INVESTIGATION OF PLANT CELL WALL PROPERTIES: A STUDY OF
CONTRIBUTIONS FROM THE NANOSCALE TO THE MACROSCALE IMPACTING
CELL WALL RECALCITRANCE
By
Jacob Dillon Crowe

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Chemical Engineering – Doctor of Philosophy
2017

ABSTRACT
INVESTIGATION OF PLANT CELL WALL PROPERTIES: A STUDY OF
CONTRIBUTIONS FROM THE NANOSCALE TO THE MACROSCALE IMPACTING
CELL WALL RECALCITRANCE
By
Jacob Dillon Crowe
Biochemical conversion of lignocellulosic biomass to fuel ethanol is one of a few
challenging, yet opportune technologies that can reduce the consumption of petroleum-derived
transportation fuels, while providing parallel reductions in greenhouse gas emissions. Biomass
recalcitrance, or resistance to deconstruction, is a major technical challenge that limits effective
conversion of biomass to fermentable sugars, often requiring a costly thermochemical
pretreatment step to improve biomass deconstruction. Biomass recalcitrance is imparted largely
by the secondary cell wall, a complex polymeric matrix of cell wall polysaccharides and
aromatic heteropolymers, that provides structural stability to cells and enables plant upright
growth. Polymers within the cell wall can vary both compositionally and structurally depending
upon plant species and anatomical fraction, and have varied responses to thermochemical
pretreatments. Cell wall properties impacting recalcitrance are still not well understood, and as a
result, the goal of this dissertation is to investigate structural features of the cell wall contributing
to recalcitrance (1) in diverse anatomical fractions of a single species, (2) in response to diverse
pretreatments, and (3) resulting from genetic modification.
In the first study, feedstock cell wall heterogeneity was investigated in anatomical (stem,
leaf sheaths, and leaf blades) and internode fractions of switchgrass at varying tissue maturities.
Lignin content was observed as the key contributor to recalcitrance in maturing stem tissues
only, with non-cellulosic substituted glucuronoarabinoxylans and pectic polysaccharides

contributing to cell wall recalcitrance in leaf sheath and leaf blades. Hydroxycinnamate (i.e.,
saponifiable p-coumarate and ferulate) content along with xylan and pectin extractability
decreased with tissue maturity, suggesting lignification is only one component imparting
maturity specific cell wall recalcitrance.
In the second study, alkaline hydrogen peroxide and liquid hot water pretreatments were
shown to alter structural properties impacting nanoscale porosity in corn stover. Delignification
by alkaline hydrogen peroxide pretreatment decreased cell wall rigidity, with subsequent cell
wall swelling resulting in increased nanoscale porosity and improved enzymatic hydrolysis
compared to limited swelling and increased accessible surface areas observed in liquid hot water
pretreated biomass. The volume accessible to a 90 Å dextran probe within the cell wall was
found to be positively correlated to both enzyme binding and glucose hydrolysis yields,
indicating cell wall porosity is a key contributor to effective hydrolysis yields.
In the third study, the effect of altered xylan content and structure was investigated in
irregular xylem (irx) Arabidopsis thaliana mutants to understand the role xylan plays in
secondary cell wall development and organization. Higher xylan extractability and lower
cellulose crystallinity observed in irx9 and irx15 irx15-L mutants compared to wild type
indicated altered xylan integration into the secondary cell wall. Nanoscale cell wall organization
observed using multiple microscopy techniques was impacted to some extent in all irx mutants,
with disorganized cellulose microfibril layers in sclerenchyma secondary cell walls likely
resulting from irregular xylan structure and content. Irregular secondary cell wall microfibril
layers showed heterogeneous nanomechanical properties compared to wild type, which
translated to mechanical deficiencies observed in stem tensile tests. These results suggest
nanoscale defects in cell wall strength can correspond to macroscale phenotypes.

To those who strive to make the world a better place

iv

ACKNOWLEDGEMENTS

I would like to thank my advisor Dr. David Hodge for all his help, and for providing me
with this opportunity to contribute to the scientific community. I would also like to thank my
committee members Dr. Shi-You Ding, Dr. Ilsoon Lee, and Dr. Bruce Dale for their guidance,
advice, and assistance with pursuing this PhD.
To my lab mates, Dr. Ryan Stoklosa, Thanaphong Phongpreecha, Lisaura MaldonadoPereira, Dr. Daniel Williams, and Dr. Glen Li, I thank you for the friendship and support, and I
wish the best in your graduate and post-graduate careers. To my collaborators Dr. Jacob Jensen,
Dr. Rebecca Garlock-Ong, Dr. Sivakumar Pattathail, Pengchao Hao, and Cliff Foster, I am
grateful to have learned so much while working with you. To the undergraduates Nicholas
Feringa, Rachael Zarger, and Henry Pan, it has been a pleasure mentoring you and working
countless hours in lab together. I expect you all will do well in your academic endeavors. To all
my other colleagues at Michigan State and abroad, I am happy and thankful to have worked with
you over these years.
To all my friends, thank you for always being around for support and encouragement. To
my family, you have made me the person I am today, I am forever grateful. To Amanda and
Maverick, without you two, I would not have made it through graduate school. Thank you for
standing by me through this journey.

v

TABLE OF CONTENTS

LIST OF TABLES

iv

LIST OF FIGURES

v

KEY TO ABBREVIATIONS

xv

Chapter 1: Introduction
Motivation: Sustainable biofuels and chemicals
Background: General outline of lignocellulosic biomass
Structure of lignocellulosic biomass
Plant cell wall polysaccharides
Lignin content and structure
Cell wall deconstruction
Enzymatic hydrolysis
Plant cell wall properties contributing to recalcitrance
Cell wall composition
Morphological fractionation and tissue maturity
Feedstock improvement through genetic manipulation
Characterization of biomass properties
Hydrophilicity
Porosity
Crystallinity
Microscopy
Cell wall extractability
REFERENCES

1
1
5
5
8
12
13
16
18
18
19
20
21
21
24
25
26
28
29

Chapter 2: Identification of developmental stage and anatomical fraction contributions to
cell wall recalcitrance in switchgrass
45
Introduction
45
Materials and methods
48
Field conditions and biomass preparation
48
Pretreatment conditions
49
Composition analysis
50
Enzymatic hydrolysis
51
Confocal microscopy
51
Glycome profiling
52
Results
52
Anatomical fraction- and internode-specific recalcitrance
53
Cell wall composition
55
Non-cellulosic glucose and extractable sugar content
58
Cell wall autofluorescence
60
Glycome profiling
65
Glycome profiling for internode specific extractability
68

iv

Discussion
Conclusions
Acknowledgements
APPENDIX
APPENDIX A: Supplementary Figures
REFERENCES
Chapter 3: Relating nanoscale accessibility within plant cell walls to improve enzyme
hydrolysis yields in corn stover subjected to diverse pretreatments
Introduction
Materials and methods
Biomass and composition analysis
Pretreatments
Enzymatic hydrolysis
Water retention value
Solute exclusion
Enzyme binding
Results and discussion
Composition analysis and enzymatic hydrolysis
Cell wall swelling in response to increasing solvent ionic strength
Accessible volume distributions determined by solute exclusion
Drying-induced pore collapse on cell wall properties and enzymatic
hydrolysis
Enzyme binding
Correlation of properties
Conclusions
APPENDIX
APPENDIX B: Supplementary figures and additional results
REFERENCES

