Lignin is a major component of cell wall biomass and decisively affects biomass utilisation. Engineering of lignin biosynthesis is extensively studied, while lignin modification often causes growth defects.We developed a strategy for cell‐type‐specific modification of lignin to achieve improvements in cell wall property without growth penalty. We targeted a lignin‐related transcription factor, LTF1, for modification of lignin biosynthesis. LTF1 can be engineered to a nonphosphorylation form which is introduced into Populus under the control of either a vessel‐specific or fibre‐specific promoter.The transgenics with lignin suppression in vessels showed severe dwarfism and thin‐walled vessels, while the transgenics with lignin suppression in fibres displayed vigorous growth with normal vessels under phytotron, glasshouse and field conditions. In‐depth lignin structural analyses revealed that such cell‐type‐specific downregulation of lignin biosynthesis led to the alteration of overall lignin composition in xylem tissues reflecting the population of distinctive lignin polymers produced in vessel and fibre cells.This study demonstrates that fibre‐specific suppression of lignin biosynthesis resulted in the improvement of wood biomass quality and saccharification efficiency and presents an effective strategy to precisely regulate lignin biosynthesis with desired growth performance.
Wood is an important carbon‐neutral biomass for production of timber products, paper pulping, chemicals and biofuels. In terms of anatomy, wood is structured mainly with thickened secondary cell walls, which primarily consist of cellulose impregnated with hemicelluloses and lignin (Kumar et al. , 20162016). Being a major component of wood, lignin is a phenolic polymer, providing cell walls with fortified mechanical strength and hydrophobicity for upright growth and transportation of water and nutrients through the vascular system (Whetten & Sederoff, 19951995).
The main building blocks of lignin are three monolignols, that is p‐coumaryl, coniferyl and sinapyl alcohol, which are polymerised via oxidative coupling into p‐hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units in lignin polymers, respectively. Lignin composition varies considerably among plant taxa and species (Boerjan et al. , 20032003). For example, lignin in gymnosperms is mainly composed of G units, whereas lignin in angiosperm dicots consists principally of G and S units (Boerjan et al. , 20032003). In monocots, especially in grasses, lignin incorporates G and S units partially decorated with γ‐acyl groups (e.g. γ‐p‐coumarates and ferulates) along with tricin flavonoid units (Lam et al. , 20192019; Ralph et al. , 20192019). Typical gymnosperm and dicot lignins contain very low or undetectable levels of H units, whereas monocot lignins tend to incorporate larger amounts of H units, albeit at still low levels (Lapierre, 19931993; Gui et al. , 20112011). As different types of lignin monomers can form different types of intermonomeric linkages in lignin polymers (Ralph et al. , 20192019; Tobimatsu & Schuetz, 20192019), the composition of lignin monomers is likely to be the key determinant of various properties of lignin and cell walls as a whole. It is believed that lignin composition varies among different cell types, such as in vessels and fibres (Boerjan et al. , 20032003; Nakashima et al. , 20082008), although the quantitative determination of cell‐type‐specific lignin composition is still challenging.
Conversely, presence of lignin in cell walls imparts recalcitrance in deconstruction of the wall materials for pulping and biofuel production (Weng et al. , 20082008; Pauly & Keegstra, 20102010). To reduce such cell wall recalcitrance, a great deal of interest has been invested to engineer lignin to reduce lignin content or modify lignin composition (Li et al. , 20032003; Chen & Dixon, 20072007; Leple et al. , 20072007; Shadle et al. , 20072007; Voelker et al. , 20102010; Gui et al. , 20112011; Eudes et al. , 20122012; Van Acker et al. , 20142014; Umezawa, 20182018; Wang et al. , 20182018). However, the lignin‐modified plants often exhibit growth defects which consequently affect biomass production, thus limiting the benefits of lignin modification (Chen & Dixon, 20072007; Voelker et al. , 20102010; Gui et al. , 20112011; Bonawitz et al. , 20142014; Van Acker et al. , 20142014). Therefore, developing optimised strategies to manipulate lignin biosynthesis without yield penalty is crucial for overcoming the adverse effects of lignin modification on plant growth.
Lignin biosynthesis‐related transcription factor 1 (LTF1), belonging to the R2R3‐MYB subfamily, binds to the promoter of lignin biosynthesis genes to represses lignin biosynthesis in Populus (Gui et al. , 20192019). Several members in this subfamily across various plant species are reported to play a role in regulation of lignin biosynthesis (Zhao & Dixon, 20112011; Rao & Dixon, 20182018; Rao et al. , 20192019). LTF1 shares a high degree of sequence homology with AtMYB4 in Arabidopsis, which is involved in regulation of the phenylproponoid metabolism pathway (Jin et al. , 20002000). In Populus, LTF1 can be phosphorylated in response to external stimuli, however the phosphorylation sites are not conserved in AtMYB4 (Gui et al. , 20192019). The phosphorylated LTF1 is degraded which leads to the release of its suppression on lignin biosynthesis and thus the phosphorylation‐null LTF1 can convey a persistent suppression of lignin biosynthesis. In this study, we developed a new strategy to modify lignin biosynthesis through a cell‐type‐specific control of the expression of phosphorylation‐null LTF1. The engineered LTF1 was introduced into Populus under the control of vessel‐specific and fibre‐specific promoters, respectively. Fibre‐specific suppression of lignin biosynthesis resulted in improvement of biomass quality without growth defects. In‐depth lignin analysis of the transgenics with cell‐type‐specific lignin modification revealed new insights into how cell‐type‐specific lignin biosynthesis is related to plant growth.
