EXPLORING CAFFEYL-LIGNIN BIOSYNTHESIS IN Cleome hassleriana AND
POLYMERIZATION OF CAFFEYL ALCOHOL IN Arabidopsis thaliana
Aaron D. Harkleroad
Thesis Prepared for the Degree of
MASTER OF SCIENCE
UNIVERSITY OF NORTH TEXAS
December 201 9
APPROVED:
Richard A. Dixon, Major Professor Brian Ayre, Committee Member Rajeev Azad, Committee Member Jyoti Shah, Chair of the Department of Biological Sciences Su Gao, Dean of the College of Science Victor Prybutok, Dean of the Toulouse Graduate School Harkleroad, Aaron D. Exploring Caffeyl-Lignin Biosynthesis in Cleome hassleriana and Polymerization of Caffeyl Alcohol in Arabidopsis thaliana. Master of
Science (Biochemistry and Molecular Biology), December 2019, 47 pp., 2 tables, 15 figures, references, 43 titles.
C-lignin (caffeyl-lignin) is a novel linear lignin polymer found in the seed coats of several non-crop plants, notably Vanilla planifolia (Vanilla), Jatropha curcas (Jatropha), and Cleome hassleriana (Cleome). C-lignin has several advantages over normal G/S- lignin, found in the majority of lignocellulosic biomass, for valorization in the context of bioprocessing: less cross-linking to cell wall polysaccharides (less recalcitrant biomass), ordered linkages between monomers (homogeneous polymer), and no branching points
(linear polymer). These properties make C-lignin an attractive replacement for native lignin in lignocellulosic biomass crops.
The seed coats of Cleome hassleriana (Cleome) synthesize G-lignin during early seed maturation, then switch to synthesis of C-lignin during late maturation. This switch to C-lignin in Cleome seed coats is accompanied by loss of caffeoyl-CoA 3-O- methyltransferase (CCoAOMT) and caffeic acid 3-O-methyltransferase (COMT) activities, along with changes in transcript abundance of several lignin related genes.
The focus of this research thesis is to understand the biochemical changes leading to
C-lignin deposition in Cleome hassleriana seed coats, and to explore the ability of
Arabidopsis thaliana seedlings to polymerize caffeyl alcohol to C-lignin.
In this thesis, candidate transcripts were implicated in C-lignin biosynthesis by differential gene expression analysis of transcripts in seed coat tissues at 8-18 days after pollination (DAP) and in non-seed coat tissues. Three candidate genes were
selected for recombinant expression and their in vitro kinetic properties were measured with potential substrates. Of the three candidates, a cinnamyl alcohol dehydrogenase
(ChCAD5) was found to have high transcript levels during C-lignin formation and have a novel preference for converting caffealdehyde to caffeyl alcohol, the precursor of C- lignin. To determine if accumulation of caffeyl alcohol is sufficient for polymerization of
C-lignin, Arabidopsis seedlings grown in a xylem induction system were supplied caffeyl alcohol. Polymerization of caffeyl alcohol was not found to occur in this Arabidopsis system, suggesting the need for a C-lignin specific polymerization mechanism.
Copyright 2019
by
Aaron D. Harkleroad
ii ACKNOWLEDGMENTS
I would like to acknowledge the support and funding by UNT start-up funds, provided to the Dixon Laboratory, along with grant #1456286 from the National Science
Foundation, Integrated Organismal Systems.
This thesis is dedicated to my family and friends who provided support and encouragement throughout my graduate work. I would like to thank my wife, Jagruti
Harkleroad, for her confidence in my scholastic abilities and for providing inspiration during challenging times. To my mom and dad, thank you for believing in me and supporting my educational journey. Thanks to my friends for listening to my esoteric
“biology talk” with earnest interest and good humor; you have all been a great source of emotional support. Thanks to all my graduate student cohorts in the UNT biology program for their comradery and stimulating conversations, I will truly miss them.
I would like to extend my greatest appreciation to my major professor, Dr.
Richard Dixon, for his amazing patience and encouragement during my graduate work.
His critical thinking and openness to hypotheses has been instrumental in my research.
I also thank the members of my graduate committee for their critical analysis and feedback. I thank the many researchers who have assisted me in my research by providing materials and expertise- there are too many to name here, but I appreciate you all. I specifically thank Chunliu Zhuo and Fang Chen for supplying substrates, Chan
Man Ha for supplying Arabidopsis seeds, Xiaolan Rao for her RNA sequencing work, and Xiaoqiang Wang for his protein modeling work.
iii TABLE OF CONTENTS
Page
ACKNOWLEDGMENTS ...... iii
LIST OF TABLES ...... vi
LIST OF FIGURES ...... vii
CHAPTER 1. INTRODUCTION ...... 1 Biosynthesis of Monolignols ...... 3 Polymerization of Lignin ...... 7 Engineering Lignin Subunits ...... 8 Hypothesis and Objectives ...... 9 Methods Overview ...... 10 1.5.1 Candidate Gene Discovery by RNA Sequencing ...... 10 1.5.2 Recombinant Expression and Characterization of Target Proteins10 1.5.3 In vitro Polymerization Systems and Induction of Lignification in vivo ...... 12 1.5.4 Analysis of Lignin Presence, Amount and Composition ...... 14
CHAPTER 2. DISCOVERY OF CANDIDATE GENES RELATED TO C-LIGNIN BIOSYNTHESIS IN Cleome SEED COATS ...... 15 Objectives ...... 15 Methods ...... 15 2.2.1 RNA Sequencing and Assembly ...... 15 2.2.2 Analysis of Differential Gene Expression ...... 15 2.2.3 Homology Mapping to Arabidopsis ...... 16 2.2.4 Protein Modeling ...... 16 Results and Discussion ...... 16 2.3.1 RNA Sequencing ...... 16 2.3.2 Differential Gene Expression ...... 16 2.3.3 Comparison of Candidate Cleome Transcripts to Arabidopsis Homologs ...... 19 2.3.4 Protein Modeling of ChCAD5 ...... 19 Conclusion ...... 21
iv
CHAPTER 3. IN VITRO CHARACTERIZATION OF RECOMBINANT CANDIDATE ENZYMES ...... 22 Objectives ...... 22 Methods ...... 22 3.2.1 Cloning, Recombinant Protein Expression, and Enzyme Purification ...... 22 3.2.2 Kinetic Assays ...... 23 3.2.3 Mixed Reactions ...... 24 3.2.4 HPLC Analysis ...... 24 Results and Discussion ...... 25 3.3.1 Expression of Recombinant Proteins ...... 25 3.3.2 Kinetic Properties of ChCCR and ChCAD4 and 5 ...... 25 3.3.3 Mixed Reactions with ChCCR and ChCAD 4 or 5 ...... 29 Conclusions ...... 32
CHAPTER 4. POLYMERIZATION OF CAFFEYL ALCOHOL AND OTHER MONOLIGNOLS IN Arabidopsis SEEDLINGS ...... 33 Objectives ...... 33 Methods ...... 33 4.2.1 Xylem Induction in Arabidopsis Seedlings and Feeding of Monolignols ...... 33 4.2.2 Staining and Microscopic Observation of Seedlings ...... 34 4.2.3 Harvesting of Shoot Tissue and Preparation of Cell Wall Residues ...... 35 4.2.4 Lignin Thioacidolysis and Analysis of Monomer Composition by GC-MS ...... 35 Results and Discussion ...... 35 4.3.1 Growth of Arabidopsis Seedlings and Induction of Ectopic Xylem 35 4.3.2 Lignin Composition and Incorporation of Supplied Monolignols .... 38 Conclusions ...... 41
CHAPTER 5. OVERALL SUMMARY AND CONCLUSIONS ...... 42
REFERENCES ...... 44
v LIST OF TABLES
Page
Table 3.1: Primers used for cloning of candidate genes...... 23
Table 3.2: Kinetic parameters for recombinant enzymes with caffeoyl-CoA and feruloyl- CoA or caffealdehyde and coniferaldehyde...... 28
vi LIST OF FIGURES
Page
Figure 1.1. The three primary monolignols that polymerize to make lignin and the caffeyl alcohol monomer from which C-lignin is derived...... 1
Figure 1.2: The lignin biosynthesis pathway, from Tobimatsu et al. 2013 ...... 4
Figure 1.3: Lignin deposition and O-methyltransferase activity in the seed coat of C. hasselerania (from Tobimatsu et al. 2013) ...... 6
Figure 2.1: Transcript levels of ChCCR1 in Cleome ...... 18
Figure 2.2: Transcript levels of ChCAD4 (green) and ChCAD5 (red) in Cleome ...... 18
Figure 2.3: Protein alignment of ChCAD4 and 5 to AtCAD2 and 5 ...... 20
Figure 2.4: Model of active site, by Dr. Xiaoqiang Wang, of ChCAD5 (blue) showing the specific amino acids (orange) interacting with caffealdehyde (yellow) (Zhuo et al. 2019)...... 21
Figure 3.1: In vitro activity of ChCCR1 with feruloyl-CoA (red) and caffeoyl-CoA (blue)...... 26
Figure 3.2: In vitro activity of ChCAD4 with coniferaldehyde (red) and caffealdehyde (blue) ...... 27
Figure 3.3: In vitro activity of ChCAD5 with coniferaldehyde (red) and caffealdehyde (blue) ...... 28
Figure 3.4: Stacked bar graph of products of in vitro mixed enzyme/substrate reactions ...... 30
Figure 4.1: Arabidopsis seedlings grown on mesh disks floating in liquid medium ...... 36
Figure 4.2: TBO staining of Arabidopsis leaves from seedlings grown with bikinin (B, C, E, F, G, and H) and without bikinin (A and D) ...... 37
Figure 4.3: Graph of lignin composition (left axis) and total lignin area (right axis) across experimental groups ...... 39
Figure 4.4: Incorporation of 13C label into G (blue) and S (red) subunits in Wt and comt mutant Arabidopsis seedlings supplied 13C-labeled coniferyl alcohol ...... 40
vii CHAPTER 1
INTRODUCTION
Lignin is the second most abundant biopolymer after cellulose and comprises a significant percentage of the cell wall in many plant tissues. It is found mainly in the stems of woody plants where it serves to strengthen the stem and facilitate water transport by acting as a hydrophobic barrier in xylem elements. Other tissues such as roots, flowers and leaves also contain lignin in vascular tissues, and seed coats also contain lignin in varying amounts.