70
75
76
77
78
83

90
90
93
93
94
95
95
96
97
98
98
102
104
107
110
111
115
117
118
123

Chapter 4: Impact of impaired xylan biosynthesis on cellulose microfibril organization in
Arabidopsis thaliana secondary cell walls
129
Introduction
129
Materials and methods
134
Plant lines and growth conditions
134
Composition analysis
134
Glycome profiling
135
Wide angle x-ray scattering
135
Tensile test
136
Scanning electron microscopy
136
Atomic force microscopy
137
Results
138
Irx mutant phenotype, composition, and extractability
138
Relative crystallinity index measured by wide angle x-ray scattering
145
Interfascicular ultrastructure is altered in irx mutants
148
Mechanical properties of irx mutants
156
Discussion
160
Conclusions
165
v

APPENDICES
APPENDIX C: Supplementary figures
APPENDIX D: Additional irx experiments
REFERENCES

vi

167
168
178
184

LIST OF TABLES

Table A.1 : Compiled AIR + destarched cell wall polysaccharides, acetate, and lignin
content for five internodes of three switchgrass anatomical fractions. Samples are presented
as an average (n = 3) with standard deviations following in Table A.2. ...................................... 78
Table A.2 : Standard deviations for compiled AIR + destarched cell wall polysaccharides,
acetate, and lignin content for five internodes of three switchgrass anatomical fractions.
Samples are presented as an average (n = 3) with standard deviations following. ....................... 79
Table 3.1 : Neutral dextran probes used in solute exclusion technique. ................................ 97
Table 4.1 : Summary features observed in SEM and AFM height images for irx mutants.
..................................................................................................................................................... 151
Table C.1 : Roughness analysis and fiber width measurements of delignified irx mutant cell
walls using AFM. Roughness parameters for average roughness (Ra) and root-mean-square
roughness (Rq) measured on AFM height images at multiple image scales. Roughness is utilized
in this case to measure cell wall uniformity. Roughness calculations were performed on three
individual images for each scale (n = 3), with standard deviations shown in table. P values
indicate statistical significance compared to wild type from t-test. ............................................ 174

iv

LIST OF FIGURES

Figure 1.1 : US primary energy consumption 2016. Taken from [2]......................................... 2
Figure 1.2 : Plant stems examined from different scales using multiple microscopy
techniques. (A) Observed whole plant of Arabidopsis thaliana, scale bar 8 cm. (B) Bright field
cross section of Arabidopsis lower stem, scale 25 Âľm. (C) Single sclerenchyma cell from orange
region in (B), scale 5 Âľm. (D) Single parenchyma cell wall from purple region in (B), scale 5
Âľm. (E) Magnified parenchyma cell, scale 2.5 Âľm. (F) Sclerenchyma secondary cell wall
surface, scale 0.75 Âľm. (G) Parenchyma primary cell wall surface, scale 500 nm. (H)
Sclerenchyma secondary cell wall surface, scale 500 nm. Adapted from [25]. ............................. 6
Figure 1.3 : Schematics diagrams of the cell wall. (A) Primary cell wall, and (B) Secondary
cell wall. Adapted from [114]. ........................................................................................................ 7
Figure 1.4 : Model of cellulose organization from individual glucan chains to fibril bundles.
Adapted from [28]........................................................................................................................... 8
Figure 1.5 : Schematic representation of hemicellulose xyloglucan structures. Individual
bond linkages between polysaccharides shown as lines are glycosidic linkages. Adapted from
[30,35]. ............................................................................................................................................ 9
Figure 1.6 : Schematic representation of hemicellulose xylan structures. Individual bond
linkages between polysaccharides shown as lines are glycosidic linkages. Adapted from [30,35].
....................................................................................................................................................... 10
Figure 1.7 : Schematic representation of hemicellulose mannan structures. Individual bond
linkages between polysaccharides shown as lines are glycosidic linkages. Adapted from [30,35].
....................................................................................................................................................... 11
Figure 2.1 : Schematic of switchgrass tillers used for this study. Tillers were harvested above
either the 5th or 6th internode, and then separated into internodes, discarding the nodes and
panicle. Internodes were further manually subdivided into leaves, leaf sheaths, and stems. Notes:
diagram is representative and not to scale. ................................................................................... 49
Figure 2.2 : Enzymatic hydrolysis glucose yields of untreated and alkaline pretreatment of
anatomical fractions. Enzymatic hydrolysis of untreated biomass was performed on AIR +

v

destarched biomass, while alkaline pretreatment was performed using 10% w/w NaOH loading
at 80°C for 1 hr. Glucose hydrolysis yields are presented as the percent of total cell wall glucan
released as monomeric glucose. Enzyme loading was 30 mg protein/g glucan for untreated
fractions and 10 mg protein/g glucan for alkaline pretreated fractions with a ratio of 2:1 CTec3 to
HTec3. Replicates (n = 3) are displayed as averages with standard deviations. .......................... 53
Figure 2.3 : Compositional highlights of (A) lignin (B) hydroxycinnamates (C) minor
polysaccharides, and (D) acetate and arabinan in switchgrass fractions with respect to
internode. Internodes I1-I5 are displayed in ascending order from left to right for each
switchgrass anatomical fraction. All composition values were quantified on an extractives free
basis (AIR). ................................................................................................................................... 55
Figure 2.4 : Native switchgrass non-cellulosic sugar composition with respect to internode.
Internodes I1-I5 are displayed in ascending order from left to right for each switchgrass
anatomical fraction. Compositions are represented as a percent of total dry weight. Replicates (n
= 3) displayed as average with standard deviations. ..................................................................... 59
Figure 2.5 : Examples of differences in macroscopic morphology, cell wall organization,
and aromatic abundance as shown by autofluorescence between switchgrass anatomical
fractions from internode I2. Confocal laser scanning microscopy carried out using laser
emission wavelengths of 405nm and 543 nm. Tissue types are denoted from the following
notation: Ep – epidermis, Fb – fiber bundles, Vb – vascular bundles, Ph – phloem, X – xylem, Pc
– Parenchyma cells. Scale bars represent 200 µm. ....................................................................... 61
Figure 2.6 : Cell wall autofluorescence cross-sections of NaOH-pretreated switchgrass
anatomical fractions from internodes I2, I3, and I4. Confocal laser scanning microscopy
carried out using an overlay of laser excitation wavelengths of 405 nm and 543 nm. Scale bars
represent 200 Âľm. ......................................................................................................................... 63
Figure 2.7 : Abundance of select major non-cellulosic glycan epitopes in chemical extracts
from stem, sheath, and leaf anatomical fractions of internodes I1 (A), and extracts from
stem internodes I1-I5 (B). Extracts were screened by ELISA using a comprehensive suite of cell
wall glycan-directed mAbs as described in materials and methods. Binding response values are
depicted as heat maps with black-red-bright yellow color scheme representing from no binding
to strongest binding. The amount of carbohydrate material recovered per gram of AIR is depicted
in the bar graphs (green) above the heat maps. The panel on the right-hand side of the heat map
shows the groups of mAbs based on the class of cell wall glycan they each recognize. .............. 65

