed for all three genes, though their patterns were different. As for CePIP1;1, compared with the initiation stage (1 DAI), significant up-regulation was observed at the early swelling stage (5 DAI), followed by a gradual decrease except for the appearance of the second peak at 20 DAI, which is something different from transcriptome profiling. As for CePIP2;1, a sudden drop of transcripts first appeared at 5 DAI, then gradually increased until 20 DAI, which was followed by a gradual decrease at two late stages. The pattern of CePIP2;8 is similar to -1;1, two peaks appeared at 5 and 20 DAI and the second peak was significantly lower than the first. The difference is that the second peak of CePIP2;8 was significantly lower than the initiation stage. By contrast, the second peak (20 DAI) of CePIP2;1 was significantly higher than that of the first one (1 DAI). Nevertheless, the expression patterns of both CePIP2;1 and -2;8 are highly consistent with transcriptome profiling.

    Figure 4.  Transcript and protein abundances of CePIP genes during tuber development. (a) Transcriptome-based expression profiling of 14 CePIP genes during tuber development. The heatmap was generated using the R package implemented with a row-based standardization. Color scale represents FPKM normalized log2 transformed counts, where blue indicates low expression and red indicates high expression. (b) qRT-PCR-based expression profiling of CePIP1;1, -2;1, and -2;8 in seven representative stages of tuber development. (c) Relative protein abundance of CePIP1;1, -2;1, and -2;8 in three representative stages of tuber development. Bars indicate SD (N = 3) and uppercase letters indicate difference significance tested following Duncan's one-way multiple-range post hoc ANOVA (p < 0.01). (Ce: C. esculentus; DAI: days after tuber initiation; DAS: days after sowing; FPKM: Fragments per kilobase of exon per million fragments mapped; PIP: plasma membrane intrinsic protein).

    Since protein abundance is not always in agreement with the transcript level, protein profiles of three dominant members (i.e. CePIP1;1, -2;1, and -2;8) during tuber development were further investigated. For this purpose, we first took advantage of available proteomic data to identify CePIP proteins, i.e., leaves, roots, and four stages of tubers (freshly harvested, dried, rehydrated for 48 h, and sprouted). As shown in Supplemental Fig. S2, all three proteins were identified in both leaves and roots, whereas CePIP1;1 and -2;8 were also identified in at least one of four tested stages of tubers. Notably, all three proteins were considerably more abundant in roots, implying their key roles in root water balance.

    To further uncover their profiles during tuber development, 4D-PRM-based protein quantification was conducted in three representative stages of tuber development, i.e., 1, 25, and 35 DAI. As expected, all three proteins were identified and quantified. In contrast to gradual decrease of CePIP2;8, both CePIP1;1 and -2;1 exhibited a bell-like pattern that peaked at 25 DAI, though no significant difference was observed between 1 and 25 DAI (Fig. 4c). The trends are largely in accordance with their transcription patterns, though the reverse trend was observed for CePIP2;1 at two early stages (Fig. 4b & Fig. 4c).

    As predicted by WoLF PSORT, CePIP1;1, -2;1, and -2;8 may function in the cell membrane. To confirm the result, subcellular localization vectors named pNC-Cam1304-CePIP1;1, pNC-Cam1304-CePIP2;1, and pNC-Cam1304-CePIP2;8 were further constructed. When transiently overexpressed in tobacco leaves, green fluorescence signals of all three constructs were confined to cell membranes, highly coinciding with red fluorescence signals of the plasma membrane marker HbPIP2;3-RFP (Fig. 5).

    Figure 5.  (a) Schematic diagram of overexpressing constructs, (b) subcellular localization analysis of CePIP1;1, -2;1, and -2;8 in N. benthamiana leaves. (35S: cauliflower mosaic virus 35S RNA promoter; Ce: C. esculentus; EGFP: enhanced green fluorescent protein; kb: kilobase; NOS: terminator of the nopaline synthase gene; RFP: red fluorescent protein; PIP: plasma membrane intrinsic protein).

    Water balance is particularly important for cell metabolism and enlargement, plant growth and development, and stress responses[2,19]. As the name suggests, AQPs raised considerable interest for their high permeability to water, and plasma membrane-localized PIPs were proven to play key roles in transmembrane water transport between cells[1,18]. The first PIP was discovered in human erythrocytes, which was named CHIP28 or AQP1, and its homolog in plants was first characterized in Arabidopsis, which is known as RD28, PIP2c, or AtPIP2;3[3,7,53]. Thus far, genome-wide identification of PIP genes have been reported in a high number of plant species, including two model plants Arabidopsis and rice[10,11,1317,5456]. By contrast, little information is available on Cyperaceae, the third largest family within the monocot clade that possesses more than 5,600 species[57].