Lignin is a complex phenolic polymer composed of cross-linked cinnamyl alcohols, called monolignols. The primary monolignols are p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol that are polymerized, respectively, to give the so-called hydroxyphenyl (H), guaiacyl (G), and syringyl (S) units of lignin shown in Figure 1.1.
The caffeyl alcohol that polymerizes to form the recently discovered caffeyl lignin (C- lignin) is also shown.
Figure 1.1. The three primary monolignols that polymerize to make lignin and the caffeyl alcohol monomer from which C-lignin is derived.
The amount of lignin available in the biosphere and its aromatic composition make it an attractive starting material for commodity chemicals and other materials.
1 Many uses of lignin have been explored- through depolymerization to yield chemical feedstocks (Xu et al. 2014), as a bulk filler in plastics and rubber (Dias et al. 2016; Jiang et al. 2014), and as a precursor for carbon fibers (Thunga et al. 2014), to name a few.
However, the chaotic nature of the lignin polymer presents several challenges to its utilization as a chemical feedstock or as a bulk material. Linkages between monomers are varied and the polymer is branched, presenting problems for depolymerization to chemical feedstocks. It is a heteropolymer composed of several different subunits, which tends to yield a variety of difficult to separate products after depolymerization.
Also, once extracted, lignin is a mixture of lignin fragments, usually with a wide polydispersity index, which may be chemically modified depending on the extraction method (Ragauskas et al. 2014). The non-homogenous size and composition of extracted lignin fragments pose a challenge to material manufacturing processes, such as carbon fiber manufacture, which rely on a homogeneous polymer precursor.
Research into lignin modification is predominantly focused on reduction of its biosynthesis or facilitation of its degradation- mainly for improved saccharification of biomass, with little research done to produce a more useful polymer. Recently, a novel type of lignin, called C-lignin, comprised only of caffeyl alcohol subunits, was discovered. This lignin appears in seed coats, but not stems or leaves, of several plants:
Vanilla planifolia (Vanilla orchid), Cleome hassleriana (Spider flower), Jatropha curcas
(Barbados nut), and other mainly exotic plant species (Chen et al. 2012). C-lignin is an ideal lignin for utilization as a chemical feedstock; it is a linear homopolymer that can be depolymerized to catechol derivatives with minor byproducts (Li et al. 2018). The linear structure of C-lignin also makes the polymer more suitable for processing into carbon
2 fibers than the branched structure of native lignin (Nar et al. 2016).
C-lignin accumulates in the seed coats of Cleome starting approximately 13 days after pollination (DAP); before this the seed coats make normal G-lignin (Tobimatsu et al. 2013). The stem of Cleome contains G/S lignin and no C-lignin. The genomes of
Cleome and several other members of the family Cleomacea have been published, and
Cleome and its relatives have been well researched due to the recent evolution of C4- type photosynthesis within the family (Bhide et al. 2014). The existing genomic resources make Cleome an ideal model system in which to study the mechanisms leading to C-lignin accumulation.
In this thesis, Cleome is used as a model organism to discover genes associated with C-lignin biosynthesis; subsequently, the recombinant protein products of these genes are in vitro characterized; and the ability of caffeyl alcohol to polymerize in
Arabidopsis is investigated. To understand the significant differences between C-lignin and native “classical” G/S lignin, a review of the processes of monolignol biosynthesis and lignin polymerization is first provided.
Biosynthesis of Monolignols
Monolignols are first synthesized in the cytosol from L-phenylalanine through the phenylpropanoid and monolignol pathways, then exported to the cell wall, and finally oxidized by peroxidases/laccases into monolignol radicals which then undergo polymerization (Takabe et al. 2001; Vanholme et al. 2010). The overall lignin biosynthesis pathway is shown in Figure 1.2.
3 Figure 1.2: The lignin biosynthesis pathway, from Tobimatsu et al. 2013. The enzymes are L-phenylalanine ammonia-lyase (PAL), cinnamate-4-hydroxylase (C4H), 4-coumarate: Coenzyme A Ligase (4CL), hydroxycinnamoyl CoA shikimate/quinate hydroxycinnamoyltransferase (HCT), p-coumaroyl shikimate 3′-hydroxylase (C3′H), caffeoyl-CoA 3-O-methyltransferase (CCoAOMT), cinnamoyl-CoA reductase (CCR), caffeic acid/5-hydroxyconiferaldehyde 3/5-O-methyltransferase (COMT), ferulate/coniferaldehyde 5-hydroxylase (F5H), cinnamyl alcohol dehydrogenase (CAD), peroxidase (POX), laccase (LAC). H, G, and S monomers are classical lignin monomers- C and 5H monomers are atypical.
4 Typical lignin is comprised of H, G, and S monomers originating from their respective cinnamyl alcohols (monolignols). The following description describes those biosynthetic steps of particular relevance for the present work. These monolignols are produced by reduction of cinnamyl aldehydes by cinnamyl alcohol dehydrogenase
(CAD). There are several CAD’s known to be functional in lignification in Arabidopsis and other plants. These CAD’s have been shown to reduce many cinnamyl aldehydes, but most CAD’s favor a certain substrate, for instance AtCAD4 and 5 from Arabidopsis prefer p-coumaraldehyde (Kim et al. 2004), whereas another CAD in Populus tremuloides (aspen) (PtSAD) prefers sinapaldehyde (Li et al. 2001). Production of caffeyl alcohol, the monolignol which is polymerized to C-lignin, also involves CAD.
AtCAD2, while not considered a classical CAD due to its low catalytic efficacy, does show preference for caffealdehyde (Kim et al. 2004). However, there has been very little in vitro research using caffealdehyde as a substrate for CAD, as it is not considered a classical monolignol.
The cinnamyl aldehyde precursors of H and G monolignols (p-coumaraldehyde and coniferaldehyde, respectively) are produced by reduction of their respective cinnamoyl-CoAs by cinnamoyl CoA reductase (CCR). Like CADs, CCRs have been shown to be multifunctional but favor one substrate over another. For instance, MtCCR1 from the model legume Medicago truncatula prefers feruloyl-CoA whereas MtCCR2 prefers caffeoyl-CoA (Zhou et al. 2010). Sinapaldehyde is produced by hydroxylation and O-methylation of the aromatic ring of coniferaldehyde, the first of these reactions being catalyzed by the cytochrome P450 enzyme ferulic acid/coniferaldehyde 5- hydroxylase (F5H).
5 Figure 1.3: Lignin deposition and O-methyltransferase activity in the seed coat of C. hasselerania (from Tobimatsu et al. 2013). A) Composition of seed coat lignin at six to 20 days after pollination (DAP), guaicyl (G) and catechyl (C). B) O-methyltransferase activity of seed coats during the same time period. C) Seed sections imaged by autofluorescence, or stained with toluidine blue, phloroglucinol, or nitroso reagent. The yellow and orange arrows indicate two sublayers. The layer indicated in orange deposits lignin at 14 DAP and after, while the layer indicated in yellow is lignified before then.