vi

Figure 2.8 : Summary of Pearson’s correlation coefficients between plant cell wall
properties and glucose hydrolysis yields for each anatomical fraction and for pooled
samples. Data demonstrate strong correlations between many properties and yields. ................ 70
Figure 2.9 : Select correlations between data relationships between either lignin content or
ferulic acid against glucose hydrolysis yield represented as scatter plots. (A) 48-hr glucose
hydrolysis yields for NaOH-pretreat biomass versus lignin content, (B) ferulate content in
relation to total lignin, and 48-hr glucose hydrolysis yield with respect to (C) fucan and (D)
arabinan content. ........................................................................................................................... 72
Figure A.1 : Cell wall autofluorescence of switchgrass organ cross-sections from internode
I2. Confocal laser scanning microscopy carried out using laser excitation wavelengths of 405 nm
and 543 nm. Tissue types are denoted from the following notation: Ep – epidermis, Fb – fiber
bundles, Vb – vascular bundles, Ph – phloem, X – xylem, Pc – Parenchyma cells. Scale bars
represent 200 Âľm. ......................................................................................................................... 80
Figure A.2 : Heat map abundance of major non-cellulosic glycan epitopes in oxalate,
carbonate, 1M KOH, 4M KOH, chlorite, and 4M KOH PC extracts from cell walls of stem,
sheath, and leaf samples of internodes I1. Extracts were screened by ELISA using a
comprehensive suite of cell wall glycan-directed mAbs as described in materials and methods.
Binding response values are depicted as heat maps with black-red-bright yellow color scheme
representing from no binding to strongest binding. The amount of carbohydrate material
recovered per gram of AIR is depicted in the bar graphs (purple) above the heat maps. The panel
on the right-hand side of the heat map shows the groups of mAbs based on the class of cell wall
glycan they each recognize. .......................................................................................................... 81
Figure A.3 : Heat map abundance of major non-cellulosic glycan epitopes in oxalate,
carbonate, 1M KOH, 4M KOH, chlorite, and 4M KOHPC extracts from stem internodes
I1-I5. Extracts were screened by ELISA using a comprehensive suite of cell wall glycan-directed
mAbs as described in materials and methods. Binding response values are depicted as heat maps
with black-red-bright yellow color scheme representing from no binding to strongest binding.
The amount of carbohydrate material recovered per gram of AIR is depicted in the bar graphs
(purple) above the heat maps. The panel on the right-hand side of the heat map shows the groups
of mAbs based on the class of cell wall glycan they each recognize. .......................................... 82
Figure 3.1 : Compositional profiles of corn stover subjected to either AHP pretreatment,
LHW, or LHW followed by AHP pretreatment. Compositional profiles for xylan, crystalline
cellulose, and Klason lignin are shown for alcohol insoluble residue (AIR) corn stover and
pretreated biomass. Composition analysis was performed in technical triplicate (n = 3), with
standard deviations shown on figure............................................................................................. 98

vii

Figure 3.2 : Enzymatic hydrolysis glucose yields of pretreated corn stover samples. Results
of never-dried hydrolysis of pretreated samples for 6 hr total hydrolysis time are plotted in blue
and dotted green, while 72 hr total hydrolysis times are plotted above in orange and striped gold.
Solid bars represented hydrolysis conditions using only 20 mg protein/g glucan cellulase loading,
while patterned bars represent hydrolysis conditions using both 20 mg protein/g glucan and 10
mg protein / g glucan hemicellulase loading. Analysis was performed in technical triplicate. .. 100
Figure 3.3 : Cell wall swelling measured using water retention in the absence and presence
of a salt. Water retention value (WRV) of never-dried corn stover at varying pretreatments
measured after incubation in water only (blue bars) or 1 M NaCl (orange bars). Samples were
performed in technical triplicate (n = 3), with standard deviation shown on figure. .................. 102
Figure 3.4 : Accessible volume distributions measured by solute exclusion. Accessible
volumes of never dried corn stover were determined using neutral dextran probes of average
molecular weight range (see Table 3.1), giving a range of effective hydrodynamic radiuses. (A)
AHP-only pretreatment conditions and (B) LHW and the two-stage pretreatments are shown on
the right figure. Samples were performed in technical triplicate (n = 3), with standard deviation
shown on figure........................................................................................................................... 105
Figure 3.5 : Structural model for pretreatment effect on cell wall organization. Transverse
section of cellulose microfibrils for (A) native cell wall, (B) AHP-pretreated cell wall, (C) LHWpretreated cell wall, and (D) two-stage AHP pretreated LHW cell wall. AHP pretreatment results
primarily in reduction in lignin content, promoting the formation of porous regions and inducing
cell wall swelling. LHW pretreatment primarily removes xylan and relocalizes lignin, increasing
accessible surface area. Two-stage pretreatments contain significant reductions in both xylan and
lignin content, increasing porous regions, and in the case of 190°C LHW, sufficient noncellulosic component removal results in cell wall aggregation. Orange squares representing
cellulose microfibrils, green lines representing xylan and gray clouds representing lignin. ...... 107
Figure 3.6 : Impact of oven drying on cell wall properties and enzymatic hydrolysis. (A)
Accessible volumes of oven-dried and rehydrated corn stover were performed similarly to Figure
3.4. (B) Water retention value (WRV) of oven dried corn stover at varying pretreatments. (C)
Hydrolysis yields for oven-dried (OD) samples with 6-hr total hydrolysis time are plotted in
blue, and 72-hr total hydrolysis time are plotted in orange. Comparative hydrolysis yields for
never-dried (ND) 6-hr and 72-hr total hydrolysis time from Figure 3.2 are shown as green dotted
and yellow striped bars respectively. Hydrolysis conditions used 20 mg protein/g glucan CTec3
and 10 mg protein / g glucan HTec3 loading. Experiments were all performed in technical
triplicate, with standard deviations are shown on each figure. ................................................... 108
Figure 3.7 : Bound enzyme fractions of CTec3 on pretreated biomass. Bound enzyme
fractions are shown as a percent of bound enzyme to total enzyme loading. Binding
viii