Populus deltoides × P. euramericana cv ‘Nanlin895’ was used in this study. About 2 kb of the PdXCP1 (Potri.004G207600) promoter region and 1.8 kb of the PdDUF579‐9 (Potri.005G141300) promoter region (Song et al. , 20142014) were cloned from Populus . After sequencing, the correct promoter fragments were fused with the 35S minipromoter (regions from −90 to + 8) (Benfey & Chua, 19901990) and subcloned into a pCAMBIA2300‐GUS vector upstream of uidA (GUS). The constructed pCAMBIA2300:PdDUF579‐9p‐35S mini‐GUS and pCAMBIA2300:PdDUF579‐9p‐35S mini‐GUS vectors were transferred into Agrobacterium strain GV3101 for Populus transformation. For cell‐type‐specific expression of phosphorylation‐null LTF1 (LTF1AA) in Populus, the full‐length of LTF1 (Potri.004G174400) was mutated to LTF1AA and fused with the tag 3Flag and then subcloned into binary vectors pCAMBIA2300:PdXCP1p‐35S mini‐GUS and pCAMBIA2300:PdDUF579‐9P‐35S mini‐GUS , respectively. Primers used in this study are listed in Supporting Information Table S1. The obtained pCAMBIA2300:PdXCP1P‐35S mini‐LTF1AA‐3Flag and pCAMBIA2300:PdDUF579‐9P‐35S mini‐LTF1AA‐3Flag vectors were mobilised into Agrobacterium strain GV3101 for genetic transformation according to the procedure (Li et al. , 20032003).
Trees were grown in a phytotron under conditions of 60% relative humidity, 12 h photoperiod at 9000 lux, and 24°C constant temperature during young stage, and then grown in a glasshouse for a further characterisation or transplanted in a field at Shanghai (Crop cultivation and breeding station, Shanghai Institute of Plant Physiology and Ecology) (39.937128°N, 121.122304°E) for field test (from March 2018 to March 2019). Trees were clonally propagated through cutting and their morphological parameters, including plant height, stem diameter, internode length, leaf blade length and width, and petiole length were measured in 2‐month‐old trees. Plant height was determined as the shortest distance between shoot tip and stem base; stem diameter was measured 5 cm up from the stem base; internode length, leaf blade length and width and petiole length were measured from the 11th to the 20th internodes counted from the shoot top.
Total RNA was extracted from the stem of Populus at the 2‐month‐old stage using a total RNA kit following the manufacturer's instructions (Omega). Transcript levels of LTF1 and secondary cell wall biosynthesis‐related genes were determined, as described previously, using a MyiQ Real‐time PCR System (Bio‐Rad). The gene expression data were normalised using the Populus actin2 gene (Potri.001G309500). Primer pairs used for real‐time PCR are as described previously (Gui et al. , 20192019).
Trees were collected to measure biomass at the 2‐month‐old stage when grown in a phytotron. To minimise the potential transpiration impact of plant weight measurement, a stem was obtained by quickly stripping off the leaves and measuring the fresh weight within 2 min. Roots were collected and washed with water to remove soil and weighed after sucking surface water from the root. After measurement of the fresh weight, all plant fractions were dried in a forced‐draft oven at 55°C for 48 h, followed by measurement of their dry weight.
The hydraulic conductance of the stem xylem in 2‐month‐old Populus trees was measured using a high‐pressure flow meter (HPFM, Dynamax) following the manufacturer's instructions. In brief, the same length of the aerial part of trees from top to bottom (about 25 cm) was collected by cutting and quickly inserting into water, and then fixing into the HPFM instrument adaptor. The conductivity and flow of water were measured using a quasi‐steady‐state flow meter. To minimise the potential impact of diurnal periodicity on hydraulic conductance, all measurements were taken between 10:00 am and 3:00 pm in the phytotron in which the trees were grown (with 60% relative humidity and 24°C constant temperature).
Phloroglucinol‐HCl staining was performed as described previously (Gui et al. , 20112011). In brief, sections were stained with 1% phloroglucinol (w/v) in 12% HCl for 5 min and immediately observed under a microscope (Olympus BX53). Toluidine blue staining of paraffin‐embedded tissue was performed as described previously (Gui et al. , 20162016). GUS staining was performed as described (Gui et al. , 20112011). Free‐hand cross‐sections of stems were incubated with a staining solution (100 mM NaPO4 (pH 7.0), 10 mM ethylene diamine tetraacetic acid (EDTA), 2 mM 5‐bromo‐4‐chloro‐3‐indolyl‐β‐GlcA, 0.5 mM K4Fe(CN)6, 0.5mM K3Fe(CN)6, 0.2% Triton X‐100, and 20% methanol) at 37°C. The reaction was stopped, and chlorophyll was extracted using 75% ethanol. For immunolocalisation, samples were fixed with acetone, paraffin‐embedded and immunolocalised with GUS‐specific antibodies, as described previously (Gui et al. , 20142014).