The precursors to G and S monolignols require O-methylation of the aromatic ring. O-Methylation is possible at different steps within the pathway shown in Figure 2
(i.e. at the CoA, aldehyde, or alcohol levels) but, according to the currently accepted pathway, it occurs preferentially at the level of the CoA and aldehyde catalyzed by caffeoyl-CoA 3-O-methyltransferase (CCoAOMT) and caffeic acid/5- hydroxyconiferaldehyde 3-O-methyltransferase (COMT), respectively (Zhou et al. 2010;
Guo et al. 2001; Ma and Xu 2008). The precursors to caffeyl alcohol are not O- methylated, therefore C-lignin accumulation could be expected to be associated a
6 pathway that does not feature CCoAOMT or COMT activity. Consistent with this idea, loss of OMT activity was shown to coincide with the appearance of C-lignin in Cleome seed coats, as shown in Figure 1.3 (from Tobimatsu et al. 2013). Similarly, RNA-seq results from V. planifolia seeds show an absence of both OMT enzymes in developing vanilla seed coats known to accumulate C-lignin (Rao et al. 2014).
Polymerization of Lignin
The final step of lignin deposition is polymerization of cinnamyl alcohols by the action of peroxidases (POX) and/or laccases (LAC) (Zhao et al. 2013). The most widely accepted mechanism of lignin polymerization is by oxidative coupling where monolignol radicals, or sometimes their conjugates, are added to the ends of a growing lignin polymer. This mechanism produces random, branched lignin polymers whose subunit compositions are based on the relative concentrations of monolignols in the cell wall, whereas their interunit linkages vary depending on each subunit’s propensity to cross- link at different positions. Thus the final structures of lignin macromolecules are incredibly varied and challenging to characterize (Ralph et al. 2004).
The ability of LAC to catalyze polymerization of caffeyl alcohol has not been reported, but a C-lignin polymer can be made in vitro from caffeyl alcohol in the presence of horseradish peroxidase and hydrogen peroxide (Chen et al., 2012).
Arabidopsis POX/LAC have been shown to possess the ability to polymerize 5- hydroxyconiferyl alcohol (5-H), various cinnamaldehydes, and monolignol p- coumarate/ferulate conjugates (Smith et al. 2015; Karlen et al. 2016). Maize cell walls, when supplied with exogenous substrates, have shown remarkable plasticity to polymerize a variety of phenolic chemicals structurally different from monolignols
7 (Grabber et al. 2010). During polymerization of C-lignin, the reactivity of caffeyl alcohol
leads to formation of benzodioxane rings through an internal cyclization reaction during
oxidative coupling, causing the C-lignin polymer to have a linear and therefore less
chaotic structure than the normal G/S lignin (Tobimatsu et al. 2013). Also, due to the
structure of the caffeyl monomer and its radical intermediates, it is thought that C-lignin
does not bond to neighboring cell wall carbohydrates during polymerization like normal
G/S lignin, thus removing a major potential for causing biomass recalcitrance (Chen and
Dixon 2007).
Engineering Lignin Subunits
The plasticity of monolignol polymerization suggests that many aromatic
compounds could be incorporated into lignin if they were present in the cell wall. This is
attractive from an engineering point of view because it opens the possibility of designing
a lignin which is either easier to remove or has added value as a bioproduct. To this
end, chemically labile linkages have been engineered into poplar lignin by expressing
an exogenous monolignol ferulate transferase which conjugates G or S monolignols to
ferulate (as feruloyl-CoA), introducing an ester bond into lignin which is able to be easily
cleaved during pretreatment of lignocellulosic biomass (Wilkerson et al. 2014). In
another approach, overexpression of F5H in poplar resulted in increased production of
sinapaldehyde and significantly increased the proportion of S-lignin, making the lignin more amenable to degradation (Stewart et al. 2009).
The presence of C-lignin in the seed coats of specific plants is notable, but for engineering and utilization as a bulk material, the polymer would need to be present in stems or other high biomass tissue. Outside of the seed coat, low levels of C-lignin (as
8 a co-polymer with G lignin) were found in modified pine tracheary element cell cultures with suppressed CCoAOMT activity (Wagner et al. 2011). However, no C-lignin has been found in any other COMT or CCoAOMT mutants to the author's knowledge, and
Arabidopsis double OMT mutants are severely dwarfed and do not contain detectable
C-lignin (Do et al. 2007). No monolignol feeding studies have been published showing polymerization of caffeyl alcohol by POX/LAC in vivo.
Hypothesis and Objectives
The hypotheses of this thesis are 1) Caffeyl alcohol accumulation and C-lignin deposition in Cleome seed coats require downregulation of both COMT and CCoAOMT; specific CCR, CAD, and LAC enzymes with preference for non-methylated monolignol precursors may also be involved. 2) Arabidopsis may have the ability to polymerize caffeyl alcohol endogenously if supplied the monolignol; if not, the need for a polymerization system with specificity for caffeyl alcohol is indicated.
The objectives of this thesis are 1) To determine if C-lignin specific CCR/CAD enzymes are involved in C-lignin formation in the Cleome seed coat. 2) To determine if endogenous Arabidopsis cell wall polymerization enzymes can polymerize supplied caffeyl alcohol.
These hypotheses and objectives were explored by 1) Measuring the relative activity of recombinant Cleome CCR and CAD proteins, which are differentially expressed coinciding with C-lignin formation, with potential substrates. 2) Monitoring incorporation of supplied caffeyl alcohol and 13C-coniferyl alcohol into lignin polymers in
Arabidopsis tracheary elements grown in a xylem induction system. To understand the rationale behind the methods used, a brief overview of the applicable methods follows.
9 Methods Overview
1.5.1 Candidate Gene Discovery by RNA Sequencing
Gene discovery refers to the process of finding a gene, or several genes, responsible for a trait of interest. One of the most direct ways is by measuring the relative transcript levels in different tissues, and/or at different time points, with and without the trait, by RNA sequencing (RNAseq). The transcript levels in these tissues, or at these timepoints, are then interrogated to find differentially expressed genes. If a trait is related to a known biosynthetic pathway, such as monolignol biosynthesis, then differentially expressed genes can be narrowed down to genes which are homologous to the enzymes which would logically affect the trait, such as CCR or CAD. Alignment and modeling of the proteins encoded by the candidate genes can be used to locate potential differences in amino acid composition which may change the enzymatic activity of the protein. An example of gene discovery related to lignin biosynthesis in the non-model organism Cunninghamia lanceolata (Chinese fir) can be found in Huang et al. (2012). It should be noted that this approach finds candidate genes; further genetic analysis would be necessary to unequivocally ascribe them a function in a particular trait.
1.5.2 Recombinant Expression and Characterization of Target Proteins
Once candidate genes in a biochemical pathway have been found, the enzymes they encode can be recombinantly expressed so that their in vitro activity can be determined. If the candidate genes do not have any glycan substitutions and are not membrane bound, they may be able to be recombinantly expressed in E. coli, otherwise a eukaryotic expression system is needed. Cloning of the candidate genes to a suitable
10 expression vector is usually done in two steps. First, the open reading frame of the gene is amplified from the cDNA of tissues of the organism of interest that express the gene, and inserted into an entry vector, which is transformed into an E. coli strain where the
vector is multiplied. Then, after sequence verification, the gene of interest is inserted
into a suitable destination vector, by restriction digestion, ligation or other methods, for
expression. Destination vectors for recombinant protein expression usually have a
purification tag to facilitate purification of the target protein after expression and an
inducible promoter which ensures the target protein is only expressed once the
appropriate inducer compound has been added to the culture medium.
Once the gene of interest is cloned to the final destination vector, the vector is
transformed into a suitable expression strain. The transformed expression strain is
grown to a certain culture density and expression of the target protein is induced by
adding the appropriate inducer compound. Expression of the target protein is tested at
different temperatures for different time periods, depending on the protein being
expressed and the expression system. Generally, the lower the expression temperature,
the longer the expression time needed, but, perhaps paradoxically, some proteins
remain more soluble in E. coli when expressed at lower temperatures. Once the protein
is expressed, it is purified from non-target host proteins using the purification tag and
analyzed by sodium dodecyl sulfate –polyacrylamide gel electrophoresis (SDS-PAGE)
to estimate purity, and the amount of protein quantified by Bradford assay. The purified
recombinant proteins can be used for in vitro characterization of enzymatic activity. An
example of cloning, recombinant expression, and in vitro characterization of a lignin
related gene (AtCCR1) can be found in Baltas et al. (2005).