concentrations were performed in technical duplicate, with standard deviations shown on figure.
..................................................................................................................................................... 110
Figure 3.8 : Comparisons between glucose hydrolysis yields, solute exclusion, water
retention value, and lignin content. (A,B) Glucose hydrolysis yields taken from Figure 3.2
were from 72 hr total hydrolysis time using an enzyme solution of CTec3 and HTec3. Xylan
content is displayed as a range for each linear trend in orange on (A). Accessible volume
distributions (C, D) were used from Figure 3.4, and lignin content was taken from Figure 3.1.
..................................................................................................................................................... 111
Figure 3.9 : Comparisons between glucose hydrolysis yields and (A) bound enzyme
fractions and (B) water retention value. Glucose hydrolysis yields taken from Figure 3.2 were
from 72 hr total hydrolysis time using an enzyme solution of CTec3 and HTec3. WRV were
taken from Figure 3.3, and bound enzyme fractions were taken from Figure 3.7. ................... 113
Figure 3.10 : Comparison of inaccessible pore volume to water retention value. Total
inaccessible pore volume was determined using solute exclusion at the effective pore size of 560
Å and compared to WRV (0 M NaCl) from Figure 3.3. ............................................................ 114
Figure B.1 : Solute-induced swelling of biomass at multiple salt concentrations. Water
retention value (WRV) of never dried and oven dried corn stover at varying pretreatments
measured without salt (0 M NaCl), and after soaking in salt solutions (0.25 -1 M NaCl). (A)
shows AHP-only pretreatment conditions, (B) shows LHW-only pretreatment conditions, (C)
shows two-stage pretreatment conditions, and (D) shows oven-dried pretreatments tested.
Samples were performed in technical triplicate (n = 3), with standard deviation shown on figure.
..................................................................................................................................................... 118
Figure B.2 : Enzyme binding curves of CTec3 on never-dried biomass and linear fits for
bound enzyme fractions. Bound CTec3 concentrations are displayed as individual data points,
with a linear best fit representing the relative ratio of bound enzyme to total enzyme loading. (A)
shows AHP-only pretreatment conditions, (B) shows LHW-only pretreatment conditions, and
(C) shows two-stage pretreatment conditions. Binding concentrations were performed in
technical duplicate (n = 2), with standard deviations shown on figure. ..................................... 119
Figure B.3 : T2 NMR relaxation curves of water interacting with pretreated biomass. (A)
Relaxation curves of AHP-only pretreatment samples, (B) relaxation curves of 160°C LHW and
two-stage pretreatments, and (C) relaxation curves of 190°C LHW and two-stage pretreatments.
Samples were performed and fitted in technical duplicate (n = 2), with standard deviations shown
on figure. ..................................................................................................................................... 119

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Figure B.4 : T2 NMR relaxation curves monocomponent T2 relaxation times. Relaxation
times were determined from a mono-exponential component fit of each relaxation curve.
Samples were performed and fitted in technical duplicate (n = 2), with standard deviations shown
on figure. ..................................................................................................................................... 120
Figure B.5 : 1H-NMR relaxometry of AHP-delignified corn stover showing (A) spin-spin
(T2) relaxation curves and (B) correlation of monocomponent exponential fits of these
relaxation curves to the corresponding WRV and glucose hydrolysis yield. ...................... 120
Figure 4.1 : Growth phenotypes of xylan irx mutants. Growth phenotypes were documented
at week 6 and 9 as indicated. Arrows in SEM cross-section (lower row, irx9 and irx15 irx15-L)
indicate the irregular xylem (irx) phenotype of collapsed xylem vessels. Scale bars for overhead
growth photos are 1.8 cm, side growth photos are 8 cm, and SEM images are 100Âľm. ............ 139
Figure 4.2 : Cell wall polysaccharide composition and xylan chain length. (A) Non-cellulosic
cell wall neutral monosaccharide content of irx mutant stems determined from TFA hydrolysis.
Standard deviations are shown (n = 12, four biological replicates with three technical replicates
for wild type, irx9, irx10, and irx15 irx15-L; n = 6, two biological replicates with three technical
replicates for irx15 irx15-L IRX15-L and irx15 irx15-L IRX15-L). * indicates statistical
significance from wild type (p < 0.05). (B) Xylan degree of polymerization (DP) percent
compared to wild type as reported by prior studies.
141
Figure 4.3 : Crystalline cellulose, lignin, and glycome profile analysis of cell wall material
of irx mutant stems. (A) chemical quantification of crystalline cellulose content (B) and ABSL
lignin content of cell wall material. Standard deviations are shown (n = 12, four biological
replicates in three technical replicates each for wild type, irx9, irx10, and irx15 irx15-L; n = 6,
two biological replicates with three technical replicates for irx15 irx15-L IRX15-L and irx15
irx15-L OE-IRX15-L). * indicates statistical significance compared to wild type (t-test, p < 0.05).
(C) Cell wall material of wild type, irx9, irx10, and irx15 irx15-L inflorescence stems was
extracted sequentially with increasing chemical severity using glycome profiling. A selected
subset of antibodies and extracts are shown, while the full profile can be found in Figure C.2 143
Figure 4.4 : WAXS diffraction pattern of irx mutant stems and relative crystallinity index.
(A), representative WAXS diffraction patterns of wild type, irx9, irx10, and irx15 irx15-L stems
showing the diffraction peaks for the [110], [1-10], [200], and [400] crystal faces. (B), relative
crystallinity indexes (RCI) of the [200] peak face calculated from the WAXS diffraction pattern,
such as the ones in (A). Standard deviations are shown (n = 4, biological replicates for wild type,
irx9, irx10, and irx15 irx15-L; n = 4, two biological and two technical replicates for irx15 irx15L IRX15-L and irx15 irx15-L OE-IRX15-L). * indicates statistical significance compared to wild
type (t-test, p < 0.05). .................................................................................................................. 145