Cell wall thickness was observed by transmission electron microscopy as described (Song et al. , 20102010). In brief, the eight stem internodes were collected and cut into 1.5–2 mm lengths and soaked in fixation buffer (3% paraformaldehyde and 0.5% glutaraldehyde in 0.1 M PBS, pH 7.4) and further fixed with 0.5% osmic acid. Then, the tissues were dehydrated and embedded in LR white resin (Polysciences, Warrington, PA, USA). Ultrathin sections were staining with uranyl acetate and lead citrate, and observed under an electron microscope (H‐7650; HITACHI, Kyoto, Japan) at 80 kV.
Xylem tissue of debarked stem from 2‐month‐old trees was collected and dried at 65°C. The dried tissue was ground to a fine powder by ball milling and used to prepare alcohol insoluble residues, according to Song et al. (20162016). The pretreated cell wall residues were used to determine lignin content using the Klason procedure, as described (Gui et al. , 20112011). For thioacidolysis and two‐dimensional (2D) nuclear magnetic resonance (NMR), dried xylem tissue was ground to fine powders by ball milling and extracted sequentially using water and 80% (v/v) ethanol to obtain extractive‐free cell wall residues (CWRs) (Tarmadi et al. , 20182018). Analytical thioacidolysis on CWR samples was performed as described previously (Yamamura et al. , 20122012) and the released lignin monomers were derivatised with N,O‐bis(trimethylsilyl)acetamide and quantified using gas chromatography‐mass spectrometry (GC‐MS; QP‐2010 Plus, Shimadzu, Kyoto, Japan) using 4,4′‐ethylenebisphenol as an internal standard (Yue et al. , 20122012). For 2D NMR analysis, CWRs were further ball‐milled and digested with crude cellulase (Cellulysin; Calbiochem, La Jolla, CA, USA) to obtain lignin‐enriched CWRs, as described previously (Tobimatsu et al. , 20132013; Lam et al. , 20172017). The lignin‐enriched CWRs (c. 25 mg) were then dissolved in dimethylsulfoxide‐d6/pyridine‐d5 (4 : 1, v/v) and subjected to NMR analysis. NMR spectra were acquired on a Bruker Biospin Avance™ III 800US system (Bruker Biospin, Billerica, MA, USA) equipped with a cryogenically cooled 5‐mm TCI gradient probe. Adiabatic 2D HSQC experiments were carried out using the standard implementation with parameters described in Mansfield et al. (20122012). Data processing and volume integration analysis was conducted using Bruker TopSpin 3.1 software (Bruker Biospin) with procedures described previously (Lam et al. , 20172017; Tarmadi et al. , 20182018).
Fresh stems of 8‐month‐old Populus trees grown in a glasshouse were prepared for mechanical strength analysis. Mechanical properties of stem were measured by a three‐point bending test using a mechanical testing machine (HY‐0580, http://www.hengyiyiqi.com) according to (Xi et al. , 20172017). Modulus of rupture (MOR) in bending was calculated according to Kern et al. (20052005).
Xylem cell wall saccharification assays were performed as described (VanAcker et al. , 20162016). Samples were collected after 3, 6, 9, 12, 24, 36, 48 or 60 h incubation with appropriate cellulase and β‐glucosidase and then the amount of glucose released was measured. Total glucan was determined using the dinitrosalicylic acid (DNS) method according to Liu et al. (20132013). To calculate total tree stem sugar yield, the saccharification of glucose yield was also presented based on total biomass of each plant stem.
In our previous study, lignin biosynthesis in Populus was persistently suppressed when LTF1 was modified to become a phosphorylation‐null form (LTF1AA) (Gui et al. , 20192019). Under the control of a constitutive promoter (cauliflower mosaic virus 35S promoter, CaMV35S), lignin biosynthesis suppression (LS) by overexpressing LTF1AA (designated as 35S‐LS ) caused severe growth defects. The transgenics displayed not only a retarded stature, shortened internodes and stem diameter, but also had short petiole length and small‐sized leaves (Fig. S1a–g). The constitutive suppression of lignin biosynthesis through overexpressing phosphorylation‐null LTF1 (LTF1AA) affected growth.
To investigate whether suppression of lignin biosynthesis in different cell types affects plant growth, we attempted to modify lignin biosynthesis under the control of vessel‐specific or fibre‐specific promoters. After examination of a number of cell‐type‐specific promoters, the promoter from PdDUF579‐9 (Potri.005G141300), which is specifically expressed in xylem fibres (Song et al. , 20142014), was used for fibre‐specific modification. The PdDUF579‐9 promoter (designated PdDUF579‐9p) was cloned and combined with a 35S minimal promoter to construct the PdDUF579‐9p‐35Smini synthetic promoter. When the synthetic promoter was fused to a β‐glucuronidase (GUS) gene (PdDUF579‐9p‐35Smini‐GUS ) (Fig. S2a) and transformed into Populus , it displayed fibre‐specific activity (Fig. S2b) For vessel‐specific modification, a promoter fragment (2068 bp) from Populus XYLEM CYSTEINE PROTEASE1 (XCP1; Potri.004G207600) (Funk et al. , 20022002) was cloned and combined with a 35S minimal promoter to generate PdXCP1p‐35Smini promoter (Fig. S2a). We confirmed that PdXCP1p‐35Smini showed an activity specific for xylem vessels when it was fused with the GUS gene (PdXCP1p‐35Smini‐GUS) and transformed into Populus (Fig. S2c).
