Abstract

   In mature pollen grains, lipids are primarily stored in the form of
   lipid droplets that provide energy and act as a carbon source for
   normal pollen development and germination. Triacylglycerol (TAG) is the
   major form of stored plant lipids. Diacylglycerol transferase, which is
   encoded by DGAT1 in Arabidopsis thaliana, is an important enzyme
   regulating triacylglycerol synthesis. Within the seeds of the DGAT1
   mutant as11, the content of TAG is significantly decreased and the
   fatty acid composition also differs from the wild type. Transcriptome
   data of mature anthers showed that the genes involved in the TAG
   synthesis pathway were downregulated in as11. Analysis of gene
   expression patterns via transcriptome data also revealed that the
   expression of PDAT1, which functions in a manner complementary to the
   DGAT1 gene, was significantly decreased in as11, whereas the
   amylopectin synthase genes SS1 and SS2 were upregulated in mutant as11.
   We also detected lower total lipid, TAG and fatty acid contents in
   mature as11 pollen, with palmitic acid (C16:0) and linolenic acid
   (C18:3) being the major fatty acids in mature pollen. The cytological
   results showed that the lipid droplet content was reduced in mature
   as11 pollen. In the binuclear pollen grain II stage, WT pollen
   contained lipid droplets that were primarily accumulated around the
   generative nucleus, whereas the pollen in the mutant as11 was rich in
   starch grains that were primarily distributed around the vegetative
   nucleus. Ultrastructural analysis indicated that during pollen
   development in as11, the amount of endoplasmic reticulum in tapetal
   cells and pollen grains decreased, whereas the Golgi body content
   increased, which directly or indirectly led to a decrease in the levels
   of lipidosomes and an increase in the starch content in as11. Changes
   in the lipid content and fatty acid composition of the mutant as11
   differ from those in the wild type during pollen development.

   Keywords: Arabidopsis thaliana, pollen development, TAG, tapetum
     __________________________________________________________________

   Diacylglycerol transferase, which is encoded by DGAT1 in Arabidopsis
   thaliana, is an important enzyme that regulates triacylglycerol
   synthesis. Transcriptome data of mature anthers showed that the genes
   involved in the TAG synthesis pathway were downregulated in DGAT1
   mutant as11. The total lipid, TAG and fatty acid contents were reduced
   in as11 mature pollen, with palmitic acid (C16:0) and linolenic acid
   (C18:3) being the major fatty acids in mature pollen. The cytological
   results showed that the lipid droplet content was lower in mature as11
   pollen compared to the wild type.

Introduction

   Pollen development in higher plants is a complicated process that
   involves the synthesis and secretion of nutriments such as
   carbohydrates, lipids, and others to provide nutrition for subsequent
   pollen development ([30]Liu et al., 2011). Tapetal cells can also
   synthesize and secrete different nutriments at various stages
   throughout development to allow for the nourishment of pollen grains
   ([31]Zhu et al., 2011; [32]Chen et al., 2013). Typically, a large
   number of nutriments are accumulated within microspores at the
   binuclear pollen grain stage, and these nutriments are primarily
   lipids, storage proteins, and/or carbohydrates ([33]Calzoni et al.,
   1990). For example, during pollen development in Lilium, a large number
   of starch grains are present within the vegetative cells at the early
   binuclear pollen grain stage. However, they begin to store lipid
   droplets as the primary nutriment in mature pollen ([34]Clément et al.,
   1994). Interestingly, during pollen development in tomato and Asian
   lotus, the opposite phenomenon is observed, with more starch grains
   than lipid droplets in mature pollen ([35]Zhu and Tian, 2015; [36]Zhang
   et al., 2019). However, during pollen development in the Hong Kong
   orchid tree, lipid droplets are present at the early stage, the lipid
   droplet and starch grain contents increase at the late binuclear pollen
   stage, and mature pollen grains store both lipid droplets and starch
   grains, but with the lipid content being dominant ([37]Zheng et al.,
   2017). At the binuclear pollen grain stage in Chinese cabbage, lipid
   droplets are abundant in both the tapetum and the pollen ([38]Xie et
   al., 2005). Additionally, the transport and transformation of
   nutriments are both directly related to pollen fertility. The innermost
   layer of anther parietal cells (the tapetum) will select, transform and
   transport nutriments entering the anther locule. Therefore, the
   abnormal accumulation of nutriments within the pollen and tapetum may
   result in pollen abortion ([39]Xie et al., 2005; [40]Xu et al., 2009;
   [41]Zhang et al., 2009; [42]Yang et al., 2017).

   Lipids and lipid derivatives are important components of pollen grains,
   particularly the mature pollen of most entomophilous flowers, and the
   lipid content can reach up to 25% of their dry weight ([43]Murphy,
   2001). Lipids that are accumulated in mature pollen can be divided into
   two categories. In the first category, the main components are phenolic
   substances and lipids. These are primarily composed of fatty acids and
   carotenoids that are synthesized and secreted by the tapetum and
   released into the anther locule to cover and deposit onto the extine to
   form the tryphine ([44]Ischebeck, 2016). The second category includes
   intracellular self-synthesized lipids that are formed during pollen
   development and is primarily comprised of polar lipids that are related
   to membrane synthesis and neutral lipids that are stored in lipid
   droplets ([45]Rodriguez-García et al., 2003; [46]Ischebeck, 2016).
   Lipid droplets in mature pollen possess one unique and relatively
   simple structure and are similar to oil bodies that are stored in plant
   seeds. Triacylglycerol (TAG) and sterol ester are the main components,
   and the outside is covered by a phospholipid monolayer membrane that is
   inlaid with a variety of proteins ([47]Thiam et al., 2013; [48]Ohsaki
   et al., 2014). There are two sites in plants where lipid droplets are
   synthesized (chloroplasts and endoplasmic reticulum), and lipid
   droplets in mature pollen are primarily synthesized by the endoplasmic
   reticulum ([49]Pyc et al., 2017). During late pollen development, lipid
   droplets are surrounded by a large number of endoplasmic reticulum
   structures to ensure that the lipid droplets are prevented from fusing
   with each other during pollen rehydration without the protection of
   oleosin ([50]Murphy, 2001). TAG is stored in large amounts within lipid
   droplets and acts as the primary source for providing energy and as an
   inert intermediate that regulates lipid synthesis ([51]Ziekiewicz et
   al., 2013). Therefore, TAG synthesis is essential for pollen
   development ([52]Zhang et al., 2009; [53]Ischebeck, 2016).

   In addition to its content within pollen, TAG exists extensively in
   plant seeds and fruits as the most primary storage form of plant oil
   ([54]Lu et al., 2003; [55]Saha et al., 2006), where it provides a
   carbon source and the nutriments necessary for seed germination and
   development ([56]Routaboul et al., 1999; [57]Zou et al., 1999; [58]Baud
   et al., 2002; [59]O’Neill et al., 2003; [60]Zheng et al., 2003;
   [61]Lung and Weselake, 2006). Diacylglycerol transacylase (DGAT) is
   encoded by the DGAT1 gene in Arabidopsis thaliana and is an important
   enzyme that catalyzes the synthesis of TAG. Compared to the wild type
   (WT), seeds from the DGAT1 gene mutant as11 exhibit a clear delay in
   development ([62]Katavic et al., 1995; [63]Hobbs et al., 1999;
   [64]Routaboul et al., 1999; [65]Zou et al., 1999; [66]Jako et al.,
   2001; [67]Lu and Hills, 2002). The DGAT1 gene mutant as11 slows DAG
   accumulation, which lowers the TAG/DAG ratio ([68]Katavic et al., 1995;
   [69]Hobbs et al., 1999; [70]Routaboul et al., 1999; [71]Zou et al.,
   1999; [72]Jako et al., 2001). The seed oil content of as11 is decreased
   by 25–35% relative to the WT, and the fatty acid composition is altered
   with an increased linolenic acid (C18:3) content ([73]Katavic et al.,
   1995; [74]Zou et al., 1999; [75]Lu and Hills, 2002). Although the oil
   content in mature as11 seeds differs from the WT, there is no
   significant difference between as11 and WT in regard to morphology
   ([76]Katavic et al., 1995; [77]Zou et al., 1999; [78]Lu and Hills,
   2002), dry weight, fresh weight and protein levels ([79]Katavic et al.,
   1995). However, as11 seeds show delayed germination, which can be
   recovered by treatment with a high concentration of mannite
   ([80]Routaboul et al., 1999; [81]Lu and Hills, 2002). Additionally, the
   sucrose content in dry seeds of as11 was higher than in the WT, and
   increased and then decreased after absorbing water and finally fell to
   the same level as that of the WT ([82]Lu and Hills, 2002).