11 For in vitro characterization of kinetic properties, a known quantity of the recombinant protein is incubated with varying concentrations of a potential substrate in
an appropriate buffer system, with the required cofactors. The reaction is conducted at a
predetermined temperature and time, then stopped by the addition of a suitable acid (or
other compound) and the substrates or products quantified. By varying the amount of
starting substrate and quantifying the amount of product produced by the enzyme, a
graph of reaction rate vs. substrate concentration can be constructed; this can show
whether the enzyme follows so-called Michaelis–Menten kinetics. By plotting the
reciprocal of the initial rate versus the reciprocal of the substrate concentration, a
Lineweaver–Burk plot can be generated from which the Km and Vmax are estimated for
each enzyme/substrate combination. Km and Vmax are standardized measurements of
an enzyme’s affinity and maximum reaction rate, respectively, for a particular substrate.
These values can be used to compare an enzyme’s relative activity with different
substrates, and to compare the activity of different enzymes to estimate their effect on
metabolic flux (Wang et al. 2014).
1.5.3 In vitro Polymerization Systems and Induction of Lignification in vivo
In vitro methods which investigate true lignin polymerization are challenging to
relate to in vivo activity because the peroxidases and lacasses which polymerize lignin
are functional in the plant cell wall, thus their activity may be affected by interactions
with cell wall structures. Outside of the cell wall, in vitro polymerization systems may not
provide entirely accurate information about polymerization mechanisms or in planta
functionality. However, a common method to investigate polymerization of monolignols
is to initiate free-radical polymerization of cinnamyl alcohols by horseradish peroxidase
12 and hydrogen peroxide. This reaction produces pseudo-lignin polymers termed dehydrogenation polymers, measurement of which is useful for testing the polymerization capabilities of potential monolignols (Méchin et al. 2007).
Another polymerization system utilizes non-living maize cell walls which are washed of monolignols and other metabolites, but retain cell wall bound peroxidases and lacasses, to polymerize potential monolignols. Compared to generation of dehydrogenation polymers, this system is a more accurate representation of lignin polymerization by POX/LAC as it involves enzymes embedded in the cell wall (Grabber et al. 1996). Using this system, several novel compounds, unrelated to known monolignols, were found to polymerize into lignin when supplied to maize cell walls
(Grabber et al. 2010).
In vivo methods for investigating polymerization of monolignols involve the
induction of ectopic lignification. The two basic approaches are cell culture of tracheary
elements (TE) and xylem induction in intact seedlings. Arabidopsis protoplast cultures
can be induced to form TEs by over-expression of the VND7 transcription factor that
controls lignification genes, resulting in upregulation of POX/LAC and other enzymes
involved in lignification. These induced TE cultures deposit lignin similar to that found in
the xylem of intact stems (Schuetz et al. 2014). Another lignin induction system uses
bikinin to activate brassinosteroid signaling in leaf disks or intact seedlings of
Arabidopsis, resulting in the formation of ectopic TEs and accumulation of lignin (Kondo et al. 2016). This xylem induction system can be used in intact seedlings grown in liquid media, which facilitates supplying the plants with monolignols, and is amenable for comparison of mutant lines.
13 1.5.4 Analysis of Lignin Presence, Amount and Composition
In planta lignin phenotypes can be explored by light microscopy of tissues
stained with many lignin-specific or partially specific stains: toluidine blue (TBO) stains
lignin and other cell wall polymers, Mäule staining reveals S-lignin, phloroglucinol stains
lignin well and reveals cinnamaldehyde end groups in the polymer (Mitra et al. 2014).
Also, the autofluorescence of lignin can be imaged by fluorescence microscopy. Of the
staining methods, TBO is the easiest to work with and is able to stain a variety of cell
wall polymers. TBO stains lignin blue-green, pectin and hemicellulose pink, and DNA
purple (Mitra et al. 2014). Staining tissues with TBO is an effective way to differentiate
lignified cells, such as TEs, from non-lignified cells, such as mesophyll cells.
Chemical analysis of lignin, such as quantification of lignin polymer and
determination of lignin composition, requires an analytical technique like gas
chromatography/mass spectrometry (GC-MS). For GC-MS analysis, the lignin polymer cannot be directly analyzed due to its polymeric nature, and it must be depolymerized first. There are several methods, but one of the most used is a process called thioacidolysis. During thioacidolysis the lignin polymers are depolymerized to generate thioethylated monomers which can be quantified by GC-MS (Harman-Ware et al. 2016).
14 CHAPTER 2
DISCOVERY OF CANDIDATE GENES RELATED TO C-LIGNIN BIOSYNTHESIS IN
Cleome SEED COATS
Objectives
• Discover genetic changes involved in C-lignin biosynthesis and deposition in
the seed coat starting at 13 DAP by investigating transcript levels before and after the
switch to C-lignin deposition in seed coats, and in tissues containing normal lignin.
• Determine if the differentially expressed genes/proteins have novel domains or are similar to other characterized genes/proteins by comparing the Cleome
sequences to Arabidopsis homologs.
Methods
2.2.1 RNA Sequencing and Assembly
RNAs from Cleome stem, bark, fiber, pith, and seed coat (at 8, 10, 12, 14, 16,
and 18 DAP) were sequenced at DOE JGI (Joint Genome Institute, Walnut Creek, CA),
using paired-end sequencing on an Illumina HiSeq 2500. The sequences were then
assembled by Dr Xiaolan Rao using Bowtie software with the available Cleome genome
(ASM46358v1) as a reference (Zhuo et al. 2019).
2.2.2 Analysis of Differential Gene Expression
The fragments per kilobase of transcript per million reads (FPKM) values of
expressed transcripts were calculated by Dr. Xiolan Rao using eXpress 1.5.0 software
(Zhuo et al. 2019). Then the levels of lignin-related transcripts were compared across
stem and seed coat tissues using spreadsheet software.
15 2.2.3 Homology Mapping to Arabidopsis
Cleome transcripts and their proteins were compared to ones in Arabidopsis
using BLASTN or BLASTP (ncbi.nlm.nih.gov) and homologs aligned using CLUSTAL
O(1.2.4) (www.ebi.ac.uk). The Zn-1, Zn-2, and NADP(H) binding domains and
substrate binding sites were annotated in the CAD alignment by comparison to known
CAD domains (Trabucco et al 2013).
2.2.4 Protein Modeling
The Cleome protein ChCAD5 was modeled and docked by Dr. Xiaoqiang Wang
using MODELLER and AUTODOCK programs, with its Arabidopsis homolog (AtCAD5)
as a template (Zhuo et al. 2019).
Results and Discussion
2.3.1 RNA Sequencing
RNA sequencing, by Dr. Xiolan Rao, of Cleome tissues resulted in 45,198 full-
length transcripts. The identity of transcripts agreed well with the reference genome of
Cleome, with 93.3% of transcripts mapping to the reference genome (Zhuo et al. 2019).
These results indicate that the RNA sequencing adequately captured the range of
Cleome transcripts, and that the resultant transcriptomes are valid for comparing across
tissue samples and seed coat time points.
2.3.2 Differential Gene Expression
Comparing transcript expression (FPKM) across seed coat developmental time
points and non-seed tissues (stem, bark, fiber, and pith) revealed several possible
candidates for seed coat specific enzymes which may be responsible for C-lignin
16 deposition starting between 12 to 14 DAP. The criteria were: 1) minimal expression in non-seed tissues, 2) expression peaking around or after 13 DAP, and 3) transcript homology to enzymes involved in phenylpropanoid biosynthesis.
Levels of both COMT and CCoAOMT transcripts fell during the period of the switch to C-lignin formation, in agreement with previous results showing loss of extractable OMT enzymatic activity in Cleome seed coats (Tobimatsu et al. 2013). This
loss of OMT activity is hypothesized to be needed for accumulation of caffeyl alcohol, as
substrate for the polymerization to C-lignin by POX/LAC; however loss of OMT activity
alone is likely not sufficient to cause deposition of C-lignin. Two CCR and CAD
transcripts showed expression patterns suggesting their involvement in the appearance
of C-lignin (Figures 2.1 and 2.2, respectively). Together these enzymes act to reduce
phenylpropanoids to their respective monolignols and it was hypothesized their activity
could be involved in caffeyl alcohol accumulation.
The transcript levels of CCR (ChCCR1) in Cleome seed coats reached maximum
at 14 DAP and were low (relative to peak expression at 14 DAP) in non-seed tissues.
This expression pattern, and the known activity of CCR in the phenylpropanoid
pathway, made CCR1 a good candidate for a role in C-lignin biosynthesis- leading to
accumulation of caffealdehyde, with subsequent conversion to caffeyl alcohol. Similarly,
CAD5 (ChCAD5) transcript levels reach a maximum at 14 DAP and the transcripts are
absent in non-seed tissues, making CAD5 a candidate for the enzyme catalyzing the conversion of caffealdehyde to caffeyl alcohol, specific to seed coat tissues. Another
CAD (ChCAD4) was found to be similarly expressed across all seed coat time points and highly expressed in lignifying stem and fiber. This CAD4 was hypothesized to be
17 enzyme involved in production of classical G/S lignin in stem and early stages of seed coat maturation.