The cell‐type‐specific expression of PdDUF579‐9p‐35Smini‐GUS and PdXCP1P‐35Smini‐GUS in Populus was further verified by immunolocalisation with GUS‐specific antibodies. PdDUF579‐9P‐35Smini‐GUS was specifically localised in fibre cells (Fig. S2d), while PdXCP1P‐35Smini‐GUS was specifically localised in vessel cells (Fig. S2e); this localisation was highly consistent with the GUS staining results (Fig. S2b,c). In addition, the localisation region of the two promoter activity was in agreement with the high expression of DUF579‐9 and XCP1 in the developing xylem region (Fig. S2e,f), according to the information documented in the AspWood resource (Sundell et al. , 20172017).
Then the two synthetic promoters were fused with LTF1AA to generate PdDUF579‐9p‐35Smini‐LTF1AA and PdXCP1p‐35Smini‐LTF1AA constructs, which were transferred to Populus. Transgenic lines harbouring PdXCP1p‐35Smini‐LTF1AA were designated as V‐LS and transgenic lines harbouring PdDUF579‐9p‐35Smini‐LTF1AA were designated as F‐LS. Thirty‐five independent F‐LS transgenic lines with similar morphologies were identified and two representative transgenic lines, F‐LS #12 and F‐LS #27 were clonally propagated for detailed characterisation. Meanwhile, 67 independent V‐LS transgenic lines were generated that displayed dramatic morphological changes. Two representative transgenic lines, V‐LS #45 and V‐LS #51 , were clonally propagated and used for further characterisation. The transgenic lines were first grown in a phytotron for 2 months (Fig. 1a) and then moved into a glasshouse (Fig. 1b). Compared with the wild‐type control, F‐LS transgenics did not show any noticeable growth change. However, the V‐LS transgenics displayed severe dwarfism, small and dark‐green leaves, and more branching (Fig. 1a,b). Specifically, the V‐LS transgenics were reduced by 50–65% in height, 35–40% in stem diameter, 60–65% in internode length and 60–70% in leaf size compared with the control (Fig. 1c,h). Consistently, stem biomass production was reduced by more than 50% in V‐LS transgenics when compared with the control, whereas F‐LS transgenics showed slight increases in both fresh and dry biomass production (Fig. S3a,d). In addition, leaf and root biomass were reduced substantially in the V‐LS transgenics (Fig. S3b,c,e,f). Collectively, fibre‐specific modification of lignin biosynthesis showed normal growth, while vessel‐specific modification resulted in growth defects and biomass reduction.
Vessels and fibres in xylem constitute a system of long‐distance water transportation and mechanical support to sustain plant growth (Boerjan et al. , 20032003). To examine how the cell‐type‐specific modification of lignin biosynthesis in xylem affected system functions such as water transportation, the hydraulic conductance of the stem was measured using a high‐pressure flow meter (HPFM) (Dynamax). The V‐LS transgenics showed a 50% reduction of hydraulic conductance, but no change was detected in the F‐LS transgenics (Fig. 1i). Meanwhile, the 35S‐LS transgenics showed a 55% reduction in stem hydraulic conductance (Fig. 1i). These results suggested that suppression of lignin biosynthesis in vessels affected stem water conductance and may be crucial for plant growth. In addition, we examined stem mechanical strength. A bending test showed the order of the stem bending force as: wild‐type > F‐LS > V‐LS transgenics (Fig. S4a). Compared with the wild‐type, the MOR displayed a reduction in the F‐LS transgenics but no significant difference in the V‐LS transgenics. This should be attributed to the smaller stem diameter in the V‐LS transgenics (Fig. S4b). The data suggested that lignin deposition in xylem vessels and fibres may have different effects on water conductance and mechanical properties.
Both V‐LS and F‐LS transgenics showed a four‐ to six‐fold increase in the LTF1 transcripts (measurements included the LTF1 and LTF1AA transcripts) compared with the control (Fig. 2a). Then, we measured the expression of secondary cell wall (SCW) biosynthesis‐related genes (lignin biosynthesis genes: 4CL1, CAld5H, COMT2 and CCoAOMT1; cellulose biosynthesis genes: CESA4, CESA7 and CESA8; and xylan biosynthesis genes: GT43B and GT8D). The transcript abundances of the key lignin biosynthesis genes 4CL1, COMT2 and CCoAOMT1 were substantially suppressed in both V‐LS and F‐LS transgenics (Fig. 2a), while the transcript abundances of cellulose and xylan biosynthesis genes were not significantly suppressed, suggesting a specific suppression of lignin biosynthesis (Fig. 2b). Interestingly, transcript abundances of the lignin biosynthesis genes in developing xylem were less suppressed in the V‐LS transgenics than in the F‐LS transgenics (Fig. 2a). This difference may reflect the ratio of vessel and fibre cell populations in xylem tissue. The expression of CAld5H, which encodes a key enzyme for S‐lignin biosynthesis (Meyer et al. , 19981998; Osakabe et al. , 19991999; Li et al. , 20032003), was reduced in the F‐LS transgenics but not in the V‐LS transgenics (Fig. 2a). These results indicated that expression levels of lignin biosynthesis genes were differentially modified through fibre‐specific or vessel‐specific manners.