    Given the crucial roles of water balance for tuber development and crop production, in this study, tigernut, a representative Cyperaceae plant producing high amounts of oil in underground tubers[28,30,32], was employed to study PIP genes. A number of 14 PIP genes representing two phylogenetic groups (i.e., PIP1 and PIP2) or 12 orthogroups (i.e., PIP1A, PIP1B, PIP1C, PIP2A, PIP2B, PIP2C, PIP2D, PIP2E, PIP2F, PIP2G, PIP2H, and PIP2I) were identified from the tigernut genome. Though the family amounts are comparative or less than 13–21 present in Arabidopsis, cassava (Manihot esculenta), rubber tree (Hevea brasiliensis), poplar (Populus trichocarpa), C. cristatella, R. breviuscula, banana (Musa acuminata), maize (Zea mays), sorghum (Sorghum bicolor), barley (Hordeum vulgare), and switchgrass (Panicum virgatum), they are relatively more than four to 12 found in eelgrass (Zostera marina), Brachypodium distachyon, foxtail millet (Setaria italic), J. effuses, Aquilegia coerulea, papaya (Carica papaya), castor been (Ricinus communis), and physic nut (Jatropha curcas) (Supplemental Table S4). Among them, A. coerulea represents a basal eudicot that didn't experience the γ WGD shared by all core eudicots[50], whereas eelgrass is an early diverged aquatic monocot that didn't experience the τ WGD shared by all core monocots[56]. Interestingly, though both species possess two PIP1s and two PIP2s, they were shown to exhibit complex orthologous relationships of 1:1, 2:2, 1:0, and 0:1 (Supplemental Table S5). Whereas AcPIP1;1/AcPIP1;2/ZmPIP1;1/ZmPIP1;2 and ZmPIP2;1/AcPIP2;1 belong to PIP1A and PIP2A identified in this study, AcPIP2;2 and ZmPIP2;2 belong to PIP2H and PIP2I, respectively (Supplemental Table S5), implying that the last common ancestor of monocots and eudicots possesses only one PIP1 and two PIP2s followed by clade-specific expansion. A good example is the generation of AtPIP1;1 and -2;6 from AtPIP1;5 and -2;1 via the γ WGD, respectively[17].

    In tigernut, extensive expansion of the PIP subfamily was contributed by WGD (2), transposed (2), tandem (5), and dispersed duplications (3). It's worth noting that, two transposed repeats (i.e., CePIP1;1/-1;3 and CePIP2;1/-2;8) are shared by rice, implying their early origin that may be generated sometime after the split with the eudicot clade but before Cyperaceae-Poaceae divergence. By contrast, two WGD repeats (i.e., CePIP1;1/-1;2 and CePIP2;2/-2;4) are shared by C. cristatella, R. breviuscula, and J. effusus but not rice and Arabidopsis, implying that they may be derived from WGDs that occurred sometime after Cyperaceae-Poaceae split but before Cyperaceae-Juncaceae divergence. The possible WGD is the one that was described in C. littledalei[58], though the exact time still needs to be studied. Interestingly, compared with Arabidopsis (1) and rice (2), tandem/proximal duplications played a more important role in the expansion of PIP genes in tigernut (5) as well as other Cyperaceae species tested (5–6), which were shown to be Cyperaceae-specific or even species-specific. These tandem repeats may play a role in the adaptive evolution of Cyperaceae species as described in a high number of plant species[14,41]. According to comparative genomics analyses, tandem duplicates experienced stronger selective pressure than genes formed by other modes (WGD, transposed duplication, and dispersed duplication) and evolved toward biased functional roles involved in plant self-defense[41].

    As observed in most species such as Arabidopsis[10,1417], PIP genes in all Cyperaceae and Juncaceae species examined in this study, i.e., tigernut, C. cristatella, R. breviuscula, and J. effuses, feature three introns with conserved positions. By contrast, zero to three introns was not only found in rice but also in other Poaceae species such as maize, sorghum, foxtail millet, switchgrass, B. distachyon, and barley[54,55], implying lineage/species-specific evolution.

    Despite the extensive expansion of PIP genes (PIP2) in tigernut even after the split with R. breviuscula, CePIP1;1, -2;1, and -2;8 were shown to represent three dominant members in most tissues examined in this study, i.e., young leaf, mature leaf, sheath, rhizome, shoot apex, and tuber, though the situation in root is more complex. CePIP1;1 was characterized as a transposed repeat of CePIP1;3, which represents the most expressed member in root. Moreover, its recent WGD repeat CePIP1;2 was shown to be lowly expressed in most tested tissues, implying their divergence. The ortholog of CePIP1;1 in rice is OsPIP1;3 (RWC-3), which was shown to be preferentially expressed in roots, stems, and leaves, in contrast to constitutive expression of OsPIP1;1 (OsPIP1a) and -1;2[5961], two recent WGD repeats. Injecting the cRNA of OsPIP1;3 into Xenopus oocytes could increase the osmotic water permeability by 2–3 times[60], though the activity is considerably lower than PIP2s such as OsPIP2;2 and -2;2[6163]. Moreover, OsPIP1;3 was shown to play a role in drought avoidance in upland rice and its overexpression in lowland rice could increase root osmotic hydraulic conductivity, leaf water potential, and relative cumulative transpiration at the end of 10 h PEG treatment[64]. CePIP2;8 was characterized as a transposed repeat of CePIP2;1. Since their orthologs are present in both rice and Arabidopsis (Supplemental Table S3), the duplication event is more likely to occur sometime before monocot-eudicot split. Interestingly, their orthologs in rice, i.e., OsPIP2;1 (OsPIP2a) and -2;6, respectively, are also constitutively expressed[61], implying a conserved evolution with similar functions. When heterologously expressed in yeast, OsPIP2;1 was shown to exhibit high water transport activity[62,6466]. Moreover, root hydraulic conductivity was decreased by approximately four folds in OsPIP2;1 RNAi knock-down rice plants[64]. The water transport activity of OsPIP2;6 has not been tested, however, it was proven to be an H2O2 transporter that is involved in resistance to rice blast[61]. More work especially transgenic tests may improve our knowledge of the function of these key CePIP genes.