Figure 2.1: Transcript levels of ChCCR1 in Cleome. The tissues shown are seed coat at 8- 18 days after pollination (DAP), stem, bark, fiber, and pith.
Figure 2.2: Transcript levels of ChCAD4 (green) and ChCAD5 (red) in Cleome. Tissues shown are seed coat at 8-18 days after pollination (DAP), stem, bark, fiber, and pith.
18 2.3.3 Comparison of Candidate Cleome Transcripts to Arabidopsis Homologs
The amino acid sequences of each candidate transcript were aligned to their
nearest Arabidopsis homologs. ChCCR1 shared 96 % similarity to its Arabidopsis
homolog (AtCCR1). ChCAD4 and ChCAD5 were aligned to AtCAD5 (a CAD shown to
be predominantly involved in lignification of Arabidopsis stems (Kim et al. 2004)) and
AtCAD2 (the only CAD shown to have a substrate preference for caffealdehyde (Kim et
al. 2004)). Conserved domains and substrate binding sites were annotated by
comparison to known CAD domains (Trabucco et al 2013). CAD has two zinc-binding
domains (Zn-1 and Zn-2). Both Cleome CADs aligned well to Arabidopsis CADs and
showed high sequence identity to AtCAD5 across conserved Zn-1, Zn-2, and NADP(H)
binding domains and most substrate binding residues (Figure 2.3). However, ChCAD5
has a histidine at position 58, whereas AtCAD5 and ChCAD4 both have a leucine at this
position. The caffealdehyde-preferring AtCAD2 also shows a substitution at this
position. This change in amino acids is significant because it is within a known
substrate-binding site for CADs. This substitution was hypothesized to affect substrate
specificity.
2.3.4 Protein Modeling of ChCAD5
The amino acid sequence of ChCAD5 was modeled by Dr Xiaoqiang Wang using
the available structure of AtCAD5 as a template. In silico docking studies with potential
substrates indicated interaction of histidine 58 with substrates (Figure 2.4) (Zhuo et al.
2019). Subsequent site-directed mutagenesis studies by Dr. Chunliu Zhuo showed that a histidine at position 58 favored binding of caffealdehyde over coniferaldehyde (Zhuo et al. 2019). The in vitro substrate preference of ChCAD5 for caffealdehyde was
19 demonstrated through in vitro assays of recombinant ChCAD5, as detailed in the next section, and supported by additional studies by Dr. Chunliu Zhuo (Zhuo et al. 2019).
Figure 2.3: Protein alignment of ChCAD4 and 5 to AtCAD2 and 5. The Zn-1, Zn-2, and NADP(H) binding domains (Trabucco et al 2013) are indicated by black boxes. The substrate binding residues (Trabucco et al 2013) are indicated above the residues by black asterisks and position 58 is marked with a black arrow. Sequence agreement is indicated by grey symbols below the residues- asterisk for fully conserved residues, colon for non- conserved residues with strongly similar properties (greater than 0.5 on point accepted mutation (PAM) 250 matrix), period for non-conserved residues with weakly similar properties (less than or equal to 0.5 on PAM 250 matrix, and a blank space for non- conserved residues with dissimilar properties.
20 Figure 2.4: Model of active site, by Dr. Xiaoqiang Wang, of ChCAD5 (blue) showing the specific amino acids (orange) interacting with caffealdehyde (yellow) (Zhuo et al. 2019).
Conclusion
Three candidate enzymes (ChCCR, ChCAD4, and ChCAD5) in the monolignol
biosynthesis pathway, potentially related to C-lignin formation, were found by
exploration of differentially expressed genes at specific times during the maturation of
the Cleome seed coat and by comparison to other non-C-lignin containing tissues. An
amino acid substitution (L58 to H58) in the active site of ChCAD5 was found by
comparing the protein sequences of ChCADs to AtCADs. Through protein modeling
studies, this substitution was shown to impact binding of substrates at the active site of
ChCAD5, and affect the enzyme’s relative activity with the C-lignin precursor caffeyl alcohol.
21 CHAPTER 3
IN VITRO CHARACTERIZATION OF RECOMBINANT CANDIDATE ENZYMES
Objectives
• Recombinantly express the candidate genes (ChCCR1, ChCAD4, and
ChCAD5) implicated in C-lignin biosynthesis in the Cleome seed coat.
• Determine the substrate specificity and kinetic properties of the corresponding enzymes through in vitro kinetic assays with potential substrates and analysis of
Lineweaver–Burk plots.
• Investigate the cooperation of ChCCR1 and ChCAD4/5 to produce caffeyl alcohol in the presence of other substrates by in vitro assays with single or mixed enzymes, incubated with single or mixed substrates.
Methods
3.2.1 Cloning, Recombinant Protein Expression, and Enzyme Purification
The open reading frames of each transcript (ChCCR1, ChCAD4, and ChCAD5) were amplified from Cleome seed coat cDNA using the primers listed in Table 1. The
PCR products were purified with QIAquick PCR purification columns (Qiagen) and inserted separately into dTOPO entry vectors, sequence verified, and cloned to pMALc5 expression vectors. Expression vectors were transformed into chemically competent E. coli (Rosetta) cells and expression induced with 300 μM isopropyl β-D-1- thiogalactopyranoside (IPTG) at 20 C for 16 hours. Enzymes were purified using amylose resin (NEB) according to vendor protocols; purification was carried out on ice or in a cold room. Protein content of enzyme preparations was quantified by Bradford assay and purity estimated by densitometry on SDS-PAGE gels
22 (https://openwetware.org/wiki/Protein_Quantification_Using_ImageJ ). Enzyme
solutions were aliquoted to sterile PCR tubes and stored at -80 C until use; the elution
buffer contained 10% glycerol and 10 mM β-mercaptoethanol to aid enzyme stability.
Table 3.1: Primers used for cloning of candidate genes.
Transcript Forward Primer Reverse Primer
ChCCR1 CACCATGCCCGTCGACGCACCT CACTCCAGGGAAAGTGAGGA
ChCAD4 CACCATGGGGAGCATTGA GGCTGCGTATTGTGTCTCCT
ChCAD5 CACCATGGGAAGGCATGAAGGAGA ACCCGTAGGTTTGTGTCGAG
3.2.2 Kinetic Assays
Coniferaldehyde and coniferyl alcohol were obtained from Sigma-Aldrich, St
Louis, MO, USA. Caffeoyl CoA, feruloyl CoA and caffealdehyde were purchased from
MicroCombichem, Wiesbaden, Germany. Caffeyl alcohol was received from Dr Rui
Katahira, National Renewable Energy Laboratory, Golden, CO, USA.
Purified enzymes were assayed in vitro in final reaction volumes of 250 μL
containing 50 mM sodium phosphate buffer (pH 7.5), 10 mM β-mercaptoethanol, 400
μM NADPH, 50 ng protein, and 0.5 to 200 μM substrates. Reactions were warmed to 30
C in 96 well UV transparent plates in a multiplate reader (Synergy MX) and started by
adding substrate solution utilizing a multi-channel pipette. The linear reaction time (4 min) was first determined by measuring the absorbance of NADPH at 340 nm. After 4 min at 30 C, the reactions were stopped with 10 μL of glacial acetic acid and put on ice.
Fifty μL of each reaction solution was mixed with an equal volume of 1:1 (v/v)
MeOH:H2O, filtered, and 90 μL injected onto an HPLC column for quantification of
reaction products as outlined below. The rate of product formation was plotted against
23 substrate concentration and the Lineweaver Burk equation was used to estimate the Km and Vmax from which other kinetic parameters were calculated.
3.2.3 Mixed Reactions
Reactions were conducted with single and multiple enzymes which were supplied single and/or multiple substrates as follows: 1) ChCCR1 with caffeoyl-CoA, 2) ChCCR1 with caffeoyl-CoA and feruloyl-CoA, 3) ChCAD4 with caffealdehyde, 4) ChCAD4 with caffealdehyde and coniferaldehyde, 5) ChCCR1 and ChCAD4 with caffeoyl-CoA, 6)
ChCCR1 and ChCAD4 with caffeoyl-CoA and feruloyl-CoA, 7) ChCAD5 with caffealdehyde, 8) ChCAD5 with caffealdehyde and coniferaldehyde, 9) ChCCR1 and
ChCAD5 with caffeoyl-CoA, 10) ChCCR1 and ChCAD5 with caffeoyl-CoA and feruloyl-
CoA.