To examine SCWs in xylem, cross‐sections of the stems at the 11th internode were dissected. In the F‐LS transgenics, lignin deposition was reduced in fibre cell walls with no apparent changes in vessel walls as determined by lignin staining using phloroglucinol‐HCl (Fig. 3a,c,d,f). By contrast, the V‐LS transgenics developed thin‐walled and deformed vessels and lignin deposition in fibre cell walls showed no change compared with the wild‐type (Fig. 3a,b,d,e). In the F‐LS transgenics, cells were well organised in xylem while, in the V‐LS transgenics, xylem cells were disorganised with deformed vessels (Fig. 3g–i).
The SCW thickness in the V‐LS transgenics was reduced by 40–50% in vessels with no change in fibres (Fig. 3j,k,m,n). By contrast, the F‐LS transgenics showed no change in SCW thickness in vessels but the SCW thickness was reduced by 35–45% in fibres (Fig. 3j,l–n). These results indicated that lignification and SCW formation were modified in vessels and fibres in a cell‐type‐specific manner. In comparison, we also examined xylem cells in the 35S‐LS transgenics. The 35S‐LS transgenics displayed distorted and collapsed vessel cells and larger but less lignified fibre cells (Fig. S5a,b). Both vessels and fibres were modified to thinner SCWs (Fig. S5c,d). The SCW thickness was substantially decreased (Fig. S5e,f) by 40% and 45% in xylem vessels and fibres, respectively (Fig. S5g,h). In addition, the majority of the vessels in the 35S‐LS transgenics displayed a wall‐deformed structure (Fig. S5i). Constitutive suppression of lignin biosynthesis resulted in defective SCW of both vessels and fibre cells in xylem.
Lignin deposition was modified in the vessels of the V‐LS transgenics and in the fibres of the F‐LS transgenics. Lignin content and monolignol composition of CWR samples prepared from the control and transgenic xylem tissues were measured using the Klason and thioacidolysis methods, respectively. The Klason lignin content was reduced by 43% in the F‐LS transgenics and by 16% in the V‐LS transgenics (Table 1). Meanwhile, lignin in the 35S‐LS transgenics was more drastically reduced by 56% (Table 1); this might reflect the lignin modification in both vessels and fibres. The F‐LS transgenics with the lignin reduction by 43% did not show any apparent growth defects (Fig. 1a,b). However, the V‐LS transgenics displayed, similarly to the 35S‐LS , serious growth defects (Figs 1, S1), although the lignin content in the V‐LS transgenics was more mildly reduced by 16%. This result suggests that cell‐type‐specific lignin deposition plays a critical role in plant growth.
|Sample||Klason lignin (% CWR)||Thioacidolysis‐derived monomer composition (μmol g−1 CWR)||Total H + G + S||Thioacidolysis‐derived monomer reduction (%)||S : G ratio|
|Control||22.18 ± 0.63a||1.64 ± 0.14||293.62 ± 23.43||354.09 ± 57.22||649.34 ± 63.99a||‐||‐||1.21 ± 0.20b|
|35S‐LS||9.85 ± 0.51d||1.80 ± 0.15||184.58 ± 28.13||76.34 ± 13.57||262.71 ± 41.80d||37a||78a||0.41 ± 0.01d|
|V‐LS||18.56 ± 0.73b||1.06 ± 0.03||238.77 ± 12.90||320.06 ± 15.34||559.88 ± 28.19b||18b||9c||1.34 ± 0.01a|
|F‐LS||12.54 ± 0.41c||1.50 ± 0.06||202.39 ± 18.38||168.82 ± 19.52||372.72 ± 37.31c||31a||52b||0.83 ± 0.04c|
Analytical thioacidolysis, which releases quantifiable H‐, G‐, and S‐type monomeric degradation products via cleavage of β–O–4 linkages in lignin polymers (Lapierre et al. , 19861986), determined substantially altered lignin subunit composition and S : G ratio in the three kinds of the transgenic lines. Consistent with the Klason lignin content data (Table 1), the total yields of the lignin‐derived monomeric products released from the three transgenic lines were all reduced compared with that determined for the wild‐type control. The lignin‐derived monomeric products were principally composed of G‐ and S‐types with much lower amounts of H‐type (about 0.2–0.6%). Our data suggested that G‐ and S‐type products were disproportionately reduced in the three transgenic lines. For the V‐LS transgenics, G‐type products were reduced by 18% and S‐type products by 9%, leading to an increase in the S : G ratio. For the F‐LS transgenics, conversely, G‐type products were reduced by 31% and S‐type products by 52%, resulting in a decrease in the S : G ratio. Given that vessel and fibre cell walls in typical dicot species are relatively enriched in G‐ and S‐lignin units, respectively (Boerjan et al. , 20032003; Nakashima et al. , 20082008), such increase and decrease of the S : G ratio in the V‐LS and F‐LS lines are in line with our idea that their lignin reduction is cell type specific for vessels and fibres, respectively (Fig. 3a–f). For the 35S‐LS transgenics, G‐type products were reduced by 37% and S‐type products by 78%. The larger reduction in S‐type products may reflect a higher percentage of fibres than vessels in xylem. Overall, our data suggest that the majority of lignin in xylem is from fibres, in which suppression of lignin biosynthesis primarily caused a reduction in S units, but suppression of lignin biosynthesis in vessels was mostly related to G‐unit reduction.