    Leaf is a photosynthetic organ that regulates water loss through transpiration. In tigernut, PIP transcripts in leaves were mainly contributed by CePIP1;1, -2;1, and -2;8, implying their key roles. During leaf development, in contrast to gradual decrease of CePIP1;1 and -2;8 transcripts in three stages (i.e. young, mature, and senescing) examined in this study, CePIP2;1 peaked in mature leaves. Their high abundance in young leaves is by cell elongation and enlargement at this stage, whereas upregulation of CePIP2;1 in mature leaves may inform its possible role in photosynthesis[67]. Thus far, a high number of CO2 permeable PIPs have been identified, e.g., AtPIP2;1, HvPIP2;1, HvPIP2;2, HvPIP2;3, HvPIP2;5, and SiPIP2;7[6870]. Moreover, in mature leaves, CePIP1;1, -2;1, and -2;8 were shown to exhibit an apparent diurnal fluctuation expression pattern that was expressed more in the day and usually peaked at noon, which reflects transpiration and the fact that PIP genes are usually induced by light[11,7173]. In rice, OsPIP2;4 and -2;5 also showed a clear diurnal fluctuation in roots that peaked at 3 h after the onset of light and dropped to a minimum 3 h after the onset of darkness[11]. Notably, further studies showed that temporal and dramatic induction of OsPIP2;5 around 2 h after light initiation was triggered by transpirational demand but not circadian rhythm[74].

    As an oil-bearing tuber crop, the main economic goal of tigernut cultivation is to harvest underground tubers, whose development is highly dependent on water available[32,75]. According to previous studies, the moisture content of immature tigernut tubers maintains more than 80.0%, followed by a seed-like dehydration process with a drop of water content to less than 50% during maturation[28,32]. Thereby, the water balance in developmental tubers must be tightly regulated. Like leaves, the majority of PIP transcripts in tubers were shown to be contributed by CePIP1;1, -2;1, and -2;8, which was further confirmed at the protein level. In accordance with the trend of water content during tuber development, mRNA, and protein abundances of CePIP1;1, -2;1, and -2;8 in initiation and swelling tubers were considerably higher than that at the mature stage. High abundances of CePIP1;1, -2;1, and -2;8 at the initiation stage reflects rapid cell division and elongation, whereas upregulation of CePIP1;1 and -2;1 at the swelling stage is in accordance with cell enlargement and active physiological metabolism such as rapid oil accumulation[28,30]. At the mature stage, downregulation of PIP transcripts and protein abundances resulted in a significant drop in the moisture content, which is accompanied by the significant accumulation of late embryogenesis-abundant proteins[23,32]. The situation is highly distinct from other tuber plants such as potato (Solanum tuberosum), which may contribute to the difference in desiccation resistance between two species[32,76]. It's worth noting that, in one study, CePIP2;1 was not detected in any of the four tested stages, i.e., freshly harvested, dried, rehydrated for 48 h, and sprouted tubers[23]. By contrast, it was quantified in all three stages of tuber development examined in this study, i.e., 1, 25, and 35 DAI (corresponding to freshly harvested tubers), which represent initiation, swelling, and maturation. One possible reason is that the protein abundance of CePIP2;1 in mature tubers is not high enough to be quantified by nanoLC-MS/MS, which is relatively less sensitive than 4D-PRM used in this study[30,46]. In fact, nanoLC-MS/MS-based proteomic analysis of 30 samples representing six tissues/stages only resulted in 2,257 distinct protein groups[23].

    Taken together, our results imply a key role of CePIP1;1, -2;1, and -2;8 in tuber water balance, however, the mechanism underlying needs to be further studied, e.g., posttranslational modifications, protein interaction patterns, and transcriptional regulators.

    To our knowledge, this is the first genome-wide characterization of PIP genes in tigernut, a representative Cyperaceae plant with oil-bearing tubers. Fourteen CePIP genes representing two phylogenetic groups or 12 orthogroups are relatively more than that present in two model plants rice and Arabidopsis, and gene expansion was mainly contributed by WGD and transposed/tandem duplications, some of which are lineage or even species-specific. Among these genes, CePIP1;1, -2;1, and -2;8 have evolved to be three dominant members that are constitutively expressed in most tissues, including leaf and tuber. Transcription of these three dominant members in leaves are subjected to development and diurnal regulation, whereas in tubers, their mRNA and protein abundances are positively correlated with the moisture content during tuber development. Moreover, their plasma membrane-localization was confirmed by subcellular localization analysis, implying that they may function in the cell membrane. These findings shall not only provide valuable information for further uncovering the mechanism of tuber water balance but also lay a solid foundation for genetic improvement by regulating these key PIP members in tigernut.

    The authors confirm contribution to the paper as follows: study conception and design, supervision: Zou Z; analysis and interpretation of results: Zou Z, Zheng Y, Xiao Y, Liu H, Huang J, Zhao Y; draft manuscript preparation: Zou Z, Zhao Y. All authors reviewed the results and approved the final version of the manuscript.