The composition of mixed reactions was identical to that used in the kinetic assays, except substrate concentrations were held constant (30 μM caffeoyl-CoA and caffealdehyde, 10 μM feruloyl-CoA and coniferaldehyde) and a 10:1 CCR:CAD enzyme ratio was used by increasing the concentration of ChCCR1 10 fold to 500 ng per 250 μL reaction. Mixed reactions were incubated, stopped, and prepared for HPLC similar to kinetic assays, except the reaction time was extended to 10 min.
3.2.4 HPLC Analysis
HPLC-UV quantification of reaction products was carried out on a Thermo
Scientific Dionex UltiMate 300 system using a Luna C18(2) reverse-phase column (5
μm particle size, 250 x 4.6 mm, Phenomenex). The following solvent gradient was used:
5% B for 5 min, to 15% B in 5 min, to 23% B in 15 min, to 33% B in 5 min, to 40% B in 5 min, to 100% B in 5 min, 100% B for 5 min, to 5% B in 5 min (mobile phase A: 1%
24 phosphoric acid in water, mobile phase B: acetonitrile). Total run time was 50 min, flow rate was 1 mL/min, and the column was heated to 30 C. Reaction products were identified by comparing UV spectra and retention times to known standards and were quantified against standard curves.
Results and Discussion
3.3.1 Expression of Recombinant Proteins
After purification and elution from amylose beads, recombinant proteins
appeared as distinct bands of the correct size on SDS-PAGE. Impurities were estimated
to be approximately 80% by summing intensities of all other bands in the lane; this
value was used to factor protein content in calculations for subsequent in vitro reactions.
Bradford assay showed the two CAD proteins were eluted at approximately equal
concentrations and at a higher concentration than the CCR protein.
3.3.2 Kinetic Properties of ChCCR and ChCAD4 and 5
The in vitro activity of ChCCR with feruloyl-CoA and caffeoyl-CoA is shown in
Figure 3.1. These results show ChCCR strongly prefers feruloyl-CoA to caffeoyl-CoA, similar to other characterized CCRs (Baltas et al. 2005; Ma 2010; Pichon et al. 1998).
The lower activity with caffeoyl-CoA suggests any significant in vivo channel from caffeoyl-CoA to caffealdehyde would require high expression of ChCCR and low levels of feruloyl-CoA in the available metabolite pool. The conditions for this channel are present during the later stages (post 13 DAP) of Cleome seed coat maturation: the expression of ChCCR peaks significantly at 14 DAP in the seed coat, to over four-fold higher transcript level than in stem or fiber tissues; and catalytic activity of COMT and
25 CCoAOMT is lost around the same time (Tobimatsu et al. 2013), presumably depleting the supply of feruloyl-CoA.
Figure 3.1: In vitro activity of ChCCR1 with feruloyl-CoA (red) and caffeoyl-CoA (blue). Results are from duplicate reactions performed under identical conditions, error bars represent standard deviation.
The in vitro activity of ChCAD4 with coniferaldehyde and caffealdehyde is shown
in Figure 3.2. These results show that the properties of ChCAD4 are similar to those of
other characterized CADs in that the enzyme shows a preference for coniferaldehyde
over caffealdehyde. There appeared to be substrate inhibition at higher concentrations
of coniferaldehyde, whereas this inhibition was not seen for caffealdehyde. The
caffealdehyde reducing activity of ChCAD4, while lower than for coniferaldehyde, shows
that the enzyme is nevertheless able to produce caffeyl alcohol. However, the transcript
26 expression pattern and in vitro activity together suggest that ChCAD4 is mainly involved in supplying substrate for G-lignin biosynthesis in the seed coat and probably G/S-lignin
biosynthesis in the stem and fiber.
Figure 3.2: In vitro activity of ChCAD4 with coniferaldehyde (red) and caffealdehyde (blue). Results are from duplicate reactions performed under identical conditions, error bars represent standard deviation.
The in vitro activity of ChCAD5 with coniferaldehyde and caffealdehyde is shown
in Figure 3.3. These results are significant in showing a rare preference for
caffealdehyde over coniferaldehyde not seen to date in CADs that have been ascribed a
role in lignification (Kim et al. 2004). The substrate preference of ChCAD5, and its high expression during Cleome seed coat maturation, strongly suggest it is responsible for an in vivo channel from caffealdehyde to caffeyl alcohol in the Cleome seed coat,
27 providing the monomer for polymerization to C-lignin. The kinetic parameters for all recombinant enzymes are shown in Table 3.2. Combined, the kinetic results suggest that a pathway from caffeoyl-CoA to caffeyl alcohol involving ChCCR and ChCAD4 or 5 is possible, and this was investigated further in mixed reactions.
Figure 3.3: In vitro activity of ChCAD5 with coniferaldehyde (red) and caffealdehyde (blue). Results are from duplicate reactions performed under identical conditions, error bars represent standard deviation.
Table 3.2: Kinetic parameters for recombinant enzymes with caffeoyl-CoA and feruloyl- CoA or caffealdehyde and coniferaldehyde.
Enzyme Substrate Vmax (nkat/mg) kM (μM) Vmax/kM Caffeoyl-CoA 9 2.5 3.6 ChCCR1 Feruloyl-CoA 75 2.2 34.1 Caffealdehyde 455 11.6 39.2 ChCAD4 Coniferaldehyde 909 17.8 51.1 Caffealdehyde 971 38.3 25.4 ChCAD5 Coniferaldehyde 426 24.8 17.2
28 3.3.3 Mixed Reactions with ChCCR and ChCAD 4 or 5
To investigate the ability of ChCCR and ChCAD4/5 to produce caffeyl alcohol in the presence of other substrates, mixtures of recombinant enzymes were incubated with potential substrates. The results from the mixed reactions are shown in Figure 3.4.
These reactions can be directly compared as they were all performed using the same
buffer conditions, in the same plate, for the same amount of time and their products
quantified in the same HPLC run. The mixed enzyme reactions were made to reflect a
high abundance of CCR relative to CAD (10:1 ratio), and the reactions containing two
substrates to contain a high abundance of caffeoyl-CoA or caffealdehyde relative to
feruloyl-CoA or coniferaldehyde (30 μM and 10 μM, respectively). In the following
paragraphs, the reactions will be referred to as reactions 1 through 10 as numbered left
to right in Figure 3.4.
Reaction 1 shows that ChCCR1 is able produce about 1.2 nmol of caffealdehyde
under these conditions; this is reflected as the rate-limiting step in reactions 5 and 9.
Comparing reaction 2 to 1 shows the preference of ChCCR1 to produce
coniferaldehyde when given a mixture of feruloyl-CoA and (higher amounts of) caffeoyl-
CoA; this relative conversion rate is reflected in reactions 6 and 10. Taken together,
these reactions show that ChCCR1 strongly prefers feruloyl-CoA as a substrate even
when high amounts of caffeoyl-CoA are available.
Reactions 3 and 4 show that ChCAD4 is able to produce significant amounts of
caffeyl alcohol even in the presence of coniferaldehyde. Reactions 7 and 8 show that
ChCAD5 produces more caffeyl alcohol than ChCAD4 under the same conditions and,
unlike ChCAD4, it does not reduce coniferaldehyde in the presence of caffealdehyde.
29 These four reactions support the hypothesis that in vivo, ChCAD5 is involved specifically in the production of caffeyl alcohol, whereas ChCAD4 is also capable of caffeyl alcohol production but is less specific.
Figure 3.4: Stacked bar graph of products of in vitro mixed enzyme/substrate reactions. Caffealdehyde (green), coniferaldehyde (yellow), caffeyl alcohol (blue), coniferyl alcohol (red). Reaction 1 contained ChCCR1 with caffeoyl-CoA; reaction 2 contained ChCCR1 with caffeoyl-CoA and feruloyl-CoA; reaction 3 contained ChCAD4 with caffealdehyde; reaction 4 contained ChCAD4 with caffealdehyde and coniferaldehyde; reaction 5 contained ChCCR1 and ChCAD4 with caffeoyl-CoA; reaction 6 contained ChCCR1 and ChCAD4 with caffeoyl-CoA and feruloyl-CoA; reaction 7 contained ChCAD5 with caffealdehyde; reaction 8 contained ChCAD5 with caffealdehyde and coniferaldehyde; reaction 9 contained ChCCR1 and ChCAD5 with caffeoyl-CoA; and reaction 10 contained ChCCR1 and ChCAD5 with caffeoyl-CoA and feruloyl-CoA. In all reactions, CCR and CAD amount was 500 ng and 50 ng, respectively; caffeoyl-CoA/caffealdehyde and feruloyl-CoA/coniferaldehyde concentrations were 30 μM and 10 μM, respectively. Results are from single reactions.