   In recent decades, the majority of studies examining the DGAT1 mutant
   as11 have primarily focused on genetics, molecular biology, physiology
   and biochemistry ([83]Katavic et al., 1995; [84]Hobbs et al., 1999;
   [85]Routaboul et al., 1999; [86]Zou et al., 1999; [87]Jako et al.,
   2001; [88]Lu et al., 2003; [89]Zhang et al., 2009; [90]Xu et al.,
   2012). Studies on the total lipid and TAG content during pollen
   development in as11 are rare. Our transcriptional analysis found that
   the expression levels of key genes in the TAG synthesis pathway were
   downregulated in mature pollen, and the amount of endoplasmic reticulum
   (which indirectly influences the lipidosome content in the tapetum) was
   different in the mutant, ultimately resulting in a decrease in the
   total lipid and TAG contents in the mature pollen and in changes in
   fatty acid composition and expression of genes related to lipid
   synthesis during pollen development.

Methods

Plant material

   Seeds of A. thaliana ecotype Columbia and DGAT1 mutant as11 were sown
   onto damp nutrient soil and cold treated for at least 2 days at 4 °C.
   Then seedlings were placed into an incubator with illumination for 16 h
   and darkness for 8 h per 24-h cycle. Plants with normal growth and
   development were used for the experiments.

Thin section and histochemical staining

   Flower buds at different developmental stages were harvested and
   processed according to a previously described method, with a few
   modifications ([91]Zhang et al., 2002). Semithin sections (1 μm thick)
   were cut using a Leica RM2255 microtome, stained with PAS and Sudan
   Black B, and then observed and photographed under a Leica DMLB.
   Ultrathin sections (60–70 nm) were cut using a Leica UC6
   ultramicrotome, stained with uranyl acetate and lead citrate, and
   observed and photographed under a Philips FEI-TECNAI 12 transmission
   electron microscope.

Total lipid and TAG quantification and total fatty acid profiles

   Mature anthers were harvested in glass tubes on dry ice and weighed
   after freeze drying. Total lipid was extracted using the
   chloroform/methanol method described in the Acyl-Lipid Metabolism
   Chapter of [92]The Arabidopsis Book (2013). TAG was fractionated by TLC
   (Silica gel 60, Merck, USA) in developing solvent (hexane:
   diethylether: acetic acid = 70: 30: 1 (v/v/v)) and visualized by iodine
   vapor. Spots were scraped for following fatty acid methyl esters (FAME)
   with TAG standard (C15:0) to quantify the content of TAG.

   FAME of TAG and anthers at the VM (vacuolate microspore), BM (binuclear
   pollen grain), SMD (second mitotic division) and MP (mature pollen
   grain) stages were produced by a direct acid-catalyzed transmethylation
   protocol presented in the Acyl-Lipid Metabolism Chapter of [93]The
   Arabidopsis Book (2013), and analyzed by GC-MS (7890A-5975C, Agilent
   Technologies, USA). GC conditions were as follows—splitless mode
   injection, injector, and flame ionization detector (FID) temperature,
   250 °C; oven temperature program—100 °C for 1 min, 15 °C min^–1 to 175
   °C for 12 min, then 2 °C min^–1 to 245 °C, holding this temperature for
   2 min.

RNA isolation and qRT-PCR

   Total RNA of anthers from the VM, BM, SMD and MP stages was extracted
   by RNA FAST200 (Fastagen, Shanghai, China). Reverse transcription was
   completed using a PrimeScript™ 1st Strand cDNA Synthesis Kit (Takara,
   Dalian, China). Expression of genes selected according to TAG synthesis
   ([94]http://aralip.plantbiology.msu.edu/pathways/triacylglycerol_biosyn
   thesis) was determined by 2× SYBR Premix Ex Taq with a CFX96 Real-Time
   PCR Detection System (Bio-Rad Laboratories, Inc., USA). Arabidopsis
   thaliana Actin2 gene was used as the housekeeping gene. Three
   biological replicates were done for each gene. The primers are listed
   in [95]Supporting Information—Table S1.

Library preparation for transcriptome sequencing

   Total mature anther RNA for each sample (3 biological replicates) was
   extracted using the TRIzol method (TIANGEN BIOTECH, Beijing) and
   treated with RNase-free DNase I (TaKaRa). RNA degradation and
   contamination was monitored on 1% agarose gels. RNA was quantified
   using an Agilent 2100 Bioanalyzer (Agilent Technologies, CA, USA), and
   its quality and integrity were assessed using a NanoDrop
   spectrophotometer (IMPLEN, CA, USA). A total of 1.5 μg RNA per sample
   was used as input material for the RNA sample preparations. Sequencing
   libraries were generated using the NEBNext^® Ultra™ RNA Library Prep
   Kit for Illumina® (NEB, USA) following the manufacturer’s
   recommendations, and index codes were added to attribute sequences to
   each sample. Briefly, mRNA was purified from total RNA using poly-T
   oligo-attached magnetic beads. Fragmentation was carried out using
   divalent cations under elevated temperature in NEBNext First Strand
   Synthesis Reaction Buffer (5×). First-strand cDNA was synthesized using
   random hexamer primers and M-MuLV Reverse Transcriptase (RNase H).
   Second-strand cDNA synthesis was subsequently performed using DNA
   Polymerase I and RNase H. Remaining overhangs were converted into blunt
   ends via exonuclease/polymerase activity. After adenylation of the 3ʹ
   ends of DNA fragments, NEBNext Adaptor with a hairpin loop structure
   was ligated to prepare for hybridization. In order to select cDNA
   fragments of preferentially 200–250 bp in length, the library fragments
   were purified with the AMPure XP system (Beckman Coulter, Beverly,
   USA). Then 3 μL USER Enzyme (NEB, USA) was used with size-selected,
   adaptor-ligated cDNA at 37 °C for 15 min followed by 5 min at 95 °C
   before PCR. Then PCR was performed with Phusion High-Fidelity DNA
   polymerase, universal PCR primers and Index (X) Primer. Finally, PCR
   products were purified (AMPure XP system) and library quality was
   assessed on the Agilent Bioanalyzer 2100 system. The library
   preparations were sequenced on an Illumina Hiseq 4000 platform by the
   Beijing Allwegene Technology Company Limited (Beijing, China) and
   paired-end 150 bp reads were generated.

Quality control

   Raw data (raw reads) of fastq format were firstly processed through
   in-house perl scripts. In this step, clean data (clean reads) were
   obtained by removing reads containing adapter, reads containing ploy-N
   and low-quality reads from raw data. At the same time, Q20, Q30,
   GC-content and sequence duplication level of the clean data were
   calculated. All the downstream analyses were based on clean data with
   high quality.

Quantification of gene expression levels

   HTSeq v 0.5.4 p3 was used to count the read numbers mapped to each
   gene. Gene expression levels were estimated by fragments per kilobase
   of transcript per million fragments mapped (FPKM).

2.8 Differential expression analysis

   Differential expression analysis of two groups was performed using the
   DESeq R package (1.10.1). DESeq provides statistical routines for
   determining differential expression in digital gene expression data
   using a model based on the negative binomial distribution. The
   resulting P values were adjusted using the Benjamini and Hochberg’s
   approach for controlling the false discovery rate. Genes with an
   adjusted P-value < 0.05 found by DESeq were classified as
   differentially expressed.

2.9 GO enrichment analysis

   Gene Ontology (GO) enrichment analysis of the differentially expressed
   genes (DEGs) was implemented by the GOseq R packages based Wallenius
   non-central hyper-geometric distribution ([96]Young et al., 2010),
   which can adjust for gene length bias in DEGs.

2.10 KEGG pathway enrichment analysis

   KEGG ([97]Kanehisa et al., 2008) is a database resource for
   understanding high-level functions and utilities of biological systems,
   such as the cell, the organism and the ecosystem, based on
   molecular-level information, especially large-scale molecular datasets
   generated by genome sequencing and other high-throughput experimental
   technologies ([98]http://www.genome.jp/kegg/). We used KOBAS ([99]Mao
   et al., 2005) software to test the statistical enrichment of
   differentially expressed genes in KEGG pathways.