To further examine lignin composition and structure, we performed solution‐state two‐dimensional (2D) nuclear magnetic resonance spectroscopy (NMR) analysis on lignin‐enriched CWR samples prepared via enzymatic removal of cell wall polysaccharides. The aromatic subregions of the short range 1H–13C correlation (HSQC) spectra displayed contour signals from G and S aromatic units along with those from p‐hydroxybenzoates (p Bz) acylating a fraction of poplar lignin (Venverloo, 19711971; Morreel et al. , 20042004; Stewart et al. , 20092009) and cinnamyl alcohol end‐groups (X1 ) (Fig. 4). Volume integration of C2–H2 signals from G and S units suggested that the S : G ratio was increased in the V‐LS transgenics (S : G, 1.2), but decreased in the F‐LS (S : G, 0.7) and 35S‐LS (S : G, 0.3) transgenics, compared with the control (S : G, 1.0) (Fig. 4). The change in the S : G ratio in each transgenic line is overall in line with the results of thioacidolysis, as described above (Table 1). In addition, we observed that the relative intensity of pBz signals was increased in the spectrum of the V‐LS lignin, but reduced in the spectra of the F‐LS and 35S‐LS lignins. As p‐benzoate preferentially acylates S units over G units in typical poplar lignin (Stewart et al. , 20092009), the data further corroborate the shifts of S : G ratio in the three transgenic lines.
The aliphatic subregions of the HSQC spectra display countersignals from the major intermonomeric linkages, such as β‐aryl ether (β–O–4, I), phenylcoumaran (β–5, II), resinol (β–β, III) and spirodienone (β–1, IV), in the lignin polymer. Volume integration of Cα–Hα signals suggested that the V‐LS transgenics had proportionally decreased β–5 units, whereas F‐LS and 35S‐LS had conversely increased β–5 units (Fig. 5). Because β–5 linkages can be only formed in G units derived from the polymerisation of coniferyl alcohol, the increase in β–5 units is consistent with the decreased S : G ratios in the 35S‐LS and F‐LS transgenics and in agreement with previous studies on transgenic plants with altered S : G ratios (Stewart et al. , 20092009; Anderson et al. , 20152015; Takeda et al. , 20172017; Takeda et al. , 20192019). Taken together, both thioacidolysis and 2D NMR data support our idea that G‐rich vessel and S‐rich fibre lignins are precisely altered in the transgenics obtained in this study.
As the F‐LS and V‐LS transgenic Populus displayed different growth under laboratory conditions, we further evaluated their growth performance in field. The transgenic trees were grown in a field in Shanghai (Crop Cultivation and Breeding Station, SIPPE, China) for a full growth season. F‐LS transgenics showed a similar growth performance with the control trees in height, stem diameter and aboveground biomass (Fig. 6a,b). However, V‐LS transgenics displayed retarded growth with reduction in height, stem diameter and aboveground biomass (Fig. 6a,b). More specifically, plant height was reduced by 56–63%, stem diameter by 39–48%, and aboveground biomass by 75–89% in the V‐LS transgenics compared with the control (Fig. 6c,e). These results showed that fibre‐specific modification of lignin biosynthesis maintained vigorous growth in fields, whereas vessel‐specific modification resulted in a growth penalty with biomass reduction. We further observed cell morphology and lignin staining of xylem tissues of the field‐grown trees. The V‐LS transgenics contained deformed or collapsed vessels with regular and normally lignified fibre cell walls (Figs 6f,g, S6a,b,d,e,g,h). By contrast, F‐LS transgenics contained less lignin in fibre cell walls, but lignin deposition was normal in vessel walls (Figs 6f,h, S6a,c,d,f,g,i). Lignin modification in specific cell types, therefore, differentially affected field growth performance.
In addition, we carried out saccharification analysis using wood stem materials. The 35S‐LS and F‐LS transgenics showed a higher efficiency of glucose release than the V‐LS transgenics (Fig. 6i). After 60 h of enzyme digestion, glucose release from the 35S‐LS and F‐LS wood material was increased by 70% and 45%, respectively, compared with the control, whereas glucose release from the V‐LS material was not significantly different from the control (Fig. 6i). It is likely that the more drastic lignin reductions in 35S‐LS and F‐LS wood materials contributed to the improved saccharification efficiency. Although saccharification efficiency was increased in 35S‐LS lines, the total glucose yield per plant was significantly reduced due to the dramatic decrease in biomass yield. As shown in Fig. 6(j), the relative glucose yield per plant reflected a combination of saccharification efficiency, glucan content and plant biomass yield. The F‐LS transgenics displayed a relative increase of glucose yield per plant by 67% compared with the control. By contrast, glucose yield per plant was decreased by 65% with 35S‐LS transgenics and by 60% with V‐LS transgenics. These results indicated that lignin modification through a fibre‐specific manner can be applicable for improvement of wood biomass production without growth penalty.