x

Figure 4.5 : Ultrastructure of irx mutants observed with SEM and AFM. Scanning electron
microscopy images of sclerenchyma cell surface texture in native wild type Arabidopsis (A,G,S),
irx9 (B,H,T), irx10 (C,I,U), irx15 irx15-L (D,J,V), irx15 irx15-L IRX15-L (E,K,W), and irx15
irx15-L OE-IRX15-L (F,L,X) stems. Atomic force microscopy images of sclerenchyma cell wall
surface texture after delignification in wild type Arabidopsis (M,Y), irx9 (N,Z), irx10 (O,AA),
irx15 irx15-L (P,BB), irx15 irx15-L IRX15-L (Q,CC), and irx15 irx15-L OE-IRX15-L (R,DD)
stems. Scale bars are 2 Âľm for low magnification SEM (A-F), 1 Âľm for intermediate
magnification SEM (G-L), and 200 nm for high magnification SEM (S-X), 500 nm for the 2.5
Âľm AFM images (M-R), and 200 nm for the 1.0 Âľm AFM images (Y-DD). ............................ 149
Figure 4.6 : Fiber widths measurements of delignified irx mutant secondary cell walls using
AFM. Fibril widths were determined from well-resolved features in 2.5 Âľm image sizes, using
five measurements for each of three individual images for each sample (n = 15). Averages of
fiber measurements are displayed as a horizontal bar for each sample on figure. ...................... 152
Figure 4.7 : Roughness analysis, fiber orientation, and 3D topography measurements of
delignified irx mutant cell walls using AFM. (A), Roughness parameters for average roughness
(Ra) and root-mean-square roughness (Rq) measured on AFM height images at 0.5 Âľm image
scale. P values indicate statistical significance compared to wild type from t-test. (B), Microfibril
orientations measured by dispersion of fibers distributed throughout an entire image at the 1.0
Âľm image scale. Standard deviations for (A) and (B) are shown on figure for three different
images (n = 3). (C), 3D surface profiles of wild type and irx9 demonstrating fiber alignment in
wild type observed at three different image angles compared to irx9. Scale bar represents 100 nm
and image scale is 0.5 Âľm. .......................................................................................................... 154
Figure 4.8 : Height distribution profiles of delignified irx mutant cell walls using AFM.
Gaussian distributions highlighting the distribution of height topography of each sample at 1 Âľm
scale. Distributions of each image triplicates (n = 3) are shown on figure. Gaussian distributions
of all image scales can be found in Figure C.11. ....................................................................... 156
Figure 4.9 : Mechanical strength of xylan irx mutant stems. Tensile stress to breaking of stem
tissue measured by universal tensile test. Box and whiskers plot is the result of twenty replicates
of one biological batch of stems (n = 20). * indicate statistical significance from wild type
(p<0.1), ** indicates statistical significance from wild type (p<0.0005). .................................. 157
Figure 4.10 : Merged nanomechanical AFM imaging of irx mutants. Nanomechanical
mapping of modulus (MPa), adhesion (nN), deformation (nm), and dissipation (eV) were scaled
to the same absolute intensities for direct comparison of distributions. Groups were designated
based upon qualitative features, with irx9, irx10, and irx15 irx15-L placed in group 1, and irx15
irx15-L IRX15-L and irx15 irx15-L OE-IRX15-L placed in group 2. Scale bar represents 200 nm,
with all images corresponding to a 1.0 Âľm size. ......................................................................... 159
xi

Figure 4.11 : Schematic of analytical techniques utilized in this study, and proposed
connections between individual techniques. ........................................................................... 164
Figure C.1 : SEM images of parenchyma cells surface texture in irx mutants. Wild type
(A,G,M), irx9 (B,H,N), irx10 (C,I,O), irx15 irx15-L (D,J,P), irx15 irx15-L IRX15-L (E,K,Q),
and irx15 irx15-L OE-IRX15-L (F,L,R) stems labeled. Scale bars are 2 Âľm for low magnification
(A-E), 1 Âľm for intermediate magnification (G-L), and 200 nm for high magnification (M-R).
..................................................................................................................................................... 168
Figure C.2 : Extended dataset of the glycome profile of irx mutants. Glycome profile of
Arabidopsis cell wall material extracted sequentially with increasing chemical severity using
oxalate, carbonate, 1M KOH, 4M KOH, acidic chlorite, and 4M KOHPC pretreatments as
defined in materials and methods. Antibody binding strength is based upon optical density, with
binding intensity presented as a color gradient from black (low binding) to red (intermediate
binding), and yellow (strongest binding). Green bars on the top indicate the amount of
carbohydrate recovered per gram of cell wall AIR for each extraction. ..................................... 169
Figure C.3 : WAXS diffraction curve processing for crystallinity and microfibril diameter
calculation. (A), WAXS diffraction curve fitting for quantification of relative crystallinity index,
with ten Gaussian fits used to fit the major crystal faces and account for amorphous
contributions. (B), Relative crystallinity index determined using the peak area method as
described in the methods. (C), Sherrer microfibril diameter determined using the width at peak
half height of the [200] crystal face. ........................................................................................... 170
Figure C.4 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis wild type. Scale bars for SEM are 2 Âľm for low magnification (left column), 1 Âľm
for intermediate magnification (middle column), and 200 nm for high magnification (right
column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the
1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the
0.25 Âľm images (fourth column). ............................................................................................... 171
Figure C.5 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx9. Scale bars for SEM are 2 Âľm for low magnification (left column), 1 Âľm for
intermediate magnification (middle column), and 200 nm for high magnification (right column).
Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the 1.0 Âľm
images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the 0.25
Âľm images (fourth column). ....................................................................................................... 171
Figure C.6 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx10. Scale bars for SEM are 2 Âľm for low magnification (left column), 1 Âľm for

xii

intermediate magnification (middle column), and 200 nm for high magnification (right column).
Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the 1.0 Âľm
images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the 0.25
Âľm images (fourth column). ....................................................................................................... 172
Figure C.7 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx15 irx15-L. Scale bars for SEM are 2 Âľm for low magnification (left column), 1
Âľm for intermediate magnification (middle column), and 200 nm for high magnification (right
column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the
1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the
0.25 Âľm images (fourth column). ............................................................................................... 172
Figure C.8 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx15 irx15-L IRX15-L. Scale bars for SEM are 2 Âľm for low magnification (left
column), 1 Âľm for intermediate magnification (middle column), and 200 nm for high
magnification (right column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first
column), 200 nm for the 1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third
column), and 50 nm for the 0.25 Âľm images (fourth column).................................................... 173
Figure C.9 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx15 irx15-L OE-IRX15-L. Scale bars for SEM are 2 Âľm for low magnification
(left column), 1 Âľm for intermediate magnification (middle column), and 200 nm for high
magnification (right column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first
column), 200 nm for the 1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third
column), and 50 nm for the 0.25 Âľm images (fourth column).................................................... 173
Figure C.10 : Distribution profiles of microfibril orientations for an entire image. Sample
image distribution profiles for each irx mutant and wild type. Dispersion was calculated based
upon the width of the gaussian distribution. ............................................................................... 174
Figure C.11 : Height distribution profiles of delignified irx mutant cell walls using AFM.
Gaussian distributions of height ranges at each image scale are displayed as individual plots of
three images for each sample. ..................................................................................................... 175
Figure C.12 : Example calculation of microfibril width. Microfibril widths were calculated
using the build in section Tool in Nanoscope. Well resolved microfibril features were used to
determine horizontal fibril widths............................................................................................... 176
Figure C.13 : Merged nanomechanical AFM imaging of all irx mutants. Nanomechanical
mapping of modulus (MPa), adhesion (nN), deformation (nm), and dissipation (eV) were scaled
xiii