    All the relevant data is available within the published article.

    This work was supported by the Hainan Province Science and Technology Special Fund (ZDYF2024XDNY171 and ZDYF2024XDNY156), China; the National Natural Science Foundation of China (32460342, 31971688 and 31700580), China; the Project of Sanya Yazhou Bay Science and Technology City (SCKJ-JYRC-2022-66), China. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

  • The authors declare that they have no conflict of interest.

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  • Cite this article

    Sassine YN, Shuleva N, El Sebaaly Z. 2022. Changes in Pleurotus ostreatus nutritional value and heavy metal profile as a result of supplementation with nano-additive. Studies in Fungi 7:12 doi: 10.48130/SIF-2022-0012
    Sassine YN, Shuleva N, El Sebaaly Z. 2022. Changes in Pleurotus ostreatus nutritional value and heavy metal profile as a result of supplementation with nano-additive. Studies in Fungi 7:12 doi: 10.48130/SIF-2022-0012

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Changes in Pleurotus ostreatus nutritional value and heavy metal profile as a result of supplementation with nano-additive

Studies in Fungi  7 Article number: 12  (2022)  |  Cite this article

Abstract: The present study evaluated the effect of Lithovit-Amino25 on the nutrient profile and heavy metal composition of Pleurotus ostreatus. The product was tested in two doses applied at three different timings: T2: 3 g kg−1/spawning, T3: 3 g kg−1/after first harvest, T4: 3 g kg−1/spawning and after first harvest, T5: 5 g kg−1/spawning, T6: 5 g kg−1/after first harvest, and T7: 5 g kg−1/spawning and after first harvest. Compared to control (T1: non-treated substrate), mushrooms’ fibers and carbohydrates increased in all treatments, recording the highest values in T4 (4.16%) and T3 (18.42%), respectively. Protein content was higher in mushrooms of substrates treated at spawning, with a 0.33% improvement in T5. Fat content decreased in T3, T4, T6, and T5. Total sugars decreased in mushrooms of treated substrates, and glucose was the dominant sugar in mushrooms. Fructose increased in mushrooms of T3 and T4. Calcium, iron, and potassium decreased in mushrooms of treated substrates. Sodium decreased in T3, T5, and T7, magnesium increased in T2, and phosphorus increased only in T2 and T7. Copper content of all treated mushrooms was in the standard safe limit (< 40 ppm), and it decreased in T2, T4, and T5 by around 2.5, 6.6, and 5.1 ppm, compared to control. However, zinc content increased in mushrooms of all treated substrates, and nickel and lead, increased by respective ranges of 2.8−11.88 ppm and 9.1−21 ppm, higher than the safe limits. The product presented a risk of heavy metal bioaccumulation even with a low dose.

    • Nowadays, a large amount of agro-industrial wastes are annually abandoned in the environment without any pre-treatment[1,2]. These wastes could be incorporated in mushroom production as a method to reduce their negative impacts on the environment which result from their hazardous disposal[35]. For instance, enormous amounts of spent mushroom substrate (SMS) resulting from mushroom cropping are discharged hazardously and need to be managed[6]. In parallel, researchers have long been utilizing SMS in mushroom production[7,8] taking advantage of its richness in lignocellulosic materials, availability, and low cost[9]. It is a nutritious substrate containing considerable amounts of minerals[10], protein and carbohydrates[11], and contains good amounts of cellulose (40%) and hemicellulose (20%)[12].

      Pleurotus ostreatus ranks as the second most cultivated mushroom in the world[13]. It is valued because it is rich in protein, fibers, vitamins and (C, D and B-complex), and amino acids, and is low in calories[14,15]. The mushroom can utilize available lignocellulosic materials[16,17], such as the SMS. Pleurotus spp. can biodegrade SMS by producing the enzymes cellulases, hemicellulases, and ligninases[18]. The subsequent utilization of the growing substrate will result in an SMS poor in nutrients and proteins[19]. As a result, such substrates are commonly being amended with protein-rich additives to ameliorate its nutritional profile, thus ensuring higher production and quality of mushrooms[20]. Furthermore, amino acids can increase the performance of the mushroom[21]. An improvement in the biological yield of oyster mushroom was found after supplementing the SMS with 3 g kg1 of a nanometric size nitrogen additive (nano-amino) applied twice during the production cycle[22]. Further, the supplement type, dose, and application timing had a major impact on the nutritional composition and heavy metal profile of P. ostreatus mushroom when the SMS substrate was supplemented with nano-urea[8].

      Eventually, the substrate nutrient composition and properties are factors determining the mushroom nutritional composition and heavy metal profile[1,8,23]. Studying the mushrooms heavy metal profile as affected by recycled substrates and its subsequent impact on human health is taking researchers interest nowadays[24]. Consequently, the present study will showcase the effect of applying nano-amino with different doses and at separate timings during the cropping cycle on the nutrient composition and heavy metals profile of oyster mushroom cultivated on a substrate containing SMS.