Lignin is synthesised specifically in the SCWs of vascular plants, fortifying the mechanical strength of walls and providing wall hydrophobicity for long‐distance transportation of water (Weng & Chapple, 20102010). During xylem differentiation in angiosperms, lignin is synthesised in fibre and vessel cells. Much evidence has accumulated to elucidate lignin biosynthesis in plants (Boerjan et al. , 20032003; Barros et al. , 20152015; Meents et al. , 20182018; Vanholme et al. , 20192019). However, the role of lignin biosynthesis in different cell types is not fully understood.
In this study, we employed a lignin‐related transcription factor gene LTF1, which had been modified to convey persistent suppression of lignin biosynthesis (Gui et al. , 20192019). The engineered LTF1 (LTF1AA) allowed us to obstruct lignin biosynthesis in specific types of cells under the control of cell‐type‐specific promoters. Constitutive expression of LTF1AA (35S‐LS) suppressed lignin biosynthesis in xylem tissue with development of defective SCWs in both xylem vessels and fibres, resulting in severe growth defects with retarded stature, shortened internodes, stem diameter and petiole length, and the small‐sized leaves of the 35S‐LS transgenic trees (Fig. 7). When lignin biosynthesis was suppressed by overexpression of LTF1AA in vessels (V‐LS), the transgenic trees developed defective SCWs in vessel and displayed a similar growth as the found for 35S‐LS (Fig. 7). However when lignin biosynthesis was suppressed by overexpression of LTF1AA in fibres (F‐LS), the F‐LS transgenic trees contained effective vessels and grew normally (Fig. 7). These results suggested that lignin biosynthesis in different cell types plays distinct roles in plant growth.
Disruptions to lignin biosynthesis often causes abnormal lignin deposition in lignified cells, such as xylem vessels, fibres and other cells, and can lead to abnormal plant growth and development including dwarfed morphology and defects in fertility (Leple et al. , 20072007; Shadle et al. , 20072007; Yang et al. , 20072007; Nakashima et al. , 20082008; Voelker et al. , 20102010; Gui et al. , 20112011; Van Acker et al. , 20142014). For example, lignin biosynthesis in endothecium cells is required for anther dehiscence and pollen release for pollination (Yang et al. , 20072007). Pod shatter also requires specific lignification of the valve margin cells to create tension for valve separation from the replum (Liljegren et al. , 20002000).
Modification of lignin biosynthesis in various type cells can disrupt proper cell function (Barros et al. , 20152015). Evidence showed that, for example, whereas mutations of lignin biosynthetic enzyme genes such as cinnamic acid 4‐hydroxylase (C4H) (Schilmiller et al. , 20092009), cinnamoyl coenzyme A reductase (CCR) (Jones et al. , 20012001) and caffeoyl shikimate esterase (CSE) (Vanholme et al. , 20132013) in Arabidopsis resulted in abnormal lignin biosynthesis, vessel cell wall defects and slow growth, particular restoration of these enzyme functions in vessels helped to rebuild vessel integrity and growth performance (Yang et al. , 20132013; Vargas et al. , 20162016; De Meester et al. , 20182018). In addition, incorporation of unconventional monolignols into lignin polymers via lignin engineering improved biomass digestibility without growth penalty in Populus (Wilkerson et al. , 20142014; Cai et al. , 20162016).
In this study, hydraulic conductance in the stem of transgenic plants was reduced after the repression of lignin biosynthesis in vessels, while fibre‐specific suppression of lignin biosynthesis did not affect hydraulic conductance. Therefore, our data demonstrate that lignin deposition in xylem vessels is required for vessel function in long‐distance transportation of water that in turn is essential for plant normal development and growth.
In the cell‐type specifically modified Populus, lignin content and lignin monomers composition were changed greatly. Lignin content was reduced by 56%, 43% and 16%, respectively, in the xylem tissue of 35S‐LS, F‐LS and V‐LS transgenics (Table 1). The reduction of lignin content reflected lignin biosynthesis, which was suppressed in both vessels and fibres under the control of a CaMV 35S promoter (35S‐LS), in fibres under the control by a fibre‐specific promoter (F‐LS), and in vessels under the control of a vessel‐specific promoter (V‐LS). In Populus, there is a higher proportion of xylem fibres than vessels in secondary xylem tissue (Takata et al. , 20192019); this could explain why suppression of lignin biosynthesis in fibres resulted in larger lignin reduction than that in vessels (Table 1).