to the same absolute intensities for direct comparison of distributions. Scale bar represents 200
nm, with all images corresponding to a 1.0 Âľm size. .................................................................. 177
Figure D.1 : Water retention values and extractive insoluble residue of cell wall material of
irx mutants. (A), Water retention values (WRV) of alcohol insoluble residue (AIR) with and
without chemical extraction with 1M KOH and extractive insoluble components (EIC) represents
solid residue after sequential extraction with 4M KOH after chlorite delignification. (B-D),
Composition of neutral monosaccharides, crystalline cellulose, and ABSL lignin content of
residue remaining after 4M KOHPC extraction. Standard deviations are shown with two
biological replicates with three technical replicates (n = 6). ...................................................... 180
Figure D.2 : Enzymatic digestibility of cell wall material of irx mutant stems. Enzymatic
hydrolysis monomeric sugar yields of glucose (A) and xylose (B) of unpretreated AIR on a basis
of % monomeric sugar observed per total available polysaccharide (g/g*100). Standard
deviations are shown (n = 6, biological duplicate with technical triplicate). ............................. 181
Figure D.3 : Cell wall compositions of Arabidopsis wild type and WAXS relative
crystallinity index after chemical extraction. (A) cell wall neutral monosaccharide content
determined from TFA hydrolysis, (B) crystalline cellulose content, and (C) ABSL lignin content
of wild type AIR, and sequentially extracted with carbonate and 1M KOH. (D) Relative
crystallinity indexes of the [200] peak face calculated from the WAXS diffraction patterns in
wild type native, AIR, and sequentially extracted with carbonate and 1M KOH. Standard
deviations are shown (n = 3, biological replicates. * indicates statistical significance compared to
wild type (t-test, p < 0.05)........................................................................................................... 182

xiv

KEY TO ABBREVIATIONS

ABSL – acetyl bromide soluble lignin
Ac - acetate
AFEX- ammonia fiber expansion
AFM -atomic force microscopy
AG – arabinogalactan
AGP – arabinogalactan pectins
AHP -alkaline hydrogen peroxide
AIR – alcohol insoluble residue
Ara- arabinan
BSA – bovine serum albumin
CBM – carbohydrate binding module
CESA – cellulose synthase complexes
CI – crystallinity index
DI – de-ionized
DP – degree of polymerization
DSC – differential scanning calorimetry
DSC-TP – differential scanning calorimetry thermoporemetry
EA – extractive ammonia
EIC – extractive insoluble components
ELISA – enzyme-linked immunosorbent assay
Ep- epidermis
ESK – ESKIMO
FA – ferulate
Fb – fiber bundle
FSP – fiber saturation point

xv

Fuc – fucan
G – guaiacyl
Gal – galactan
GAX – glucuronoarabinoxylan
GFP – green fluorescent protein
GH – glycosyl hydrolase
GHG – greenhouse gas
GlcA – glucuronic acid
GUX – glucuronyl transferases
GX – glucuronoxylan
GXMT – glucuronic acid methyl transferase
H – p-hydroxyphenyl
HG – homogalacturanan
HPLC – high performance liquid chromatography
Irx – irregular xylem
LHW – liquid hot water
LUT – look up table
mAb – monoclonal antibody
Man – mannan
MeGlcA – Methyl glucuronic acid
MLG – mixed linkage glucan
ND – never-dried
NMR – nuclear magnetic resonance
NREL – national renewable energy laboratory
OD – oven-dried
Pc – parenchyma cells
PC – post-chlorite
pCA – p-coumarate

xvi

Ph – phloem
RCI -relative crystallinity index
RFS – renewable fuel standard
RG – rhamnogalacturonan
Rha – rhamnan
S – syringyl
SEM – scanning electron microscopy
TBL – TRICHOME BIREFRINGENCE-LIKE
TEM - transmission electron microscopy
TFA – trifluoracetic acid
Vb – vascular bundle
WAXS – wide angle x-ray scattering
WRV – water retention value
X – xylem
XG – xyloglucan
XGA – xylogalacturanan
Xyl – xylan

xvii

Chapter 1 : Introduction
Motivation: Sustainable biofuels and chemicals
There exists a growing political, cultural, and economic movement towards pursuing
policy and technologies to mitigate the human carbon footprint on climate change. From the
2015 Paris Agreement, predictions of a global reduction of 50% of total greenhouse gas (GHGs)
emissions by 2050 are required to limit the mean temperature rise between 2 -2.5°C [1]. Fossil
derived energy accounts for 80% of total global energy consumption [2], with both changes to
energy utilization and source required to meet the maximum GHG predicted for a limited
temperature rise [3]. In 2016, an estimated 10% of total energy consumption was derived from
renewables in the United States (Figure 1.1), with 15% of total electrical power generated from
renewables such as solar and wind. Advances in solar and wind technologies are anticipated to
reduce implementation costs in the coming years [4], allowing renewables to compete
economically on the electrical grid with fossil sources.

1

Figure 1.1 : US primary energy consumption 2016. Taken from [2].

Energy for transportation accounts for 29% of total energy consumption in the United
States (Figure 1.1), with 92% of that energy derived from petroleum sources. Current
technological prospects exist to offset energy dense liquid transportation fuels with alternative
sources such as electrification of vehicles, hydrogen production, or other non-drop-in fuels [5],
however significant advances to energy storage technology, production, and infrastructure for
development and distribution are still required for successful implementation of these
technologies. Biofuels represent a challenging, yet opportune technology that can supplement
petroleum derived transport fuels using existing infrastructure [6], provide reductions in carbon
emissions, and improve non-petroleum derived human wealth and development during the global
transition away from fossil fuels [7].

2

The Renewable Fuel Standard (RFS) was implemented as part of the Energy Policy Act
of 2005, and expanded under the Energy Independence and Security Act of 2007, setting
mandatory volumes of renewable fuels to offset petroleum derived fuels [8]. Current volumes of
first generation corn ethanol are near 15.3 billion gallons a year in the US, with an average of
43% reduction in GHG emissions compared to gasoline [9]. Corn ethanol, while establishing
logistics and transport infrastructure for ethanol distribution and blending, is volume limited due
to land allocation between corn purposed for food production versus ethanol production.
Cellulosic ethanol, represents another source of reduced carbon for bioethanol production with
the potential for higher reductions of GHGs compared to gasoline, availability of feedstock, with
an estimated 1.3 billion tons of economically feasible and sustainable feedstock [10], and is
slated for 16 billion gallons of total production in the RFS by 2022 [8]. Cellulosic bioethanol is
divided into three major feedstock unit operations to produce cellulosic bioethanol: (1) feedstock
pretreatment to improve cell wall accessibility for (2) enzymatic hydrolysis to produce hexose
and pentose sugars capable of providing anaerobic microbes a high titer stream of sugars for (3)
fermentation of sugars to bioethanol or other value-added products.
Recent progress in cellulosic bioethanol processing has resulted in the construction of
pioneer second generation cellulosic ethanol plants [11], however the current predicted volume
requirements of cellulosic ethanol set by the RFS are not being met. This is due in part to the
difficulties encountered in upstream processing and scale up [12], as well as recent increases in
oil and gas drilling in the form of hydraulic fracturing and oil sands reducing petroleum prices
and lowering the incentive for investing in alternative forms of energy security [13].
If cellulosic bioethanol is to come to fruition on a large scale, technological
improvements need to be implemented to reduce capital costs and improve process economics

