Overexpression of a Prefoldin β subunit gene reduces biomass recalcitrance in the bioenergy crop Populus

Summary Prefoldin (PFD) is a group II chaperonin that is ubiquitously present in the eukaryotic kingdom. Six subunits (PFD1‐6) form a jellyfish‐like heterohexameric PFD complex and function in protein folding and cytoskeleton organization. However, little is known about its function in plant cell wall‐related processes. Here, we report the functional characterization of a PFD gene from Populus deltoides, designated as PdPFD2.2. There are two copies of PFD2 in Populus, and PdPFD2.2 was ubiquitously expressed with high transcript abundance in the cambial region. PdPFD2.2 can physically interact with DELLA protein RGA1_8g, and its subcellular localization is affected by the interaction. In P. deltoides transgenic plants overexpressing PdPFD2.2, the lignin syringyl/guaiacyl ratio was increased, but cellulose content and crystallinity index were unchanged. In addition, the total released sugar (glucose and xylose) amounts were increased by 7.6% and 6.1%, respectively, in two transgenic lines. Transcriptomic and metabolomic analyses revealed that secondary metabolic pathways, including lignin and flavonoid biosynthesis, were affected by overexpressing PdPFD2.2. A total of eight hub transcription factors (TFs) were identified based on TF binding sites of differentially expressed genes in Populus transgenic plants overexpressing PdPFD2.2. In addition, several known cell wall‐related TFs, such as MYB3, MYB4, MYB7, TT8 and XND1, were affected by overexpression of PdPFD2.2. These results suggest that overexpression of PdPFD2.2 can reduce biomass recalcitrance and PdPFD2.2 is a promising target for genetic engineering to improve feedstock characteristics to enhance biofuel conversion and reduce the cost of lignocellulosic biofuel production.

For cellulose and hemicellulose isolation, the extractives-free samples were delignified by peracetic acid with 5.00 g loading per g biomass. The solution consistency was adjusted to 5% with deionized (DI) water and the holopulping was conducted at room temperature for 24 h with magnetic stirring. The solid residue, designated as holocellulose, was washed with excessive DI water (18.0 MΩ) and air dried at room temperature for 24 h. A sub-portion of the air-dried holocellulose (100 mg) was consecutively extracted at 25°C with 17.5% (wt/v) NaOH solution (5.00 mL) for 2 h, followed by 8.75% NaOH solution (10.00 mL) for an additional 2 h. The alkaline slurry was then filtered and rinsed with 5 mL of 1% acetic acid leading to a liquid fraction and a solid residue. The solid residue, namely α-cellulose, was washed with an excess of DI water and air dried for the analysis of cellulose DP. The liquid fraction, rich in hemicellulose, was adjusted to pH 6-7 with anhydrous acetic acid. Hemicellulose was then precipitated by adding three volumes of 100% ethanol to the liquid fraction. Hemicellulose was then obtained by centrifugation at 8000 rpm (267π rad/s) for 5 min and freeze dried for 24 h.