It is believed that lignin monomer composition varies among cell types. The current understanding of cell‐type‐specific lignin composition is primarily based on evidence provided by ultraviolet (UV) light microscopy, FT‐IR microspectroscopy or X‐ray analysis (Saka et al. , 19881988; Foster et al. , 20102010; Gorzsas et al. , 20112011). However, more precise determination of lignin monomer composition in different cell types still remains to be addressed. The specific suppression of lignin biosynthesis in different types of cells in the secondary xylem tissue, and examination of the expression of lignin biosynthetic genes and the lignin composition in the transgenics illustrated a quantification of lignin monomers in fibres and vessels. Interestingly, CAld5H, the key enzyme for S‐lignin biosynthesis (Meyer et al. , 19981998; Osakabe et al. , 19991999; Li et al. , 20032003), was not suppressed in the V‐LS transgenics, but significantly suppressed in the F‐LS transgenics, suggesting that S‐lignin biosynthesis may take place primarily in fibre cells. This idea was further verified by thioacidolysis and 2D NMR analyses. Suppression of vessel lignin biosynthesis resulted in more G‐unit reduction, whereas suppression of fibre lignin biosynthesis conversely resulted in more S‐unit reduction (Table 1; Fig. 4). In line with these observations, 2D NMR further revealed that phenylcoumaran (β–5) linkages unique in G‐lignin units were proportionally reduced in the transgenics of vessel‐specific lignin suppression (V‐LS) whereas the transgenics of fibre‐specific lignin suppression (F‐LS) produced lignin with more β–5 units. These results provided a line of quantitative evidence suggesting that, in xylem tissue vessel, cells are enriched in G‐lignin while fibre cells are predominantly composed of S‐lignin.
Lignin plays various roles in different cell types for plant growth and development. Meanwhile, lignin is also a limiting factor in utilisation of the most abundant and renewable lignocellulosic biomass for production of fibre materials and biofuels (Chen & Dixon, 20072007; Vanholme et al. , 20132013; Van Acker et al. , 20142014). To overcome this lignin barrier in biomass deconstructions, a great deal of interest has been invested in modification of lignin biosynthesis through manipulating key genes encoding the pathway enzymes and transcription factors involved in lignin biosynthesis. For example, reduction in lignin content has been achieved by downregulation of 4‐coumarate:coenzyme A ligase (4CL) in poplar (Hu et al. , 19981998; Voelker et al. , 20102010), caffeic acidO‐methyltransferase (COMT) in switchgrass (Fu et al. , 20112011), 4‐coumarate 3‐hydroxylase (C3H) and hydroxycinnamoyl CoA:shikimate hydroxycinnamoyl transferase (HCT) in alfalfa (Shadle et al. , 20072007). In addition, combinational modification of multiple lignin biosynthesis genes to alter lignin monomer composition and content were achieved in poplar and rice (Li et al. , 20032003; Gui et al. , 20112011).
Although the strategies to modify lignin biosynthesis genes to reduce lignin content and increase biomass saccharification efficiency have significant potential for biotechnology applications, traditional strategies for lignin reduction through a constitutive or a tissue‐specific manner often cause stunted growth and reductions in biomass yield that impose considerable limitations on their application. To optimise the benefit from lignin modification, we carried out a precise regulation of lignin biosynthesis in specific cell types using LTF1, a transcription factor. LTF1 is able to modulate expression of the multiple key genes in the lignin biosynthesis pathway and regulate the entire metabolic flux to lignin deposition (Gui et al. , 20192019). In addition, the engineered LTF1 (LTF1AA) is more stable and can provide a persistent suppression of lignin biosynthesis. Suppression of lignin biosynthesis in fibre cells improved biomass production without growth defects, whereas suppression of lignin biosynthesis in vessel cells caused growth defects and substantial reduction in biomass yield.
Lignin suppression in different cell types showed different growth performance, biomass yield and glucose yield through saccharification. Based on our analysis, the glucose yield after saccharification was increased by 67% in the transgenics with fibre‐specific suppression of lignin biosynthesis, while the transgenics with vessel‐specific suppression of lignin biosynthesis showed a 65% reduction in glucose yield. Modification of lignin in fibres displayed a significant potential to reduce cell wall recalcitrance in biomass conversion, as well as improve biomass production without growth penalty. Next, a larger scale of field test would be necessary to evaluate the applicability of this approach of lignin engineering. Nevertheless, this study opens an avenue to more precisely manipulate lignin deposition and improve lignocellulosic biomass property and production.
We thank Xiaoyan Gao, Jiqin Li and Zhiping Zhang for their support in transmission electron microscopy and Hironori Kaji and Ayaka Maeno for their support in NMR experiments. Part of this study was conducted using the facilities in the DASH/FBAS at RISH and the NMR spectrometer in the JURC at ICR, Kyoto University. This work was supported by the National Natural Science Foundation of China (grant no. 31630014), Chinese Ministry of Agriculture (grant no. 2018ZX08020002), the Chinese Academy of Sciences (grant no. XDB27020104), the Youth Innovation Promotion Association CAS (grant no. 2017318), the Chinese Ministry of Science and Technology (grant no. 2016YFD0600104), the Japan Society for the Promotion of Science (JSPS; grant nos. 16K14958, 16H06198 and 17F17103), RISH, Kyoto University (Mission‐linked Research Funding, no. 2016‐5‐2‐1), and the China/Japan Scientific Cooperation Programme by NSFC/JSPS.