3

such as value-added co-production such as lignin valorization [14, 15]. Pioneer plants have
identified key technical challenges associated with continuous upstream production and
achieving plant operating capacity, which may be addressed in time through ‘learning by doing’,
however, other challenges outside of the industrial plant still exist [11]. Development and
implementation of upstream feedstock supply chains is both a technical and capital investment
hurdle; unlike other commodity chemicals with established logistics, lignocellulosic biomass
lacks existing infrastructure [12]. In addition, lignocellulosic feedstocks have poor transport and
handling properties, with low densities, high moisture content, and poor-flowability
characteristics, often necessitating on-field drying and densification for long distance transport
[16].
Understanding feedstock properties from the fundamental perspective may provide an
avenue for addressing a few outstanding challenges limiting economic feasibility of
lignocellulosic bioethanol; biomass heterogeneity is a significant variable impacting upstream
feedstock handling properties, as well as downstream biomass conversion economics. At the root
of feedstock heterogeneity, the plant cell wall is a complex and diverse structure that is still not
fully understood. As a result, the focus of this dissertation will be to investigate factors
influencing plant cell wall properties in order to understand and improve cell wall hydrolysis for
the production of fermentable sugars. In Chapter 2, biomass heterogeneity will be investigated
in anatomical and developmentally distinct fractions of a single switchgrass cultivar to
understand cell wall variability between fractions. Furthermore, these differences will be
investigated in the context of cell wall deconstruction towards improving biomass processing.
In Chapter 3, the role of chemical pretreatment impacting nanoscale cell wall properties
is investigated to understand cell wall factors impacting cellulose accessibility and enzymatic

4

digestibility. Nanoscale porosity has shown to play an integral role in improving cell wall
deconstruction, and will be investigated in the context of changes to cell wall structure and
rigidity.
In Chapter 4, the role of individual components impacting secondary cell wall structure
is investigated using a series of xylan deficient irregular xylan (irx) mutants. The purpose of this
work is to examine the role of altered polymer structure to understand the role of higher order
interactions impacting cell wall organization, as well as demonstrate synergy between multiple
analytical techniques to characterize altered cell wall structure.

Background: General outline of lignocellulosic biomass
Structure of lignocellulosic biomass
A large fraction of the total terrestrial plant mass resides in plant cell walls, an integral
component of all plant cells. Cell walls are important features of plant cells that perform essential
features ranging from defining cell shape and cell type, to providing a physical barrier from
extracellular interaction [17]. The general model of plant cell wall is comprised of both a primary
and secondary cell wall, with vastly different functions for each.
The primary cell wall makes up a thin physical layer that encapsulates cells and inhibits
movement during creep, or expansion of the cell wall during plant growth [18]. Composed
primarily of cellulose microfibrils imbedded in a matrix of complex polysaccharides, the primary
cell wall resembles a web of fibrous cellulose weaving and forming distinct lamellar layers [19,
20]. Polysaccharides associated with cellulose microfibrils in primary cell wall interact via noncovalent interactions, such as hydrogen bonding or van-der Waals interactions, and form a
complex 3D imbedded matrix. Xyloglucans, one class of hemicelluloses within the primary cell
wall of dicots, are an essential polysaccharide involved in cell wall loosening during extension.

5

The primary mechanism of interaction with cellulose microfibrils is hypothesized through
association of a series of parallel microfibrils together via a tethered network model, however,
recent evidence in xyloglucan deficient xxt1/xxt2 mutants has put that model into question [21,
22]. Pectins are also present in the primary cell wall, with highly hydrophilic structures likely
contributing to reductions in cellulose-cellulose interactions, and promoting inter-microfibril
spacing [23].

Figure 1.2 : Plant stems examined from different scales using multiple microscopy
techniques. (A) Observed whole plant of Arabidopsis thaliana, scale bar 8 cm. (B) Bright field
cross section of Arabidopsis lower stem, scale 25 Âľm. (C) Single sclerenchyma cell from orange
region in (B), scale 5 Âľm. (D) Single parenchyma cell wall from purple region in (B), scale 5
Âľm. (E) Magnified parenchyma cell, scale 2.5 Âľm. (F) Sclerenchyma secondary cell wall
surface, scale 0.75 Âľm. (G) Parenchyma primary cell wall surface, scale 500 nm. (H)
Sclerenchyma secondary cell wall surface, scale 500 nm. Adapted from [25].

6

The secondary cell wall is present only in fully developed cells post elongation, and is
deposited in certain cell types to provide mechanical strength and resistance to osmotic
pressures. Secondary cell walls are significantly thicker than primary cell walls, constitute a
large portion of the total cell wall mass, and are composed predominately of cellulose,
hemicelluloses, and lignin [24]. There are two types of secondary cell walls, the parenchymatype and the sclerenchyma-type secondary cell wall. Parenchyma-type are usually partially
lignified and act as support tissues, while sclerenchyma-type secondary cell walls are found in
differentiated treachery elements and sclerenchyma fibers [25]. Sclerenchyma-type secondary
cell walls are fully lignified, and formed by alternating layers of sheet-like cellulose microfibrils
accompanied by hemicelluloses and separated by lignified layers [26]. Many genes specific to
secondary cell wall formation are highly active in sclerenchyma-type secondary cell walls only
[24]. Example schematics of both the primary and secondary cell wall are highlighted in Figure
1.3.

Figure 1.3 : Schematics diagrams of the cell wall. (A) Primary cell wall, and (B) Secondary
cell wall. Adapted from [114].

7

Plant cell wall polysaccharides
The major classes of polysaccharides in the cell wall include cellulose, hemicelluloses,
and pectins. Cellulose is linear chain of glucosidic β-1,4-linked glucose units comprised of a
repeat unit of two anhydroglucose rings, referred to as cellobiose, with a variable chain length
anywhere from 10,000 – 15,000 individual glucose units depending on the source material [27].
Intra-chain hydrogen bonding between hydroxyl groups and adjacent oxygen molecules
maintains the linear configuration of cellulose, with interchain hydrogen bonding and van-der
Waals between individual cellulose chains resulting in parallel stacking and formation of
cellulose elementary fibrils [28]. Elementary fibrils further aggregate and form larger
microfibrils through hydrogen bonding. A traditional two-phase model has been used to describe
cellulose structure, with regions of highly ordered cellulose crystals formed termed crystalline
cellulose, while cellulose microfibrils of low structural order are referred to as amorphous
cellulose [29].

Figure 1.4 : Model of cellulose organization from individual glucan chains to fibril bundles.
Adapted from [28].

Hemicelluloses compared to cellulose are significantly more heterogeneous, comprised
broadly of xyloglucans, xylans, mannans, and mixed linkage glucans. Xyloglucans (XyG) as
described earlier are primarily found in the primary cell wall of dicots and are hypothesized to
8

interact and form cellulose- xyloglucan networks. The predominant xyloglucan structure in
dicots is the fucogalactoxyloglucan structure (FucogalactoXyG), while monocots favor the
arabinogalactoxyloglucan (arabinogalactoXyG) structure [30]. Xyloglucan backbones are
comprised of β-1,4-lined glucose units similar to cellulose, that are substituted at various
positions by xylosyl residues depending upon plant species. In most species, xylosyl residues are
further substituted with galacturonosyl, arabinoxyl, or other glycosyl residues, with galactosyl
residues often further substituted by fucosyl and acetyl units depending upon the species [30].