2.11 Data analysis

   Figures were plotted by Origin2018 (Microcal Software Inc.,
   Northampton, MA). All histograms were processed with Excel (Microsoft,
   Seattle, WA). The data were statistically analyzed using SPSS version
   20.0 (SPSS Inc., Chicago, IL). One-way ANOVA (Duncan’s multiple range
   test) was performed and differences were considered significant at p <
   0.05.

3 Results

3.1 Transcriptome analysis revealed candidate genes involved in TAG
biosynthesis

   Using the Illumina HiSeq4000 sequencing platform and PE150 sequencing
   strategy, 141,560,394 raw read pairings were measured and 139,274,010
   clean read pairings were obtained after quality control. A total of
   10,988 differential genes were identified, of which 5,175 were
   upregulated and 5,813 were downregulated in as11 ([100]Fig. 1). GO
   enrichment analysis of differentially expressed genes resulted in
   10,562 genes being annotated into 30 subclasses of two GO classes:
   biological processes and cellular components. A total of 6573 DEGs were
   identified as being involved in cellular processes in the biological
   process domain, and 7910 and 7920 DEGs were identified as being
   involved in cell parts and cells in the cellular component
   morphogenesis domain respectively. KEGG enrichment analysis of
   differentially expressed genes identified 2109 DEGs. In as11, 2105 DEGs
   were upregulated and 704 DEGs were downregulated. They were mainly
   enriched in metabolic pathways (383 upregulated, 460 downregulated)
   such as biosynthesis of secondary metabolism, carbon metabolism,
   biosynthesis of amino acids, fatty acid metabolism and fatty acid
   degradation (fatty acid metabolism) [[101]see Supporting
   Information—Fig. S1]. Forty-eight of the 69 related genes identified in
   the fatty acid metabolism pathway were differentially expressed, and 41
   of the 60 genes identified in the glycerolipid metabolism pathway were
   differentially expressed. Among the 43 genes identified in the fatty
   acid biosynthesis pathway, 28 were differentially expressed. According
   to the TAG synthesis pathway ([102]Fig. 2), 8 key genes with changes in
   expression were identified ([103]Table 1), and subsequent studies of
   these were carried out.

Fig. 1.

   [104]Fig. 1.
   [105]Open in a new tab

   Differentially expressed genes in WT and as11. Volcano plot of DEGs
   between WT and as11. Each dot in the volcano plot represents a gene;
   the abscissa represents the logarithm of the difference multiple of
   gene expression between the WT and as11 lines, and the ordinate
   represents the negative logarithm of statistically significant changes
   in gene expression. The larger the absolute value of the abscissa, the
   greater the multiple difference of expression quantity between the two
   lines; the larger the vertical coordinate value, the more significant
   the differential expression level, and the more reliable the selected
   DEGs. The green dots represent downregulated DEGs, the red dots
   represent upregulated DEGs, and the blue dots represent no DEGs. In
   comparing the gene expression in the WT and mutant, 10988 DEGs were
   consistently expressed in both the WT and mutant, 5175 were upregulated
   and 5813 were downregulated in the mutant.

Fig. 2.

   [106]Fig. 2.
   [107]Open in a new tab

   Overview of the TAG synthesis pathways in Arabidopsis. The genes marked
   in red were chosen for the expression profile analysis, and were
   downregulated in as11 according to the RNA-seq data. DHAP,
   dihydroxyacetone phosphate; GPDH, glycerol-3-phosphate dehydrogenase;
   G3P, glycerol-3-phosphate; GPAT, glycerol-3-phosphate acyltransferase;
   LPA, lysophosphatidic acid; LPAAT, lysophosphatidic acid
   acyltransferase; PA, phosphatidic acid; PAP, phosphatidic acid
   phosphatase; DAG, diacylglycerol; DGAT, diacylglycerol acyltransferase;
   CPT, CDP-choline: diacylglycerol cholinephosphotransferase; PDCT,
   phosphatidylcholine: diacylglycerol cholinephosphotransferase; PLC,
   phospholipase C; PLD, phospholipase D. PDAT,
   phospholipid:diacylglycerol acyltransferase; TAG, triacylglycerol.

Table 1.

   Genes with changes of glycerolipid etabolism in transcriptomics.
   DEGs EC No. Gene Description log2F PValue Express differences
   AT2G41540 1.1.1.8 Glycerol-3-phosphate dehydrogenase [NAD(+)] GPDHC1,
   cytosolic (GPDHc1) -0.4291 2.86E-23 Down
   AT1G06520 2.3.1.15 Glycerol-3-phosphate acyltransferase 1 (GPAT1)
   -0.54064 2.24E-25 Down
   AT2G38110 2.3.1.15 Glycerol-3-phosphate 2-O-acyltransferase 6 (GPAT6)
   -0.98036 1.45E-137 Down
   AT5G60620 2.3.1.15 GPAT9 -0.49674 1.05E-11 Down
   AT5G42870 3.1.3.4 Phosphatidate phosphatase PAH2 (PAH2) -0.31911
   3.12E-07 Down
   AT5G13640 2.3.1.158 Phospholipid:diacylglycerol acyltransferase 1
   (PDAT1) -0.49734 8.15E-18 Down
   AT3G12120 1.14.19.6
   1.14.19.22 FAD2 -0.20497 1.72E-08 Down
   AT1G74960 2.3.1.179 3-oxoacyl-[acyl-carrier-protein] synthase II,
   chloroplastic (FAB1) -0.23545 5.45E-06 Down
   AT5G24300 2.4.1.21 Starch synthase, chloroplastic/amyloplastic (SS1)
   0.23372 0.0021172 Up
   AT3G01180 2.4.1.21 Starch synthase, chloroplastic/amyloplastic (SS2)
   0.19539 0.0029552 Up
   [108]Open in a new tab

3.2 The lower expression of genes related to lipid synthesis and content of
total lipid and TAG in mature as11 pollen

   Based on transcriptome data, we next tested the expression of lipid
   synthesis related genes of wild type and mutant as11 in mature pollen
   grains. Our results indicated that at the mature pollen grain stage,
   the expression of GPAT (GPAT1, GPAT6, and GPAT9) and PDAT1, which are
   involved in seed TAG synthesis, was downregulated in as11 ([109]Fig.
   3). The GPDHc1 gene, which regulates the transition from
   glycometabolism to lipid metabolism, was expressed at a higher level in
   the WT ([110]Fig. 3). The expression analysis of the key genes related
   to fatty acid synthesis revealed that the expression of FAB1, which
   catalyzes the conversion of C16:0-ACP to C18:0-ACP, and the key enzyme
   gene FAD2, which can catalyze the production of linoleic acid (C18:2)
   was distinctly upregulated in the WT ([111]Fig. 3). However, it is
   interesting that the expression of starch synthetase genes,
   particularly the SS1 and SS2 genes that are primarily responsible for
   the regulation of amylopectin synthesis, was higher in as11 ([112]Fig.
   3).

Fig. 3.

   [113]Fig. 3.
   [114]Open in a new tab

   The relative expression of genes in the TAG synthesis pathway in the
   mature pollen grains. Significant differences are indicated by *P <
   0.05 and **P < 0.01. Bars show the standard error, except when it was
   too small to be visible.

   Based on the lower expression of lipid synthesis related genes in as11,
   we further determined the content of total lipid and TAG in mature
   pollen grains of the WT and as11. The total lipid in mature pollen
   grains of as11 was 26% less than that in the WT ([115]Fig. 4A). The TAG
   content of the total lipid was also significantly lower in as11
   ([116]Fig. 4B). As a result, the total content of TAG in mature as11
   pollen grains was distinctly lower than that in the WT ([117]Fig. 4C).
   We next measured the fatty acid composition of the mature WT and as11
   pollen grains, and found no significant difference in content
   ([118]Fig. 4D). Palmitic acid (C16:0) and linolenic acid (C18:3) were
   the main fatty acids in mature pollen in both the WT and as11. The
   content of linoleic acid (C18:2) and linolenic acid (C18:3) was a
   little higher in the WT. The stearic acid (C18:0) content followed the
   inverse pattern.

Fig. 4.