Reactions 5 and 9 show the combined ability of ChCCR1 and ChCAD4 or
ChCAD5, respectively, to produce caffeyl alcohol from caffeoyl-CoA. Comparing the reactions shows ChCAD4 actually produces more caffeyl alcohol than ChCAD5; this
30 was unexpected in light of the previous kinetics results but may reflect the observation that ChCAD4 has a slightly higher catalytic rate at very low levels of caffealdehyde- as
the aldehyde may be supplied at low levels by ChCCR1. Alternatively, this result could
indicate inhibition of ChCAD5 by caffeoyl-CoA.
Reactions 6 and 10 show the combined ability of ChCCR1 and ChCAD4 or
ChCAD5, respectively, to produce caffeyl alcohol and coniferyl alcohol when supplied a
mixture of caffeoyl-CoA and (lower amounts of) feruloyl-CoA. Comparing these
reactions shows that ChCAD4 produces a significant amount of coniferyl alcohol along
with caffeyl alcohol whereas ChCAD5 produces exclusively caffeyl alcohol. However,
ChCAD5 produces less caffeyl alcohol than ChCAD4, and this may be due to the
reasons mentioned in the previous paragraph. Inhibition of ChCAD5 by the buildup of
coniferaldehyde can be ruled out because this is not seen when comparing reaction 7 and 8.
Taken together, the mixed reactions provide in vitro evidence of a pathway from caffeoyl-CoA to caffeyl alcohol involving ChCCR1 and either ChCAD4 or ChCAD5. The pathway involving ChCAD4 is seemingly more efficient but less specific for caffe(o)yl metabolites, whereas the pathway involving ChCAD5 is less efficient but is exclusive for the production of caffeyl alcohol. Considering the mixed reaction results and the expression profiles of the genes in the Cleome seed coat during C-lignin formation, it is likely ChCAD4 and 5 both supply caffeyl alcohol to be polymerized to C-lignin, but
ChCAD5 probably plays a larger role due to its approximately seven-fold higher transcript expression levels. Also, the exclusivity of ChCAD5 for caffeyl alcohol production when incubated with both caffealdehyde and coniferaldehyde (Figure 11,
31 condition 8) may be responsible for the lack of G/C heteropolymer observed in the seed coats (Tobimatsu et al. 2013).
Conclusions
Three genes (ChCCR1, ChCAD4, and ChCAD5) hypothesized to be involved
with C-lignin formation in Cleome seed coats were recombinantly expressed and their
activity with different substrates characterized. ChCCR1 was shown to have a substrate
preference similar to other CCRs, preferring feruloyl-CoA to caffeoyl-CoA. ChCAD4 was
also shown to have activity similar to other CADs, preferring coniferaldehyde to
caffealdehyde. However, ChCAD5 was shown to have a novel preference for
caffealdehyde over coniferaldehyde. The activities of mixed recombinant enzymes fed
mixed substrates were also investigated. These mixed reactions showed ChCCR1 with
either ChCAD4/5 can produce caffeyl alcohol when supplied a combination of feruloyl-
CoA and caffeoyl-CoA, but only ChCCR1 with ChCAD5 exclusively produces caffeyl
alcohol. The expression profile and substrate preference of ChCAD5 make it probable
that it is crucial to accumulation of caffeyl alcohol, and thus C-lignin deposition, in
Cleome seed coats.
32 CHAPTER 4
POLYMERIZATION OF CAFFEYL ALCOHOL AND OTHER MONOLIGNOLS IN
Arabidopsis SEEDLINGS
Objectives
• Determine if accumulation of caffeyl alcohol is sufficient to guarantee
formation of C-lignin by exploring if Arabidopsis seedlings have the ability to polymerize
supplied caffeyl alcohol in an ectopic xylogenesis system, along with 13C- labeled
coniferyl alcohol as a positive control.
• Examine the impact of COMT knockout on lignin production in the above
monolignol feeding system by comparing lignin formation in Arabidopsis comt knockout
seedlings (in which fed caffeyl alcohol cannot be methylated to coniferyl alcohol) in
parallel with Wt.
Methods
4.2.1 Xylem Induction in Arabidopsis Seedlings and Feeding of Monolignols
The particular xylogenesis system (ectopic xylem differentiation in seedlings in
response to brassinosteroid) was selected over protoplast TE induction systems
because it allowed for feeding experiments to be conducted within ten days utilizing
existing mutant lines. Following a modified method from Kondo et al. (2015), seedlings
were grown in six-well culture plates in liquid medium and induced to form ectopic xylem
with bikinin while being supplied specific monolignols (or no monolignols for the
negative control). A laminar flow hood and sterile instruments were used during
manipulation of seeds or seedlings and when changing media. Wild type (Wt) and comt
null mutant seeds were each grown in five experimental conditions (1, no monolignols
33 without bikinin; 2, no monolignols with bikinin; 3, 13C-coniferyl alcohol with bikinin; 4, caffealdehyde with bikinin; and 5, caffeyl alcohol with bikinin). Experimental and control groups were grown in triplicate for a total of 30 separate experimental samples.
In short, sterilized and vernalized Wt (Col) and comt homozygous null mutant
(SALK_135290) seeds (approximately 40 per well) were placed on sterile nylon mesh
(200 micron, componentsupplycompany.com, part number U-CMN-200) disks floating on sterilized liquid medium [2.2 g/L MS salts (no vitamins), 10 g/L sucrose, 500 mg/L
MES, pH to 5.7 with KOH] in 6-well culture plates and grown under 24 h fluorescent lighting at 25 C with 100 rpm shaking. After six days, the original medium was removed and replaced with sterile induction medium [2.2 g/L MS salts (no vitamins), 50 g/L glucose, 1.25 mg/L 2,4-D, 0.25 mg/L kinetin, 15 μM bikinin (except for no-bikinin
controls), pH to 5.7 with KOH] containing the previously specified monolignols (60 μM
each), along with no bikinin or monolignols in the control groups.
To ensure infiltration of induction medium and fed compounds into the leaf tissue
of the seedlings, the mesh disks, with attached seedlings, were flipped upside down so
that shoots were immersed in the liquid medium and a vacuum was applied twice.
Seedlings were flipped right-side-up and grown for another three days under the
previously described growth conditions.
4.2.2 Staining and Microscopic Observation of Seedlings
Several seedlings from each experimental group were collected and stained with
toluidine blue (TBO) following established protocols (Pradhan Mitra and Loqué, 2014).
The stained seedlings were observed by light microscopy and the amount of cell walls
stained for lignin was compared between induced and uninduced groups.
34 4.2.3 Harvesting of Shoot Tissue and Preparation of Cell Wall Residues
Seedlings were removed from induction medium and washed in distilled water four times to remove any remaining medium. The shoots of seedlings were separated from roots using a flat razor blade to cut shoots along the mesh surface. The shoot tissues of each group (pooled from approximately 40 seedlings per well) were collected in 2 mL microcentrifuge tubes and stored at -80C until preparation of cell wall residue.
Cell wall residue (CWR) was prepared by first drying the tissues in a 60 C oven and
pulverizing in a ball mill, then repeatedly extracting with methanol and chloroform until
cell wall extractives had been removed.
4.2.4 Lignin Thioacidolysis and Analysis of Monomer Composition by GC-MS
Lignin content and composition was determined by thioacidolysis. Following
established lab procedures (Lapierre et al. 1995; Yamamura et al. 2012), 7 mg of CWR
from each group (and CWR from vanilla seed as a C-lignin positive sample) was subjected to thioacidolysis and derivatization. Afterwards, the derivatized lignin solutions were analyzed for monomer composition by GC-MS following procedures established in the Dixon Lab, based on the approach of Lapierre et al (1995). Lignin monomer composition was compared across experimental conditions and incorporation of supplied monolignols was measured by quantifying 13C labeled monomers (from 13C-
coniferyl alcohol) or caffeyl monomers.
Results and Discussion
4.3.1 Growth of Arabidopsis Seedlings and Induction of Ectopic Xylem
The growth of Arabidopsis seedlings in six well plates is shown in Figure 4.1.
Seedlings grown under these conditions did not show any noticeable differences in
35 morphology or growth rate due to experimental feedings (eg. supplied monolignols or bikinin). This indicates the seedlings from different treatment groups grew similarly in the system and can be compared without having to account for any gross differences in seedling development.
Figure 4.1: Arabidopsis seedlings grown on mesh disks floating in liquid medium. The seedlings pictured have been growing approximately eight days with two days of xylem induction and feeding. Experimental treatments are labeled on the lids of each plate: A) 13C-coniferyl alcohol with bikinin ; B) caffealdehyde with bikinin; C) caffeyl alcohol with bikinin; D) no monolignols with bikinin; and E) no monolignols without bikinin. The ‘W’ and ‘C’ labels at the top of each plate stand for Wt and comt seedlings, respectively.