Methods S2. Saccharification assay.
Dried and Wiley-milled (40 mesh) stems of the Populus control and transgenic plants were used for saccharification assays. In brief, biomass was extracted with α-amylase (Spirizyme Ultra, 0.25%) and αglucosidase (Liquozyme SC DS, 1.5%) in 0.1 M sodium acetate (24 h, 55°C, pH 5.0) to remove possible starch content; followed by an ethanol (95% v/v) Soxhlet extraction for an additional 24 h. After drying overnight, 5 mg (±0.5 mg) of biomass was weighed in triplicate into one of 96 wells in a solid Hastelloy microtitre plates and 250 μL of water was added. Samples were then sealed with silicone adhesive, Teflon tape. For pretreatment, the samples were reacted at 180°C for 17.5 min, after cooled, 40 μL of bufferenzyme stock [8% CTec2 (Novozymes, Bagsvaerd, Denmark) (excess enzyme loading of 70 mg/g biomass) in 1 M sodium citrate buffer] was added. The samples were then gently mixed and left to statically incubate at 50°C for 70 h. After incubation, an aliquot of the saccharified hydrolysate was diluted and tested using megazymes GOPOD (glucose oxidase/peroxidase) and XDH assays (xylose dehydrogenase). Results were calculated using standard curves created from mixtures of glucose and xylose.
Raw fastq file reads were filtered and trimmed using the JGI QC pipeline. Using BBDuk (https://sourceforge.net/projects/bbmap/), raw reads were evaluated for sequence artifacts by kmer matching (kmer=25) allowing 1 mismatch, and detected artifacts were trimmed from the 3' end of the reads. RNA spike-in reads, PhiX reads and reads containing any Ns were removed. Quality trimming was performed using the phred trimming method set at Q6. Following trimming, reads under the length threshold were removed (minimum length 25 bases or 1/3 of the original read length; whichever was longer). Raw reads from each library were aligned to the P. trichocarpa reference genome (https://phytozome.jgi.doe.gov/pz/portal.html#) using TopHat2 (Kim et al., 2013). Only reads that mapped uniquely to one locus were counted. FeatureCounts (Liao et al., 2014) was used to generate raw gene counts, which were used to evaluate the level of correlation between biological replicates. DESeq2 (Love et al., 2014) was subsequently used to determine which genes were differentially expressed between pairs of conditions (P value < 0.05).

Methods S4. Metabolomic analysis by gas chromatography-mass spectrometry (GC-MS).
The leaf tissues were ground with liquid nitrogen in a chilled mortar and pestle with ~ 50 mg FW; and were subsequently twice extracted with 2.5 mL 80% ethanol overnight and then combined prior to drying a 1 mL aliquot in a nitrogen stream. Sorbitol was added before extraction as an internal standard to correct for differences in extraction efficiency, subsequent differences in derivatization efficiency, and changes in sample volume during heating. Dried extracts were dissolved in 500 μL of silylation-grade acetonitrile, followed by the addition of 500 μL N-methyl-N-trimethylsilyltrifluoroacetamide (MSTFA) with 1% trimethylchlorosilane (TMCS) (Thermo Scientific, Bellefonte, PA), and samples then heated for 1 h at 70°C to generate trimethylsilyl (TMS) derivatives (Tschaplinski et al., 2012). After 2 days, 1-μL aliquots were injected into an Agilent Technologies Inc. (Santa Clara, CA) 5975C inert XL GC-MS, fitted with an Rtx-5MS with Integra-guard (5% diphenyl/95% dimethyl polysiloxane) 30 m × 250 µm × 0.25 µm film thickness capillary column. The standard quadrupole GC-MS was operated in the electron impact (70 eV) ionization mode, targeting 2.5 full-spectrum (50-650 Da) scans per second, as described previously (Tschaplinski et al., 2012). Metabolite peaks were extracted using a key selected ion, characteristic m/z fragment, rather than the total ion chromatogram, to minimize integrating co-eluting metabolites. The extracted peaks of known metabolites were scaled back up to the total ion current using predetermined scaling factors. Peaks were quantified by area integration and concentrations normalized to the quantity of the internal standard (sorbitol) recovered, amount of sample extracted, derivatized, and injected. A large user-created database (> 2400 spectra) of mass spectral electron impact ionization (EI) fragmentation patterns of TMS-derivatized compounds, as well as the Wiley Registry 10th Edition combined with NIST 2014 mass spectral database, were used to identify the metabolites of interest to be quantified. Unidentified metabolites were denoted by their retention time as well as key mass-to-charge (m/z) ratios and partial naming given the typical identity of specific m/z. Figure S1. GO enrichment of genes co-expressed with PtPFD2.2. Figure S2. cis-acting elements in promoter region of PtPFD2.2. Figure S3. Protein structure of PtPFD2.2.
(a) Secondary protein structure of PtPFD2.2 with potential post-translation modification sites. Yellow, red and blue pins represent phosphorylation, sumoylation, and ubiquitination sites, respectively. Conserved Prefoldin_2 motifs identified from pfam database is shown in the blue box (13-118 aa). (b) Protein 3D structure of PtPFD2.2 (b) Protein 3D structure of PtPFD2.2. Figure S4. Expression of PdPFD2.2 in control (Ctrl) and two overexpression lines through qRT-PCR.