   [119]Fig. 4.
   [120]Open in a new tab

   Comparison of total lipid, TAG levels and relative content of total
   fatty acid levels between anthers of the WT and as11. A. The content of
   total lipid. Based on 100 mg freeze-dried mature anthers, the content
   of total fat in WT mature anthers was 25.71 mg, and that of as11 was
   18.88 mg, 26.57% lower than that of the WT. B. The TAG of total lipid.
   1 mg of the total lipid was separated by TLC, and GC-MS was used to
   calculate the TAG content: 6.60 μg/mg (TAG/total lipid) in WT mature
   anthers and 3.48 μg/mg in as11 anthers. Compared with the WT, the
   content was 47.21% lower in the mutant. C. The total TAG in the anthers
   of the WT and as11. The content of TAG in the WT was 169.60 μg per 100
   mg of freeze-dried mature anthers, while the content of TAG in as11 was
   65.32 μg, a decrease of 61.48% compared with that in the WT. D. The
   relative content of total fatty acid levels at the mature pollen grain
   stage. Significant differences are indicated by *P < 0.05 and **P <
   0.01. Three biological replicates were performed.

   We also measured the fatty acid composition of wild type and mutant
   as11 pollen grains at the VM (vacuolate microspore), BN (binuclear
   pollen grain), and SMD (second mitotic division) stages, and found that
   the content of linolenic acid (C18:3) was much higher in as11 than in
   the WT at the SMD stage, presenting a ‘trade-off’ with linoleic acid
   (C18:2), palmitic acid (C16:0) and stearic acid (C18:0). The relative
   content of fatty acids at the VM stage was used as the standard to
   obtain the relative ratio of palmitic acid (C16:0), stearic acid
   (C18:0) and linolenic acid (C18:3) at four developmental stages. We
   found that at the SMD stage, the ratios of palmitic acid (C16:0) and
   stearic acid (C18:0) were lower in as11 than in the WT ([121]Fig. S3).
   Accordingly, the ratio of linolenic acid (C18:3) was significantly
   higher in as11 than in the WT at the same stage. When the grains were
   mature, the ratio of the three fatty acids in both recovered the same
   between WT and as11.

3.3 The pollen ultrastructure changed during as11 pollen development

   Based on the developmental characteristics of the tapetum and pollen
   grains in the WT (Col ecotype) and according to the 13 growth stages of
   the entire developmental period in the wild-type Arabidopsis thaliana
   ecotype Wassilewskija ([122]Owen and Makaroff, 1995), we divided the
   mature pollen grain stages of Arabidopsis thaliana ecotype Columbia
   into mature I and mature II stages, giving a total of 14 growth stages.
   The growth stages of mutant as11 were divided on the basis of pollen
   development.

   Histochemical staining results indicated that there was no significant
   difference in the accumulation and distribution of nutrients during
   pollen development in either the WT or as11 from the pre-meiosis stage
   to the released microspore stage ([123]Fig. S2). At the vacuolate
   microspore stage, black granular lipid could be observed only in the
   tapetum of the WT ([124]Fig. 5A). At the binuclear pollen grain II
   stage, starch grains surrounded the vegetative nucleus in both the WT
   and as11 according to PAS staining ([125]Fig. 5B and [126]b); however,
   the mutant exhibited more starch accumulation. After counterstaining
   with Sudan Black B, there was obvious lipid accumulation in both the
   tapetum and the pollen grains of the WT and as11, and lipid droplets
   within the pollen grains were predominantly distributed around the
   generative nucleus ([127]Fig. 5C and [128]c). Furthermore, the content
   of lipid droplets within WT pollen was significantly higher than that
   in as11. When pollen grains developed to the second mitosis I stage,
   the number of starch grains within the pollen grains decreased
   significantly ([129]Fig. 5D and [130]d). Tapetal cells began to
   degrade, and lipid accumulation within the pollen grains continued to
   increase, with more lipid droplets in the WT ([131]Fig. 5E and [132]e).
   At the mature stage WT pollen grains were rich in lipid droplets
   ([133]Fig. 5F), and the lipid droplet content was significantly higher
   ([134]Fig. 5f). Both mature WT and as11 pollen only contained a small
   amount of starch grains ([135]Fig. 6F and [136]f).

Fig. 5.

   [137]Fig. 5.
   [138]Open in a new tab

   Anther cross sections of WT (A–F) and mutant as11 (a–f) with PAS and
   Sudan Black B staining at different developmental stages. A, a:
   vacuolate microspore stage stained with PAS and Sudan Black B, A. Lipid
   was accumulated in the tapetum (arrows). B, b, C, c: binuclear pollen
   grain II stage (B and b were stained with PAS only, C and c were
   stained with PAS and Sudan Black B). B, b: starch grains were
   accumulated around the vegetative nucleus (arrows). C, c: lipid
   droplets deposited around the generative nucleus (arrows). D, d, E, e:
   second mitosis division I (D and d were stained with PAS only, E and e
   were stained with PAS and Sudan Black B), a few starch grains in the
   pollen grains; E, e: lipid droplets were accumulated in the tapetum
   cells and pollen grains, with fewer in the mutants (e). F, f: mature
   pollen grain II stage stained with PAS and Sudan Black; F, middle layer
   and tapetum had disappeared, and lipid droplets were accumulated in
   pollen grains, with fewer in the mutants (f). Bar = 10 μm. Ep,
   epidermis; En, endothecium; ML, middle layer; T, tapetum.

Fig. 6.

   [139]Fig. 6.
   [140]Open in a new tab

   The ultrastructure of pollen grains in the WT (A–D) and as11 (a–d) at
   different developmental stages. A, a: released microspore II stage; A:
   small lipid droplets occurred in microspores; a: the content of Golgi
   bodies was higher in microspores compared with WT. B, b: bicellular
   pollen grain I stage; B: vegetative nucleus was surrounded by plastids
   with starch grains (arrowheads); b: plastids with starch grains
   distributed around vegetative nucleus (arrowheads). C, c: binuclear
   pollen II stage; C: starch grains were observed around the vegetative
   nucleus (white arrowheads), and lipid droplets were deposited at the
   surface of the generative nucleus (black arrowheads); c: starch grains
   increased (white arrowheads) and lipid droplets were present at the
   surface of the generative nucleus (black arrowheads). D, d: mature
   pollen grain II stage (d-1 is partial enlarged view of d); D: lipid
   droplets and electron-transparent vesicles (black arrows) were
   abundant; d: a small number of lipid droplets were stored in mature
   pollen, which contained abundant electron-dense vesicles (d-1 white
   arrows) and electron-transparent (d and d-1 black arrows) covered with
   endoplasmic reticulum. N, nucleus; Nu, nucleolus; ER, endoplasmic
   reticulum; P, plastid; M, mitochondrion; G, Golgi bodies; L, lipid
   droplet; GC, generative cell; VN, vegetative nucleus.

   To explore the results at the cytological level, the ultrastructure of
   both the WT and as11 was observed at different developmental stages. At
   the released microspore II stage, the microspores had a centrally
   located nucleus that was surrounded by a small amount of endoplasmic
   reticulum, and these microspores began to secrete substances to form
   the pollen intine ([141]Fig. 6A and [142]a). By this stage, small lipid
   droplets had already begun to form in the WT ([143]Fig. 6A), while more
   Golgi bodies were present in as11 ([144]Fig. 6a). At the binuclear
   pollen grain I stage, a lenticular-shaped generative cell produced by
   an asymmetric mitosis was located on one side of the pollen grain and
   surrounded by a continuous intine. Several plastids with starch grains
   were present around the vegetative nuclei of both WT and as11 plants
   ([145]Fig. 6B and [146]b). For further development to the binuclear
   pollen grain II stage, generative cells were free in the cytoplast of
   the vegetative cells, and these cells contained abundant vesicles and
   organelles such as mitochondria, plastids, and others. Lipid droplets
   precipitated at the surface of the generative nucleus, and plastids
   containing starch grains were distributed around the vegetative nuclei
   in both the WT and as11 ([147]Fig. 6C and [148]c). Compared to the WT,
   the content of lipid droplets was reduced but plastids that were rich
   in starch grains were more numerous in mutant as11 at this stage. When
   pollen grains matured, the cytoplasm of WT vegetative cells was rich in
   lipid droplets and endoplasmic reticulum and contained numerous
   electron-transparent vesicles ([149]Fig. 6D). In contrast, lipid
   droplets and endoplasmic reticulum were lacking in as11 pollen
   ([150]Fig. 6d); however, electron-transparent vesicles that were
   covered with endoplasmic reticulum ([151]Fig. 6d and d-1) and
   electron-dense vesicles ([152]Fig. 6d-1) were both abundant in the
   cytoplasm.

3.4 The pollen ultrastructural changes were related to ultrastructural
changes in the elaiosomes and the lipidosomes in the tapetum

   Most of the nutrients in pollen grains come from the tapetum, so we
   also observed and compared the ultrastructure of associated organelles
   in the WT and mutant as11 tapetal cells at different developmental
   stages, with particular focus on the developmental processes of the
   elaiosomes and the lipidosomes.