    • The substrate used was a 1:1, w/w mixture of wheat straw and spent mushroom substrate. The latter was procured by 'Gourmet' farm (at Byblos, Lebanon) and was previously used to grow oyster mushroom. It was subjected to a sun-drying process for 1 week and then shopped for size reduction. Thereafter, the mixture of SMS and wheat straw was pasteurized for 8 h using hot water (60–65 °C) and then allowed to cool to 25 °C for spawning. The substrate properties determined by a series of analytical tests were as follows: C:N ratio = 43:1 (determined by CHN Carlo-Erba elemental analyzer, Model 1106, Italy), moisture content: 85.6% (by Moisture Analyzer), organic matter: 82.8% dry weight (by loss of ignition over 24 h at 430 °C), pH (1:5, w/v): 5.2 (by pH meter: UltraBasic-UB10 Denver Instrument, USA), total proteins: 7.5% dry weight (by Micro-Kjeldahl method using N × 6.25[25], and total carbohydrates: 30.5% dry weight (using the Anthrone method)[26].

    • Supplementation of the growing substrate applied a nitrogen-rich fertilizer (Lithovit-Amino25), containing 16 water-soluble vegetable l-amino acids, composed of calcium carbonate (50.0%), calcium oxide (28.0%), silicon dioxide (9.0%), total nitrogen (3.0%), magnesium oxide (1.8%), iron (0.5%), and manganese (0.02%)[27]. It was used in two separate doses and at three different timings. The experimental design was carried out for full factorial testing of the effect of two factors: product dose and timing of product application, through the following treatments: T2: 3 g kg−1/spawning, T3: 3 g kg−1 /after first harvest, T4: 3 g kg−1 /spawning and after first harvest, T5: 5 g kg−1/spawning, T6: 5 g kg−1/ after first harvest, and T7: 5 g kg−1/spawning and after first harvest. Each treatment was applied to 10 bags (10 replicates/treatment). To apply the nitrogenous additive at different timings, two solutions of two different concentrations (3 g L−1 and 5 g L−1) were prepared, and from each solution 0.5 l was sprayed on the substrate according to the corresponding treatment, keeping the substrate moisture content at 60%.

    • Spawning of the substrate was carried out at the 5% rate using a grain spawn of the strain M2175, procured from Mycelia Company (Deinze, Belgium)[22]. Polyethylene bags filled by the spawned substrates were then incubated at 25 °C in dark conditions. Inside the incubation room, relative humidity was maintained around 80%−90% by an ultrasonic mist maker (Hotsale 7 L h−1) throughout the incubation period (14 d). At the end of the vegetative growth phase, fruit induction was triggered by reducing CO2 levels (to 900–2,300 ppm by ventilation), lighting (using 200 lx light source), and cooling the growing room to a temperature of 15 °C. At this stage, the relative humidity was 88–90%. Regulation of room temperature and relative humidity during incubation and fruit induction applied a humidity-temperature meter (Lutron HT-3007SD).

    • Several analytical tests served for determining the mushrooms’ chemical composition, using 100 g of fresh mushrooms (pileus and stipe) of each treatment. The macro-Kjeldahl method was used to determine the total protein content with the conversion factor N × 4.38[28]. Total carbohydrates were determined using the Anthrone method[26]. Fiber analysis was carried out by applying the Weende technique[29]. After extracting a known weight of powdered mushroom sample with ethyl ether, the analysis of fat content was performed using Aldrich® Soxhlet extraction apparatus Z556203. Mushroom samples were boiled in water for 30 min to analyze soluble sugars, and 20 μl of the filtrate was then used for normal phase extraction using High Performance Liquid Chromatography at 30 °C (column NH2 column: 250 mm × 4.5 mm ID, flow rate of 1.2 ml min–1). Sugar identification applied a refractive index detector (RID), mobile phase: mixture of polar-non-polar solution, calibration: using a 2 point concentration), comparing with standards prepared from stock solution of sugars to get concentrations approximate to the sample. Mineral composition (Ca, Mg, K, Mg, Na, Fe, and Mn) was determined by adding 2.8 ml of HNO3 (65%) to 5–6 g of samples, digesting at 150 °C for 1 h, filtrating with 100 ml of distilled water, and subjecting the filtrate to Inductively Coupled Plasma Mass Spectrometry (ICP-MS). Phosphorus content was determined by spectrophotometry[30]. Nickel, copper, lead and zinc content were measured by Atomic Absorption Spectrophotometer (Perkin Elmer, Model Analyst 400, USA) after digesting the mushroom samples with a mixture of HNO3, H2SO4, and H2O2 (4:1:1) (12 ml per 1 g of sample). The mixture was then boiled at 150 °C for 4 h, and diluted to 25 ml with deionized water. Similarly, a blank digest was prepared. For calibration, standard solutions were prepared by diluting stock solutions (1,000 mg L−1; Sigma and Aldrich, Burlington, USA) of each metal.

      Fresh samples of mushrooms were used for the analysis and results (mean values of 3 replicates ± standard deviation) were converted and expressed as percentage dry weight.

    • Data analysis applied the One-way ANOVA and means were compared by Duncan’s multiple range test at p < 0.05 using SPSS25 program.