36 Microscopic observations of leaves from seedlings stained with TBO are shown in Figure 4.2. The leaves from seedlings without bikinin (A) show blue stained lignin mainly in the veins of the leaf, whereas leaves from seedlings with bikinin (B and C) show blue staining cell walls outside of the veins, indicating formation of ectopic xylem
(red arrows). These ectopic xylem elements are shown at higher magnification in induced leaves (G and H), and featured notable xylem-like spiral cell wall thickenings which are absent in uninduced leaves.
Figure 4.2: TBO staining of Arabidopsis leaves from seedlings grown with bikinin (B, C, E, F, G, and H) and without bikinin (A and D). Areas with ectopic xylem formation are indicated by red arrows in B and C. Ectopic TEs are indicated by red arrows in E and F.
37 The induction of ectopic xylem in bikinin-containing groups indicates these
seedlings are upregulated in lignin biosynthesis and polymerization, similar to other
xylem induction systems in protoplasts or tissue culture. This system has the advantage
that it uses intact seedlings with functioning vascular tissue and, equally important,
experimental manipulations can be performed with existing mutant lines.
4.3.2 Lignin Composition and Incorporation of Supplied Monolignols
The lignin compositions of the seedlings and vanilla seed (C-lignin control) are shown in Figure 4.3. There are notable differences between groups supplied bikinin and no bikinin. Groups without bikinin have less total lignin, which is composed primarily of H subunits, whereas groups supplied bikinin have significantly more total lignin which is composed of much greater amounts of G units. This indicates that the ectopic xylem in bikinin-supplied groups has a significantly different lignin composition from the normal lignin found in uninduced seedlings, and is more similar to the lignin found in non- induced seedlings with more advanced vascular development.
Differences between ectopic lignification in the Wt and comt background can also be seen. Wt plants induced with bikinin produce more total lignin with a higher percentage of G and S subunits, whereas comt seedlings have less total lignin with a lower percentage of G subunits and very little S subunits. These differences in G and S subunits are also seen in the uninduced groups. These results indicate that the differences in lignin composition between lignin mutant backgrounds is conserved and reflected in the bikinin-induced groups.
Clear differences in lignin composition between groups supplied various monolignols were not noted. Importantly, there was no incorporation of C subunits in
38 groups supplied caffeyl alcohol or caffealdehyde. A very small background of C subunits
(0.1-0.25% of total lignin) was noted across all groups, but this may be an artifact of the thioacidolysis process (Fang Chen, personal communication). However, there was some variation in total lignin among monolignol treatments. Generally, Wt seedlings induced with bikinin produce more lignin when not fed any monolignols. Also, both Wt and comt seedlings supplied with caffealdehyde had less total lignin than other groups fed monolignols. This difference between monolignol treatments may be due to the ability of the supplied monolignols to inhibit lignin biosynthesis or other cell wall developmental processes.
Figure 4.3: Graph of lignin composition (left axis) and total lignin area (right axis) across experimental groups. Total lignin area (black), syringyl units (green), guaiacyl units (yellow), catechyl units (red), p-hydroxyphenyl units (blue). Results are averaged from groups grown in triplicate; except for Wt fed caffeyl alcohol with bikinin, Wt fed no monolignols with bikinin, and Vanilla seed- which are biological duplicates. Error bars for total lignin area represent standard deviation.
39 The percent incorporation of 13C label into G and S subunits in Wt and comt
seedlings fed 13C-labeled coniferyl alcohol is shown in Figure 4.4. In the Wt,
incorporation of the label into S subunits is significantly higher than in the comt mutant.
Incorporation of coniferyl alcohol into S units at first seems paradoxical. First, there is
much more G than S lignin in these seedlings, so the seemingly high amount of label
incorporation into S lignin is somewhat unexpected. Second, it might have been
assumed that there would be no incorporation into S lignin in the comt mutant; however
there may be some residual OMT activity or another minor pathway by which a small
amount of labeled coniferyl alcohol could be converted to sinapyl alcohol.
Figure 4.4: Incorporation of 13C label into G (blue) and S (red) subunits in Wt and comt mutant Arabidopsis seedlings supplied 13C-labeled coniferyl alcohol. Results are from biological triplicates, error bars represent standard deviation.
40 Since incorporation of 13C label was found in groups fed 13C-coniferyl alcohol, but
no C-lignin was found in groups fed caffeyl alcohol or caffealdehyde, it can be
hypothesized that this Arabidopsis system is capable of polymerizing supplied
monolignols, but that caffeyl alcohol is not able to be polymerized in this system.
Reasons for the lack of caffeyl alcohol polymerization are not known but are probably
related to the inability of Arabidopsis laccases and/or peroxidases to catalyze radical
oxidation of caffeyl alcohol. It may be that a monolignol polymerization enzyme with
special activity toward caffeyl alcohol is needed in order to polymerize C-lignin, and that
this enzyme is active in the seed coats of Cleome, Vanilla, and other C-lignin containing
species.
Conclusions
A novel xylem induction system in intact Arabidopsis seedlings was used to study
the ability of Wt and comt mutants to produce lignin after being supplied with different
monolignols. Differences in lignin composition were apparent between seedlings with or
without induced xylem, and between Wt and comt backgrounds. This system was
proven to incorporate 13C-labeled coniferyl alcohol into G/S lignin, but did not incorporate caffeyl alcohol into C-lignin. The reason behind the lack of caffeyl alcohol polymerization remains elusive but is probably due to Arabidopsis lacking a laccase or
peroxidase with the needed activity. If such a gene can be identified, the Arabidopsis
seedling system would be an excellent way to test its involvement in C-lignin
biosynthesis.
41 CHAPTER 5
OVERALL SUMMARY AND CONCLUSIONS
C-lignin formation in Cleome was investigated by measuring the transcript levels
of potential lignin pathway–related genes in different tissues and selecting candidate
genes based on their expression patterns during C-lignin deposition. Three differentially
expressed candidate genes for involvement in the monolignol biosynthesis pathway
were selected: ChCCR1 expression reached a maximum during C-lignin deposition in seed coats but was also expressed in the stem, ChCAD4 expression was relatively constant as the seed coat matured and was expressed in the stem, and ChCAD5 expression reached a maximum during C-lignin formation but was expressed at low levels in the stem. Protein modeling of ChCAD5 indicated an amino acid substitution
(L58 to H58) that may explain interaction with potential substrates.
The three candidates were recombinantly expressed for in vitro characterization of enzyme activity in E. coli. Kinetic results for the recombinant enzymes indicated that
ChCAD5 had a novel preference for caffealdehyde over coniferaldehyde, whereas
ChCCR1 and ChCAD4 preferred the classical G-lignin precursors feruloyl-CoA and coniferaldehyde, respectively. Thus it was concluded that ChCAD5 may be involved in
C-lignin biosynthesis in Cleome seed coats.
When recombinant ChCAD4 was supplied a mixture of coniferaldehyde and caffealdehyde, both coniferyl alcohol and caffeyl alcohol were produced, whereas only caffeyl alcohol was produced in similar incubations with ChCAD5. When ChCCR1 and
ChCAD4/5 enzymes were mixed and supplied caffeoyl-CoA and/or feruloyl-CoA, the
ChCCR1/ChCAD5 combination produced some caffeyl alcohol but no coniferyl alcohol,
42 whereas the ChCCR1/ChCAD4 combination produced a significant amount of coniferyl alcohol. Under in vivo conditions, the final monolignol product may be removed by transport across the plasma membrane, affecting the final equilibrium concentrations of
products. With the limitations of the in vitro enzyme approach in mind, the results of the
enzyme/substrate mixing experiments are not inconsistent with a model in which
ChCCR1 and ChCAD5 may work together to create a caffeyl alcohol specific channel in
the seed coat, which would explain the lack of G/C homopolymer found in Cleome seed
coats.
An Arabidopsis xylem induction system in seedlings was used to explore the
ability of Arabidopsis to polymerize caffeyl alcohol. During this experiment, it was found
that neither Wt nor comt knockout seedlings incorporated caffeyl alcohol into lignin,
whereas 13C labeled coniferyl alcohol was incorporated into the G and S subunits of
lignin. This suggests that Arabidopsis lacks the ability to polymerize caffeyl alcohol to C-
lignin, possibly because it is lacking a specific POX or LAC activity to initiate oxidative
radical polymerization of the caffeyl alcohol monomer. However, this special POX or
LAC activity is likely functional in the seed coats of Cleome, vanilla, and other plants
which contain C-lignin. Finding this enzyme(s) will likely be crucial for developing
strategies to engineer C-lignin as a bioproduct in vegetative tissues of crop plants.
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