   At the post-meiosis and cytokinesis stage in WT tapetum, a small number
   of Golgi bodies were present in the cytoplasm, but plastids were
   abundant, and osmiophilic deposits appeared to be internal ([153]Fig.
   7A). Osmiophilic deposits were not obvious within plastids in the
   mutant tapetal cells, and the Golgi bodies were present ([154]Fig. 7a).
   At the tetrad stage, plastids in WT tapetum began to accumulate
   internal osmiophilic and electron-transparent deposits. The endoplasmic
   reticulum was abundant and primarily located between the nucleus and
   the cytomembrane. The Golgi bodies were highly developed ([155]Fig.
   7B). The tapetum of mutant as11 exhibited an increase in the content of
   endoplasmic reticulum and these structures were also rich in Golgi
   bodies actively secreting vesicles. Osmiophilic deposits were also
   present in plastids ([156]Fig. 7b). When microspores were released, in
   the WT tapetum numerous vesicles were stacked along the edge of
   plasmalemma, and the swollen endoplasmic reticulum was more obvious.
   The plastids were not significantly altered ([157]Fig. 7C). In as11,
   the Golgi bodies were very abundant and were swollen with rich
   secretory vesicles. Osmiophilic droplets appeared to be increased
   within plastids ([158]Fig. 7c). At the stage of vacuolate microspores
   in WT tapetum, the endoplasmic reticulum was still plentiful, and a
   large number of plastoglobules that exhibited a centered rectangular
   chip structure began to accumulate in plastids. Additionally, numerous
   expanded Golgi bodies and exuberant secretory vesicles developed into a
   relatively stable elaiosomes. New lipid-like structures called
   lipidosomes also became visible, which was surrounded by extensive
   endoplasmic reticulum ([159]Fig. 7D). At the vacuolate microspore stage
   in as11 tapetum, vesicles harboring electron-dense materials increased
   in number, and plastoglobules began to accumulate within plastids. At
   this stage, lipidosomes were not observed in mutant tapetal cells
   ([160]Fig. 7d). At the binuclear pollen grain I stage, the
   plastoglobules formed a centered rectangular chip structure in plastids
   that subsequently became relatively stable elaiosomes. Additionally, a
   small number of lipidosomes appeared ([161]Fig. 7e). As pollen grains
   continued to develop, the Golgi bodies were reduced and vesicles
   possessing electron-dense inclusions were still present in the WT
   tapetum. The elaiosomes remained stabilized, and the lipidosomes
   increased and fused with each other to form sheeting structures that
   were surrounded by plentiful endoplasmic reticulum ([162]Fig. 7E). At
   the binuclear pollen grain II stage, the tapetal cells were primarily
   occupied by elaiosomess and lipidosomes, and the content of both
   structures reached its maximum ([163]Fig. 7F). The development of
   mutant tapetal cells during the late binuclear pollen grain stage was
   not significantly different from that of the WT ([164]Fig. 7f). The
   content of elaiosomes and liposomes, which occupied most of the volume
   in tapetal cells, also reached its peak at the binuclear pollen grain
   II stage; however, the lipidosome content was not as rich as that of
   the WT ([165]Fig. 7f). At the second mitotic division I stage, the
   tapetal cells of both the WT and as11 were degraded. The electron
   density of the lipidosomes increased, and the structure of the
   elaiosomes remained stable. The structures were both released into the
   anther loculus after the plasma membrane had degraded, and they were
   deposited as a covering for the extine of pollen grains to form the
   tryphine ([166]Fig. 7G and [167]g). At the second mitotic division II
   stage, the tapetum of the WT and as11 continued to release degraded
   cellular inclusions, and ultimately, these cells became vacuolate
   sheeting lipid structures that were fused by the lipidosomes. It should
   be noted that the lipid structure in the mutant as11 was less
   electron-dense and did not fill the entire tapetal residual structure
   compared to the observations in the WT ([168]Fig. 7H and [169]h).

Fig. 7.

   [170]Fig. 7.
   [171]Open in a new tab

   The ultrastructure of the tapetum in the WT and as11 at different
   developmental stages. A, a: Post-meiosis, pre-cytokinesis stage. A: A
   few osmiophilic plastoglobules (white arrowheads) were visible in
   plastids. A: There were no plastoglobules in the cells. B, b: Tetrad
   stage. B: Osmiophilic and non-osmiophilic plastoglobules (black
   arrowheads) were accumulated in plastids. B: Osmiophilic plastoglobules
   (white arrowheads) occurred in plastids. C, c: Released microspore I
   stage. C: The endoplasmic reticulum content was abundant. C: The Golgi
   bodies were abundant with rich secretory vesicles. D, d: Vacuolate
   microspore stage. D: Rectangular chip-shaped substances (arrows) were
   presented in plastoglobules. D: Plastids started to accumulate
   non-osmiophilic plastoglobules (white arrowheads). E, e: Vacuolate
   microspores. E: Elaiosomes (black asterisks) became stable and
   lipidosomes (white asterisks) occurred. e: Rectangular chip-shaped
   substances (white arrows) were present in the plastoglobules.
   Elaiosomes (black asterisks) became stable and lipidosomes (white
   asterisks) occurred. F, f: Binuclear pollen grain I stage. Vesicle
   containing electron-dense materials fused with plasmalemma to release
   inclusions to anther locule (black arrows), and lipidosomes (white
   asterisks) coalesced with each other to form sheeting structures. G, g:
   nuclearpollen grain II stage. Elaiosomes (black asterisks) and
   lipidosomes (white asterisks) occupied most of the space in the
   tapetum. H, h: Second mitotic division I. H: Tapetal plasmalemmas broke
   down (white arrows) and freed lipidosomes and elaiosomes to deposit on
   the surface of the pollen extine. H: Tapetal plasmalemmas break down
   (white arrows) and freed lipidosomes and elaiosomes to anther locule.
   I, i: Second mitotic division II stage. I: Tapetal cells degraded
   completely, only leaving lamellate lipid structures fused by
   lipidosomes (black arrows). Tapetal cells degraded completely, only
   leaving lamellate lipid structures fused with lipidosomes (black
   arrows). N, nucleus; Nu, nucleolus; ER, endoplasmic reticulum; P,
   plastid; M, mitochondrion; G, Golgi bodies.

   We randomly calculated the area of the lipidosomes and elaiosomes in 20
   tapetal cells from the WT and mutant at the binuclear pollen grain
   stage, and found that the area of lipidosomes in WT tapetal cells was
   3.49 times that of the mutant, while the area of the elaiosomes in
   mutant tapetal cells was 2.26 times that of the WT ([172]Fig. 8).

Fig. 8.

   [173]Fig. 8.
   [174]Open in a new tab

   The area ratio of lipidosomes and elaisomes between WT and as11.

4 Discussion

4.1 Fatty acid metabolism-related DEGs involved in as11

   Transcriptome sequencing is now a routine experimental method for
   differential gene expression, quantitative gene expression, and
   transcriptional identification ([175]Mao et al., 2021). [176]Kim et al.
   (2016) analyzed and identified genes related to fatty acid biosynthesis
   in Perilla frutescens (L.) var frutescens by transcriptomic analysis.
   Studies on changes in the fatty acid metabolism in Arabidopsis DGAT1
   mutant as11 seed have been relatively comprehensive, and the transcript
   profiling of developing WT and as11 seed showed differences in the
   fatty acid metabolism gene expression ([177]Zou et al., 1999;
   [178]Aulakh and Durrett, 2019), but the genes related to fatty acid
   biosynthesis during pollen development of the mutant have not yet been
   clarified. [179]Zhang et al. (2021) found fatty acid metabolism genes
   in Ophiocordyceps sinensis by transcriptomics. In the current study, a
   total of 10,988 differential genes were identified, of which 5,175 were
   upregulated and 5,813 were downregulated according to the transcriptome
   analysis ([180]Fig. 1).