    • Results in Table 1 show that the product application resulted in a significant reduction in fat content for mushrooms of T3, T4, T6, and T7, but a significant increase in this component in mushrooms of T2 (by around 0.05% compared with control). Fiber and carbohydrates content increased significantly (p < 0.05) in the majority of treatments compared with control. The product applied in a dose 3 g kg−1 caused a higher increase in fiber content of mushrooms compared with the dose 5 g kg−1; this effect was especially pronounced with the lowest product dose applied twice (T4), causing the highest fiber content in mushrooms (4.16%). Improvement in carbohydrates content was the highest with 3 g kg−1 nano-amino applied after first harvest (T3: 18.42%), followed by that obtained with 5 g kg−1 applied at spawning (T5: 10.56%). Protein content recorded a significant improvement (0.33%) in mushrooms of T5 compared to control. Total sugars including glucose and sucrose were lower in mushrooms of substrates subjected to nano-amino application. However, there was a punctual increase of fructose in mushrooms obtained in substrates treated with 3 g kg−1 after first harvest (T3), and at both tested timings (T4).

      Table 1.  Composition (%fw) of P. ostreatus obtained from supplemented substrates.

      Fats
      Fiber
      Total
      carbohydrates
      Total
      proteins
      Total
      sugars
      Fructose
      Glucose
      Sucrose
      T10.16 ± 0.02d2.69 ± 0.25b4.36 ± 0.35a2.92 ± 0.13d0.18 ± 0.02e0.005 ± 0.00a0.17 ± 0.02d0.01 ± 0.00b
      T20.21 ± 0.01e3.56 ± 0.02d4.33 ± 0.07a2.95 ± 0.04d0.021 ± 0.00ab0.005 ± 0.00a0.011 ± 0.00ab0.005 ± 0.00a
      T30.05 ± 0.01a3.94 ± 0.02e18.42 ± 0.03e2.82 ± 0.03c0.047 ± 0.00d0.012 ± 0.00c0.03 ± 0.01c0.005 ± 0.00a
      T40.09 ± 0.02c4.16 ± 0.01f7.49 ± 0.02c2.24 ± 0.02a0.023 ± 0.00bc0.009 ± 0.00b0.009 ± 0.00ab0.005 ± 0.00a
      T50.17 ± 0.02d3.30 ± 0.16c10.56 ± 0.02d3.25 ± 0.01e0.021 ± 0.00ab0.005 ± 0.00a0.011 ± 0.00ab0.005 ± 0.00a
      T60.052 ± 0.01ab2.43 ± 0.02a7.10 ± 0.16b2.23 ± 0.01a0.025 ± 0.00c0.005 ± 0.00a0.015 ± 0.00b0.005 ± 0.00a
      T70.07 ± 0.02b2.81 ± 0.02b7.44 ± 0.01c2.64 ± 0.02b0.015 ± 0.00a0.005 ± 0.00a0.005 ± 0.00a0.005 ± 0.00a
      p-value
      Dose0.000.000.000.090.440.000.021.00
      Timing0.000.000.000.000.000.000.001.00
      Dose × Timing0.010.000.000.000.000.000.041.00
      T1: control, T2: 3 g kg−1/spawning, T3: 3 g kg−1/after first harvest, T4: 3 g kg−1/spawning and after first harvest, T5: 5 g kg−1/spawning, T6: 5 g kg−1/after first harvest, T7: 5 g kg−1/spawning and after first harvest. Means in the same column followed by different letters are significantly different at p < 0.05.

      Findings in Table 2 showed that although it is a good source of calcium and iron, nano-amino couldn’t increase these nutrients in produced mushrooms. On the contrary, calcium, iron, and also potassium was significantly (p < 0.05) lower in mushrooms obtained from treated substrates than in control. Further, mushrooms obtained in treated substrate had a manganese content comparable (T3, T4, T5, T6, and T7) or significantly lower (T2) than that of mushrooms obtained in control substrate. Sodium content decreased by 0.0015%, 0.0012% and 0.0010% were noted respectively in the treatments T3, T5, and T7. Overall, zinc content increased in mushrooms of all treated substrates. Magnesium content increased punctually in T2 by around 0.003% compared to control. Phosphorus content increased in T2 and T7, by 0.22% and 0.15% respectively.

      Table 2.  Mineral composition of P. ostreatus mushrooms cultivated on supplemented substrates.

      Ca (%fw)K (%fw)Mn (ppm)Fe (ppm)Na (%fw)Mg (%fw)P (%fw)Zn (ppm)
      T10.0036 ± 0.00d0.36 ± 0.04d1.4 ± 0.00b22.0 ± 0.00f0.0080 ± 0.00c0.020 ± 0.00e0.68 ± 0.01c42.50 ± 0.1a
      T20.0018 ± 0.00c0.37 ± 0.00d0.9 ± 0.00a14.0 ± 0.00a0.0083 ± 0.00c0.023 ± 0.00f0.90 ± 0.16d65.28 ± 0.1b
      T30.0012 ± >0.00b0.29 ± 0.00c1.3 ± 0.00b20.0 ± 0.00d0.0065 ± 0.00a0.015 ± 0.00b0.64 ± 0.01bc71.22 ± 0.1c
      T40.0010 ± 0.00a0.24 ± 0.00a1.3 ± 0.00b18.0 ± 0.00c0.010 ± 0.00d0.014 ± 0.00a0.62 ± 0.01bc72.18 ± 0.1cd
      T50.0018 ± 0.00c0.31 ± 0.00c1.1 ± 0.00ab16.0 ± 0.00b0.0068 ± 0.00ab0.016 ± 0.00c0.46 ± 0.02a46.30 ± 0.2a
      T60.0013 ± 0.00b0.26 ± 0.00b1.1 ± 0.00ab21.0 ± 0.00f0.0079 ± 0.00c0.015 ± 0.00b0.57 ± 0.02b83.10 ± 0.2e
      T70.0014 ± 0.00b0.30 ± 0.00c1.1 ± 0.00ab20.0 ± 0.00e0.0070 ± 0.00b0.017 ± 0.00d0.83 ± 0.02d76.48 ± 0.1d
      p-value
      Dose0.010.040.410.000.000.000.000.46
      Timing0.000.000.090.000.000.000.000.00
      Dose × Timing0.010.000.150.000.000.000.000.00
      T1: control, T2: 3 g kg−1/spawning, T3: 3 g kg−1/after first harvest, T4: 3 g kg−1/spawning and after first harvest, T5: 5 g kg−1/spawning, T6: 5 g kg−1/after first harvest, T7: 5 g kg−1/spawning and after first harvest. Means in the same column followed by different letters are significantly different at p < 0.05.

      The heavy metal analysis of mushrooms (Table 3) revealed that the copper content of all treated mushrooms met the WHO standard safe limit (2009) (< 40 ppm). Compared to control, copper content of T2, T4 and T5 mushrooms was reduced by around 2.5, 6.6, and 5.1 ppm respectively. Despite the dose and timing of supplementation, mushrooms of treated substrates had increasing and dramatic values of nickel and lead, which increased by respective ranges of 2.8−11.88 ppm (2.8−11.88 mg kg−1) and 9.1−21 ppm (9.1−21 mg kg−1).

      Table 3.  Heavy metals in P. ostreatus mushrooms cultivated on supplemented substrates.

      Copper (ppm)Nickel (ppm)Lead (ppm)
      T113.90±0.1de8.80±0.2a6.20±0.1a
      T211.38 ± 0.1c14.70 ± 0.2e15.30 ± 0.2b
      T313.28 ± 0.1d16.30 ± 0.2f18.70 ± 0.2d
      T47.28 ± 0.1a12.86 ± 0.1c19.78 ± 0.1e
      T58.76 ± 0.1b20.68 ± 0.1g18.00 ± 0.2cd
      T614.00 ± 0.2de11.60 ± 0.2b27.20 ± 0.1f
      T714.22 ± 0.1e14.28 ± 0.1d17.66 ± 0.1c
      p-value
      Dose0.000.000.00
      Timing0.000.000.00
      Dose × Timing0.000.000.00
      T1: control, T2: 3 g kg−1/spawning, T3: 3 g kg−1/after first harvest, T4: 3 g kg−1/spawning and after first harvest, T5: 5 g kg−1/spawning, T6: 5 g kg−1/after first harvest, T7: 5 g kg−1/spawning and after first harvest. Means in the same column followed by different letters are significantly different at p < 0.05.
    • Mushroom protein content was higher when substrates were supplemented at spawning rather than other timings. It could be that high nitrogen doses accumulating in the growing substrate could counteract the assimilation of amino acids and sugars from the substrates, causing a lower synthesis of proteins, sugars, and carbohydrates in mushrooms. The initial nitrogen level is a crucial factor for the microbiota development within the substrate[31]. When nitrogen is excessive in the substrate it plays a negative effect on the growth and development of the mycelium in the growing substrate. An increase of P. ostreatus protein content by 33.6% was found when the mushroom was cultivated on sugar cane bagasse supplemented with urea[32].

      The total carbohydrate content of the substrate usually decreases after the first harvest. This is because fungi consume them along with other nutrients during growth[33]. Therefore, the higher carbohydrate content contained in the mushrooms of T3 and T6 (supplementation after first harvest) in comparison with control cases could be explained by the fact that the product, with high nitrogen content, has boosted the degradation of the substrate lignocellulose hence facilitating carbohydrate metabolism. Improvement in mushroom carbohydrates could be linked to a better degradation of lignin in the growing substrate. The highest decrease in substrate lignin was reported after a low dose of nano-amino was applied at double timings during P. ostreatus production cycle[22].

      Further, to obtain the amino-acids from substrates, the mushroom needs first to degrade the substrate protein via extracellular enzymatic secretion. The mushroom could then synthesize proteins. The product applied in the present study is initially rich in amino-acids, providing a more easily available form of amino acids compared to those obtained after the biodegradation of substrate’ proteins. As a result, mushrooms treated with nano-amino at spawning had higher protein content than those treated at later stages of the production cycle. But, the double application of the product did not essentially ameliorate the protein synthesis in mushrooms, probably because of high nitrogen accumulating in the substrate and negatively affecting the mushroom growth and metabolism. Moreover, the application of nano-supplement (nano-urea) to spent mushroom substrate was reported to improve the protein content in produced mushrooms[8].

      Carbohydrate foods are important source of fiber, with positive physiological effects on human health[34]. In the human body, proteins and other nitrogenous compounds are constantly broken down and contribute to the amino acid/nitrogen pool, from which precursors and amino acids are reused to produce enzymes, hormones, immune- functioning proteins, and other essential compounds[35].