   DGAT is an important enzyme catalyzing TAG synthesis in the Kennedy
   pathway, which is of great significance for TAG synthesis and
   accumulation ([181]Zou et al., 1999; [182]Lu and Hills, 2002; [183]Lu
   et al., 2003). In addition to the Kennedy pathway, [184]Dahlqvist et
   al. (2000) found that phospholipid:diacylglycerol acyltransferase
   (PDAT) in Arabidopsis thaliana can catalyze the synthesis of TAG. The
   GPDHc1 gene expression product is glycerol-3-phosphate dehydrogenase
   with NAD dependence, which can directly catalyze the formation of
   dihydroxyacetone phosphate (DHAP) in the process of sugar metabolism to
   generate glycerol-3-phosphate (G3P), and then directly enter the
   Kennedy pathway to provide the glycerol framework for TAG synthesis. It
   plays an important role in the mutual conversion of sugars and lipids
   ([185]Shen et al., 2006; [186]To et al., 2012). The FAB1 gene
   expression product β-Ketoacyl-ACP synthase II (KAS II) catalyzed the
   elongation of fatty acids from C16:0-ACP to C18:0-ACP ([187]Hirano et
   al., 2001; [188]Pidkowich et al., 2007).

   Phosphatidylcholine:diacylglycerol choline phosphotransferase PDCT can
   transfer choline from C18:2/3-PC to C18:1-DAG and generate C18:2/3-DAG
   and C18:1-PC. The former can continue to synthesize C18:2/3-TAG. The
   latter, as a substrate for fatty acid dehydrogenase (FAD), continues to
   be desaturated to produce C18:2/3-PC ([189]Lu et al., 2009). PAH2 can
   encode phosphatidic acid phosphatase (PAP) and catalyze the production
   of diacylglycerol DAG ([190]Eastmond et al., 2010). The DGAT activity
   of as11 seeds decreased by 40–70% relative to the WT, which affected
   the amount of oil accumulation in seeds, especially the synthesis of
   TAG ([191]Katavic et al., 1995; [192]Zou et al., 1999), whereas the
   amount of α-linolenic acid (18:3) almost doubled ([193]Zou et al.,
   1999; [194]Jako et al., 2001; [195]Lu and Hills, 2002), this being the
   main fatty acid for storing TAG in mature seeds ([196]Zou et al.,
   1999). In the GO enrichment analysis of differentially expressed genes,
   10,562 genes were annotated into 30 subclasses of two GO classes:
   biological processes and cellular components. Following a KEGG
   enrichment analysis of differentially expressed genes in fatty acid
   metabolism and fatty acid degradation and based on the results of
   functional enrichment analysis of DEGs, we selected fatty acids as the
   focus of research on the differences between WT and as11. The
   transcriptomic data of as11 revealed that the expression levels of
   GPGHC1, GPAT1, GPAT6, GPAT9, PDAT1, PAH2 and FAB1 all decreased in
   as11. So, 7 genes related to TAG biosynthesis were selected for further
   analysis ([197]Table 1).

4.2 Lipid accumulation and fatty acid composition change during pollen
development in as11

   Previous studies demonstrated that the content of oil and TAG in mature
   seeds of the Arabidopsis DGAT1 gene mutant as11 was distinctly lower
   than that in the WT ([198]Katavic et al., 1995; [199]Zou et al., 1999;
   [200]Lu and Hills, 2002). The DGAT1 gene can also reduce TAG
   accumulation in other nutritional organs of Arabidopsis ([201]Slocombe
   et al., 2009). Additionally, the TAG content in mature seeds, roots,
   stems, leaves, and petals was decreased in tobacco after silencing of
   the DGAT1 gene ([202]Zhang et al., 2005). Antisense suppression of the
   DGAT1 gene in Brassica napus not only decreased the oil content of
   mature seeds, but also reduced the seed yield and germination rate and
   resulted in highly abnormal development ([203]Lock et al., 2009).
   Although plant pollen is very small, it is extremely abundant in
   nutrients such as lipid droplets and/or starch, which are needed to
   ensure that there is sufficient energy and material to promote the
   normal germination of pollen. Any abnormal accumulation of nutrients
   during pollen development may cause pollen sterility ([204]Zhang et
   al., 2002, [205]2009; [206]Nashilevitz et al., 2009; [207]Li et al.,
   2012; [208]Yang et al., 2017; [209]Gao et al., 2019). As the main
   component of lipid stored in pollen grains, triacylglycerol (TAG) not
   only serves as an energy and carbon source, but it also participates in
   important metabolic processes such as membrane lipid remodeling and has
   great significance in regard to normal development and successful
   pollination ([210]Yang and Benning, 2018). DGAT, the enzyme catalyzing
   the synthesis of TAG, is important for TAG synthesis and accumulation.
   It not only has a high content and activity in developing seeds, but
   also widely exists in different organs such as leaves, petals, fruits,
   and anthers ([211]Hobbs et al., 1999; [212]Zou et al., 1999; [213]Lu
   and Hills, 2002; [214]Lu et al., 2003). It was reported that during
   pollen development in Arabidopsis thaliana ([215]Owen and Makaroff,
   1995; [216]Zajac, 1997; [217]Zhang et al., 2002) and Brassica napus
   ([218]Platt et al., 1998), lipid droplets first begin to accumulate at
   the binuclear pollen grain stage where they are mainly distributed
   around the generative nucleus, and these lipid droplets gradually
   increase as pollen develops until the pollen has matured. This
   phenomenon is not limited to cruciferous plants, but can also be
   observed in the pollen development of other plants such as Lycium
   barbarum ([219]Xu et al., 2009) and Pancratium maritimum ([220]Konyar,
   2018), indicating that the binuclear pollen stage of pollen development
   is a critical period for the accumulation of important nutrients. The
   current study found that although DGAT1 gene mutation did not cause
   pollen abortion, it seriously affected the lipid accumulation
   characteristics and content during pollen development. At the binuclear
   pollen grain II stage, although lipid droplets that accumulated in
   vegetative cells of mutant as11 increased compared to the previous
   stage and were mostly distributed around the generative cell nucleus,
   the content was significantly less than that of the WT. In contrast,
   the number of plastids containing starch grains surrounding the
   vegetative nucleus was significantly higher in the mutant than in the
   WT ([221]Fig. 5C and [222]c; [223]Fig. 6C and [224]c). As the pollen
   developed, lipid continued to accumulate in pollen grains. Mature as11
   pollen was not as rich in lipid droplets as WT pollen ([225]Fig. 5F and
   [226]f; [227]Fig. 6D and [228]d). Furthermore there was less total
   lipid and TAG in mature as11 pollen than in the WT ([229]Fig. 4A and
   [230]C). Previous studies revealed that seed oil in mutant as11 was
   25–35% lower than in the WT ([231]Zou et al., 1999) and oil accumulated
   in seeds to approximately 70% of that in the WT ([232]Xu et al., 2012).
   Our results revealed that the DGAT1 gene also affected lipid and TAG
   synthesis and accumulation in pollen grains, and that mutating the
   DGAT1 gene reduced the lipid and TAG content in pollen compared to that
   in seeds.

   Fatty acids are important components of lipids. [233]Heyl (1923) found
   that the pollen of ragweed contained saturated fatty acids such as
   lauric acid (C12:0), carmine acid (C14:0), and palmitic acid (C16:0)
   and unsaturated fatty acids such as oleic acid (C18:1) and linoleic
   acid (C18:2). [234]Ching (1962) and [235]Standifer (1966) detected
   fatty acids in pollen of five pine species and dandelion species
   respectively, and found that the contents of palmitic acid (C16:0),
   stearic acid (C18:0), oleic acid (C18:1), linoleic acid (C18:2) and
   linolenic acid (C18:3) were higher. Thus the fatty acids of 16-carbon
   palmitic acid (C16:0) and 18-carbon fatty acids are the main types in
   mature pollen. [236]Zheng et al. (2003) measured fatty acids in mature
   pollen of WT and GPAT1-1 mutants in Arabidopsis thaliana. They found
   that mutation of GPAT1 affected the synthesis of linolenic acid (C18:3)
   in pollen. The fatty acids that accumulated in mature pollen of the WT
   and as11 mutant were similar to those that accumulated in Arabidopsis
   seeds ([237]Lu et al., 2009; [238]Napier and Graham, 2010), and,
   consistent with the types of fatty acids in pollen measured by
   [239]Zheng et al. (2003), palmitic acid (C16:0) and linolenic acid
   (C18:3) were the main fatty acids accumulated in the current study
   ([240]Fig. 4D). However, we also measured the pollen fatty acid
   composition at the VM, BN, and SMD stages, and observed dynamic changes
   in fatty acid content during pollen development. The results showed
   that linolenic acid (C18:3) was the main fatty acid in pollen of the WT
   and as11 during the second mitosis stage, with higher levels in the
   mutant ([241]Fig. S3). Moreover, changes in the relative content ratio
   of linolenic acid in the anthers at each developmental stage ([242]Fig.
   S3) indicated that the content and synthesis rate of linolenic acid
   (C18:3) in as11 pollen grains were highest from the BN stage to the SMD
   stage. It has been reported that the DGAT1 gene can affect the
   composition of fatty acids accumulated in seeds of Arabidopsis
   thaliana, in which the level of α-linolenic acid (C18:3) accumulation
   in mature seeds of as11 was nearly doubled compared with that of the WT
   ([243]Katavic et al., 1995; [244]Hobbs et al., 1999), being the main
   storage fatty acid for TAG in mature seeds ([245]Zou et al., 1999). In
   conclusion, the DGAT1 gene of Arabidopsis thaliana does not affect the
   type and content of fatty acids accumulated in pollen, but can affect
   the synthesis rate of linolenic acid. Therefore, we speculate that
   DGAT1 affects the expression of genes related to linolenic acid (C18:3)
   synthesis.

4.3 Lipid accumulation and fatty acid composition may be related to the
changes in TAG biosynthesis

   In addition to the Kennedy pathway, TAG was found to be available
   through phospholipid:diacylglycerol acyltransferase (PDAT)-mediated
   catalysis of DAG using phospholipids in Arabidopsis thaliana
   ([246]Dahlqvist et al., 2000). In Arabidopsis seeds, PDAT is not the
   primary determinant of TAG synthesis, as the low transcript levels of
   PDAT1 in WT seeds were unchanged in DGAT1 mutant seed ([247]Aulakh and
   Durrett, 2019). However, after mutating the DGAT1 gene, the expression
   of the PDAT1 and LPCAT2 genes was upregulated, and these pathways
   became the primary pathways for TAG synthesis and resulted in
   approximately 70% more oil accumulation in seeds than that in the WT
   ([248]Xu et al., 2012). The failure to obtain a double mutant after
   crossing the DGAT1 gene mutant and the PDAT1 gene mutant indicated that
   the functions of DGAT1 and PDAT1 are complementary during pollen
   development ([249]Zhang et al., 2009). However, our research found that
   the expression levels of PDAT1 were higher in mature WT pollen grains.
   The expression of GPAT9, which is involved in seed TAG synthesis
   ([250]Shockey et al., 2016), was also not clearly upregulated in as11.
   [251]Zheng et al. (2003) found that the TAG content in flower buds of
   GPAT1 gene mutants was decreased by approximately 10%, indicating that
   the GPAT1 gene is involved in TAG synthesis in floral organs. The GPAT6
   gene, which is mainly expressed in the floral organs including the
   tapetum and microspores, could cause abnormal accumulation of nutrients
   in pollen and defects in pollen wall development by affecting the lipid
   metabolism of the tapetum, and this gene can also cause premature
   anther senility ([252]Li et al., 2012). It was reported that the GPAT1
   and GPAT6 genes exerted important effects on the normal development of
   Arabidopsis pollen ([253]Zheng et al., 2003; [254]Li et al., 2012).
   Additionally, the expression of the PAH2 gene promotes the
   transformation of sugar to lipids and provides the glycerin skeleton
   for TAG synthesis, which catalyzes DAG synthesis and the GPDHc1 gene
   ([255]Shen et al., 2006; [256]Eastmond et al., 2010; [257]To et al.,
   2012). Recent research shows that the soybean GPDHc1 gene is involved
   in lipid synthesis in the endoplasmic reticulum, and increases the
   content of C18:1 fatty acid in TAG by increasing the diacylglycerol (de
   novoDAG) produced by the G3P pathway ([258]Zhao et al., 2021). So, we
   suggest that the high expression levels of GPAT1 and GPAT6 genes in the
   WT, in accordance with the trend for increased GPDHc1 and PAH2
   expression in the same pathway of DAG synthesis, promotes TAG synthesis
   ([259]Fig. 4) and the synthesis of endoplasmic reticulum in the
   tapetum. The synthesis of lipidosomes that are rich in TAG is closely
   related to that of the endoplasmic reticulum. Therefore, the high
   expression of GPAT1 and GPAT6 in the WT could promote the synthesis of
   endoplasmic reticulum in the tapetum. As a result, the obvious
   lipidosomes that were observed in the tapetal cells could provide raw
   materials for lipid synthesis in pollen.

   DGAT1 gene expression not only affects the content of oil in seeds, but
   can also alter the type of fatty acid accumulation. Compared to that in
   the WT, the accumulation of linolenic acid (C18:3) was twice as high in
   as11 mature seeds ([260]Katavic et al., 1995; [261]Hobbs et al., 1999;
   [262]Routaboul et al., 1999; [263]Zou et al., 1999; [264]Jako et al.,
   2001; [265]Lu and Hills, 2002), and it is the main fatty acid stored in
   the TAG of mature seeds ([266]Zou et al., 1999). Our study indicated
   that the main fatty acids in mature mutant as11 pollen grains did not
   differ from that in the WT ([267]Fig. 2D). However, during the SMD
   stage, linolenic acid (C18:3) became the main fatty acid in as11, based
   on the change trend of relative content of linolenic acid (C18:3)
   during development ([268]Fig. S3). It was concluded that DGAT1
   increases the content of linolenic acid (C18:3) by regulating the
   expression of FAD2 during pollen development. In DGAT1 mutant seed, the
   transcript levels of FAD2 encoding the ER-localized desaturases
   responsible for the synthesis of 18:2 and 18:3 were higher than that of
   the WT, and the desaturase genes that introduce double bonds in various
   plastidial glycerolipids were unaffected, like FAB2 ([269]Aulakh and
   Durrett, 2019). It was reported that the linolenic acid (C18:3) content
   was decreased in mutant GPAT1-1 mature pollen grains and seeds
   ([270]Zheng et al., 2003), and increased in mutant GPAT9 mature seeds,
   compared to the WT ([271]Shockey et al., 2016). Consistent with
   previous work, GPAT1 was slightly upregulated in mutant seed ([272]Zou
   et al., 1999; [273]Aulakh and Durrett, 2019). Therefore, the high
   expression of GPAT1 and slight upregulation of GPAT9 gene expression in
   the WT may lead to a slight increase in linolenic acid (C18:3) content,
   and we speculated that this could be one of the reasons for the
   upregulation of FAD2 gene expression. Furthermore, scanning electron
   microscopy indicated that there were few mature pollen grains in the
   mutant GPAT1-1, and this significantly reduced pollen fertility
   ([274]Zheng et al., 2003). Based on this, the changes in pollen fatty
   acid composition may also exert effects on pollen fertility. The
   observed normal pollen morphology and self-pollination of as11
   ([275]Katavic et al., 1995), however, indicate that to guarantee pollen
   sterility, the fatty acid composition of as11 mature pollen grains
   should be significantly different. When the pollens were mature, the
   composition of the main fatty acids in as11 tended to be similar to
   that in the WT ([276]Fig. 5D). It was shown that in order to guarantee
   pollen sterility, FAD2 expression decreased, which prevented the
   accumulation of linolenic acid, with the composition of fatty acids in
   the as11 mature pollen gradually tending to be the same as the
   composition of the main fatty acids in WT pollen grains. Thus,
   expression of the DGAT1 gene affects lipid accumulation and fatty acid
   composition during pollen development, but does not affect pollen
   fertility.

   In conclusion, the DGAT1 gene can affect the accumulation of TAG in
   pollen grains by affecting the expression levels of GPDH, GPAT1, GPAT6,
   GPAT9 and PAH2, and affects the composition of fatty acids in pollen by
   affecting the expression levels of FAB1 and FAD2.