      Generally, mushrooms are known to have low total soluble sugar content[36]. As observed, glucose was the most abundant type of sugar found in produced mushrooms, but it was significantly reduced in all mushrooms of treated substrates compared to control. The substrate used to grow P. ostreatus is formed by wheat straw, containing around 36% cellulose, which, when broken down by the mushrooms’ enzymes, secretes simple sugars, like glucose[37]. A lower degradation of cellulose in substrates treated with nano-amino may have caused a lower assimilation of sugars from the substrates, causing lower sugar content in mushrooms. Nano-amino application caused a lower cellulose biodegradation[22]. Further, the sugar composition of P. ostreatus mushrooms obtained in the present study is close to that obtained after supplementing SMS with nano-urea[8].

      In general, the mineral composition of mushrooms is normally affected by the substrate’s mineral profile[38]. Also, the substrates' pH may affect the heavy metal bioaccumulation and favor the absorption of certain minerals at the expense of others[39]. The product applied, initially rich in calcium carbonate (CaCO3: 50%) could increase the substrate pH, resulting in a variable mineral profile of mushrooms obtained in the different treatments. The effects of nitrogen supplementation on mineral uptake levels is directly related to the substrate composition[32].

      Further, it is well known that high lignin decomposition by P. ostreatus could be linked to a high MnP liberation in the substrate[40]. This liberation may have enhanced the MnP enzymatic activity in treated substrates richer in Mn due to nano-amino application. Manganese content in treated substrates may have been completely used by MnP at the stage of mycelial run which inhibited its translocations to mushrooms in a further stage. The substrate supplemented at spawning with a product dose of 3 g kg−1 showed higher lignin degradation compared to non-treated substrate[22]. A reduction of food sodium content is favored for blood pressure control[1] (https://meadowmushrooms.co.nz/storage/wysiwyg/files/final-nutritional-analysis-of-meadow-mushrooms-a-summary.pdf).

      The competition between metals in soil affects the absorption of some of these metals by wild mushrooms[41]. This may suggest a serious metal competition occurring in treated substrates and favoring the absorption of zinc at the expense of calcium and iron from the substrates. P. ostreatus is rich in phosphorus and phosphorus-rich foods are good contributors in human nutrition[14]. However, high levels may inhibit calcium absorption causing weak bones, itchy skin, and joints pain that can lead to mineral bone disorders in chronic kidney disease[42]. Foods rich in protein and carbohydrates were associated with zinc accumulation[43]. In the current study, all treated mushrooms with nano-amino showed high zinc levels above the safe level (60 ppm) set by the WHO[44] except when the growing substrate was supplemented at spawning with a dose of 5 g kg−1 (T5).

      Heavy metal concentrations in edible and non-edible mushrooms are associated with mineral substrates or heavily contaminated areas such as large cities and industrial sites[45]. Oyster mushroom absorbs heavy metals from the substrate through its spacious mycelium[46]. Certain metals, such as Ca, Cu, Fe, K, Mg, Mn, Na, Ni, and Zn are biologically active in fungi[47]. High nickel levels in mushrooms could lead to serious toxicity[48]. Effectively, the levels of nickel detected in mushrooms from treated substrates were higher than the safe range of 0.05–5 ppm given for plant foods (https://nap.nationalacademies.org/read/20096/chapter/2 ). Moreover, lead content in control mushrooms was higher than that previously reported by Quarcoo & Adotey[46] (0.04 mg kg1) and the values recommended by the EU commission (https://eur-lex.europa.eu/LexUriServ/LexUriServ.do?uri=CONSLEG:2001R0466:20060701:EN:PDF) and WHO[44] (0.3 mg kg–1 and 2 mg kg–1, respectively). Ca, Fe, K, Mn, and P are required for normal human physiological function, but prolonged overexposure to Cu and Pb may cause neurological dysfunction or overt disease. High levels of Zn, Ni, and Cu are for instance neurotoxic and may lead to seizures[49]. The mushrooms’ heavy metals profile obtained in treated substrates is of concern. The exceptional reduction in Cu content in a few cases of treated mushrooms might have occurred because of the competition posed by Ni, Pb, and Zn, as suggested previously by Sassine et al.[8].

    • Investigating the nano-amino effect on P. ostreatus nutritional attributes showed that such a treatment could be beneficial causing a general improvement of proteins, carbohydrates, and fiber content, reduction in total sugars, coupled with punctual phosphorus increase and sodium decrease in mushrooms. Using the product once in low or high dose seems to be more advantageous than twice for carbohydrates and protein metabolism. However, even with the lowest dose applied, a risk of nickel and lead accumulation was observed suggesting that the product may have been better tested in lower doses.

      • The authors declare that they have no conflict of interest.

      • Copyright: © 2022 by the author(s). Published by Maximum Academic Press, Fayetteville, GA. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
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    Sassine YN, Shuleva N, El Sebaaly Z. 2022. Changes in Pleurotus ostreatus nutritional value and heavy metal profile as a result of supplementation with nano-additive. Studies in Fungi 7:12 doi: 10.48130/SIF-2022-0012
    Sassine YN, Shuleva N, El Sebaaly Z. 2022. Changes in Pleurotus ostreatus nutritional value and heavy metal profile as a result of supplementation with nano-additive. Studies in Fungi 7:12 doi: 10.48130/SIF-2022-0012

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