4.4 The lower lipid content in as11 may be indirectly related to the
development of elaiosomes and lipidosomes in tapetal cells

   As the innermost anther parietal cells, tapetal cells possess the
   function of absorbing and transporting polysaccharides or lipids to
   provide nutrients for the development and germination of pollen.
   Therefore, abnormal metabolism in tapetal cells will directly affect
   the development of microspores in the anther loculus and the
   macromolecular nutrient synthesis and accumulation that occurs in these
   cells. In addition to pollen grains, the tapetum also secretes lipids
   to nourish microspores during anther development ([277]Zhu et al.,
   2011; [278]Chen et al., 2013). Studies examining gene expression
   related to TAG synthesis using in situ labeling have shown that the
   anther is the second most active tissue besides the seed
   ([279]Piffanelli et al., 1997, [280]1998; [281]Murphy, 2001). In regard
   to the lipid cytological changes in the tapetum and pollen grains,
   [282]Hernandez-Pinzón et al., (1999) isolated two organelles, the
   elaiosomes and the tapetosome, that were related to lipid synthesis by
   the tapetum of Brassica napus. The former is a type of oil-containing
   organelle that functions in the process of plant pollen development
   that involves sterol esters, TAG, and protein ([283]Hernandez-Pinzón et
   al., 1999; [284]Hsieh and Huang, 2004). The latter is a specific
   organelle in tapetal cells ([285]Wu et al., 1997), and is rich in TAG
   and some proteins similar to seed oleosin. The tapetosome is closely
   related to the endoplasmic reticulum during anther development and is
   generally believed to be formed by rough endoplasmic reticulum budding
   ([286]Hsieh and Huang, 2005). Similar structures have been observed in
   the tapetum of the Arabidopsis thaliana ecotype Columbia ([287]Zhang et
   al., 2002; [288]Suzuki et al., 2013), and these structures are called
   lipidosomes. The proteins in lipidosomes and sterol esters in
   elaiosomes are primarily used to synthesize the pollen coat. The
   specific function of TAG located within these structures is unclear,
   but it may be transported through the pollen coat by esterase or
   β-oxidation to the inner pollen for further metabolism and utilization
   ([289]Zhang et al., 1994; [290]Hernandez-Pinzón et al., 1999). In this
   study, we found that despite the fact that the lipidosomes and
   elaiosomes were both present in the tapetal cells of WT and as11
   mutants, there were differences in timing of occurrence and their
   characteristics.

   First, plastids in the WT began to exhibit osmiophilic deposits at the
   post-meiosis and pre-cytokinesis stage ([291]Fig. 7A), and reached a
   steady state during the vacuolate microspore stage when the lipidosomes
   formed ([292]Fig. 7E). Plastids of mutant as11 began to exhibit
   osmiophilic deposits at the tetrad stage ([293]Fig. 7e), and reached a
   relatively steady state at the binuclear pollen grain I stage when the
   lipidosomes formed ([294]Fig. 7f). Both organelles filled almost the
   entire tapetum in the WT and mutant at the binuclear pollen grain II
   stage ([295]Fig. 7G and [296]g), when the content reached its maximum.
   At the second mitosis stage, the tapetum was degraded and these
   structures were released into the anther loculus where they covered the
   surface of pollen grains. The area ratio of lipidosomes and elaiosomes
   showed that the area of lipidosomes was higher in the WT tapetal cells,
   while the area of elaiosomes was higher in mutant tapetal cells.
   Throughout the entire process of pollen development, the content of
   endoplasmic reticulum in WT tapetal cells was generally richer than
   that in as11, and the lipidosomes were primarily formed by rough
   endoplasmic reticulum budding ([297]Platt et al., 1998; [298]Ting et
   al., 1998; [299]Hsieh and Huang, 2004, [300]2005). The MAGL gene (a
   homolog of AtMAGL8) which encodes the monoacylglycerol lipase (MAGL) in
   Arabidopsis thaliana, results in male sterility, and the lipidosome
   content in the tapetum was higher than that in the mutant ([301]Gao et
   al., 2019). Based on this, the lipidosomes that were rich in TAG in the
   mutants appeared later and were less abundant than those observed in
   the WT. [302]Zheng et al. (2003) examined the ultrastructure of GPAT1
   mutant anthers and found that the fusion of endoplasmic reticulum with
   plasma membranes occurred rarely and endoplasmic reticulum’s dilation
   was generally decreased. The ultrastructure of the GPAT6 gene mutant
   anther also indicated that the content of the endoplasmic reticulum in
   tapetal cells was reduced and was difficult to detect in mature pollen
   grains ([303]Li et al., 2012). The above results indicate that the
   relevant genes within the TAG synthesis pathway may also exert a
   certain impact on the development of the pollen. It has been reported
   that DGAT, an important enzyme encoded by the DGAT1 gene, catalyzes the
   synthesis of TAG and is located on the endoplasmic reticulum
   ([304]Routaboul et al., 1999; [305]Zou et al., 1999; [306]Jako et al.,
   2001; [307]Kaup et al., 2002; [308]Lu and Hills, 2002; [309]Ramli et
   al., 2005). Thus, it is justifiable that in mutant as11, the decreased
   DGAT enzyme activity resulted in a lower content of endoplasmic
   reticulum in tapetal cells and pollen grains.

   The elaiosomes contain only small amounts of TAG, and the reduction in
   DGAT activity should have no significant effect on its content.
   However, our study found that the content of elaiosomes in the mutant
   was higher than that in the WT. We speculated that the mutant may
   supplement TAG accumulation by increasing the number of elaiosomes.
   Compared to the WT, the content of Golgi bodies, which is closely
   related to the position of the elaiosomes, was relatively rich in
   mutant tapetum development. Studies have shown that the Golgi bodies
   are closely associated with the secretion of polysaccharides in plant
   cells ([310]Buvat, 1989), as their membranes possess a variety of
   enzymes involved in polysaccharide synthesis. We considered that in the
   process of as11 pollen development, due to the reduced TAG content, the
   SS1 and SS2 genes related to starch synthesis were clearly upregulated
   at the mature pollen grain stages to increase polysaccharide substances
   ([311]Fig. 5B). As a result, the content of Golgi bodies that were
   associated with polysaccharide secretion also increased. Additionally,
   lipids are involved in the processing of secretory vesicles of the
   Golgi bodies ([312]Kirchhausen, 2000; [313]Shemesh et al., 2003;
   [314]Weiss and Nilsson, 2003; [315]Bethune et al., 2006). In
   particular, DAG is involved in the production of COP vesicles during
   the transport process facilitated by the Golgi-endoplasmic reticulum
   ([316]Asp et al., 2009; [317]Martinez-Alonso et al., 2013), suggesting
   that DAG has a specific relationship with the transport of Golgi bodies
   and endoplasmic reticulum. The content of endoplasmic reticulum systems
   in tapetal cells of the mutants was less than that in the WT, and the
   reduction in TAG synthesis led to an increase in DAG accumulation. The
   Golgi body secreted vesicles increased instead of decreased endoplasmic
   reticulum vesicles.

   Based on our findings, we concluded that the DGAT1 gene regulates
   tapetum cell structure during pollen development. DGAT1 indirectly
   regulates the activity characteristics of the lipidosomes and
   elaiosomes by affecting the contents of the endoplasmic reticulum and
   Golgi bodies and their secretory vesicle activity, and DGAT1 ultimately
   affects the lipid accumulation in mature pollen grains.

Supporting Information

   The following additional information is available in the online version
   of this article –

   Fig. S1. General bioinformatics analysis of all unigenes. A. GO
   classifications of the Venn diagram of annotated unigenes. B. Histogram
   of the KOG categories.

   Fig. S2. Anther cross sections of WT (A–F) and mutant as11 (a–f) with
   histochemical staining of PAS and Sudan Black at different
   developmental stages. A, a: premeiosis I stage, the anther wall
   composed of four layers, respectively epidermis, endothecium, middle
   layer and tapetum. The microspore mother cell has a large and distinct
   nucleus (arrow); B, b: premeiosis II stage, the microspore mother cells
   begin to deposit callose, the tapetum cells contract, and cytoplasm
   contains numerous vesicles (arrow). C, c: meiosis stage; C, the
   microspore mother cell was surrounded by callose, and the tapetum cells
   contracted with dense cytoplasm and a larger vesicle (arrow); c, the
   mother cell of the spore surrounded the callose, the tapetum cells
   contracted, with many small vesicles (arrow); D, d: Tetrad stage, after
   cytokinesis, the microspore mother cell formed four mononuclear
   microspores, which were enclosed and separated by callose (arrow). E,
   e: Released microspore I stage, callose wall was degraded and an
   irregular-shaped microspore was released into the pollen sac; only one
   nucleus was located in the microspore (arrow). F, f: Released
   microspore I stage, the microspore became more rounded and had thick
   walls. Ep, epidermis; En, endothecium; ML, middle layer; T, tapetum.

   Fig. S3. Relative content of total fatty acids (A–C) and rate of
   relative content of three fatty acids (D–G) in WT and mutant as11 at
   different developmental stages. VM, ring-vacuolate microspores; BN,
   binuclear pollen grains; SMD, second mitotic division; MP. Significant
   differences are indicated by *P < 0.05 and **P < 0.01. Three biological
   replicates were performed.

   Table S1. The primer sequences for qRT-PCR.
   plad012_suppl_Supplementary_Material
   [318]Click here for additional data file.^ (1.1MB, pdf)

Acknowledgements