Figure 1.5 : Schematic representation of hemicellulose xyloglucan structures. Individual
bond linkages between polysaccharides shown as lines are glycosidic linkages. Adapted from
[30,35].

Xylans are the major hemicellulose present in secondary cell walls of monocots and
dicots. Glucuronoxylan (GX) is the predominant structure of xylan in dicots, consisting of β-1,4linked xylosyl residues as the backbone, decorated with ι-1,2-linked glucuronic acid (GlcA) and
4-O-methyl-glucuronic acid (MeGlcA) [31], and O-acetyl groups at the O-2 and O-3 position
[32]. A tetrasaccharide sequence of 4-β-D-Xyl-(1-4)-β-D-Xyl-(1-3)-ι-L-Rha-(1-2)-ι-D-Gal-(1-4)D-Xyl

is found in some dicots including Arabidopsis, and is thought to play a terminating role in

xylan chain length during biosynthesis [33, 34]. Glucuronoarabinoxylan (GAX) is the main

9

xylan found in monocot grass primary and secondary walls. In addition to a similar backbone
and GlcA and MeGlcA substitution and acetylation patterns, GAX also contains arabinosyl
substitutions to the backbone at the Îą-1,2 and Îą-1,3 positions [35]. Unique to grass cell walls,
arabinosyl units may be further substituted through ester and ether linkage of hydroxycinnimates,
acting as an anchor point for lignification [36].

Figure 1.6 : Schematic representation of hemicellulose xylan structures. Individual bond
linkages between polysaccharides shown as lines are glycosidic linkages. Adapted from [30,35].

Mannans are the major hemicellulose in secondary cell walls of gymnosperms [37], and
are classified into homomannan, glucomannan, galactomannan, and galactoglucomannan [35].
Homomannans and galactomannan have a backbone formed by β-1,4-linked mannosyl units,
while glucomannan and galactoglucomannan contain both mannose and glucose units in a β-1,4linked configuration [30]. Mannosyl residues may be substituted with ι-1,6-linked galactosyl
units, and can be acetylated at the O-2 and O-3 positions [38, 39]. Mixed linkage glucans
(MLGs) are found in the cell walls of grasses, and are thought to play a role as a storage polymer
[40]. MLGs are unsubstituted, linear chains of glucose units with randomly distributed β-1,4linked cellobiose, cellotriose, and cellotetrose sub-units. Although not as prevalent as a

10

hemicellulose, MLGs can constitute an appreciable fraction of cell polysaccharide in secondary
cell walls of grasses [41].

Figure 1.7 : Schematic representation of hemicellulose mannan structures. Individual bond
linkages between polysaccharides shown as lines are glycosidic linkages. Adapted from [30,35].

Pectins are the third group of polysaccharides present in the cell wall, and are thought to
facilitate cell wall strength through non-covalent and covalent interactions with other cell wall
polymers [42]. There are four major groups of pectins in non-aquatic plants, broken into
homogalacturanans (HG), xylogalacturanan (XGA), and rhamnogalacturanan I and II (RG I &
II). The ratio of pectin structures is highly varied depending upon plant species, however HG is
usually the most prevalent pectin, followed by RG I and other pectins constituting smaller
fractions [43]. HG is comprised of unbranched Îą-1,4-linked D-GalUA that can be further
substituted with other sugar residues such as xylosyl units in XGA, or with acetyl groups at the
O-2 and O-3 positions or methyl esterification at the C-6 carboxyl position. RG II consists of a
stretch of HG backbone that is substituted with four complex size chains, containing a total of 12
different types of glycosyl residues [44]. RG II is highly conserved among vascular plants, [45]
and likely plays a critical role in maintain cell wall integrity. RG I is the only pectin lacking a
HG backbone, and instead is a branched polymer with Îą-1,4-D-Gal-Îą-1,2-L-Rha repeat. Rha
residues are further substituted with either β-1,4-linked galactan branches that are further

11

substituted with branched arabinose units, or arabinogalactan side chains, with acetylation of the
backbone occurring at the O-3 position [42].
Lignin content and structure
Lignin is a complex heteropolymer found in plant secondary cell walls that provides
compression strength, limits cell wall mobility, and prevents extracellular interaction with the
cell wall. Lignin content in biomass, although varying considerably depending upon plant
species and tissue maturity can range from 10-30% dry weight [46]. Chemically, lignin is
derived mainly from three p-hydroyphenyl alcohols, p-coumaryl, coniferyl, and sinapyl alcohols
which are precursors for the lignin monolignols p-hydroxyphenyl (H), guaiacyl (G), and syringyl
(S) units. Other non-monolignol aromatic monomers such as hydroxycinnimates p-coumatate,
ferulate, and sinapate found in grasses, are derived from the lignin biosynthesis pathway are also
incorporated into lignin, and can constitute an appreciable fraction of quantifiable lignin
monomers depending upon species [47].
Lignin is thought to form within the cell wall through radical oxidative polymerization
catalyzed by peroxidases and laccases [48]. A range of bond configurations exist within lignin,
however the primary bonds quantified are categorized into labile β-O-4 and ι-O-4 aryl-ether
bonds, and condensed C-C bonds such as β-5 phenylcoumaran, β-β resinol, and 5-5 aryl-aryl
bonds, or 4-O-5 and 5-O-4 biphenyl ethers which are significantly more resistant to chemical
degradation [49].
In addition to the bond configurations described, monocot grasses incorporate
hydroxycinnimates into lignin through ester and ether bonding. Specifically, ferulate monomers
can be incorporated into lignin by various ether and C-C bonds [47], however they are also
known to ester bind to arabinosyl residues on GAX, forming covalent linkages between
12

hemicellulose and lignin [50]. Ferulates may also dimerize, forming diferulates, and may play an
important role as initiation sizes for lignin formation or anchoring lignin in grass cell walls [51].
P-coumarates are also in abundance in grass cell walls, and are mainly bonded through
esterification to the gamma-position of phenylpropanoid sidechains of S lignin, and unlike
ferulate, form few crosslinked structures [52]. P-coumarates are thought to aid in the formation
of syringyl-rich lignins during later stages of lignification, however their exact role is unknown
[50]. Generally, grasses have a more condensed lignin structure with higher phenolic hydroxyl
content compared to dicots [36], and coupled with alkali labile bonds formed from
hydroxycinnimates are highly soluble in alkali solution, with large fractions of total lignin
soluble at room temperature [53].
Cell wall deconstruction
Biomass recalcitrance, imparted by cell wall structural and compositional features, is a
key obstacle in successful employment of lignocellulosic biomass as a feedstock. Limited
accessibility of polysaccharide degrading enzymes within the cell wall makes biomass
conversion on an industrially relevant time scale unfeasible in native biomass [54], requiring the
alteration of biomass feedstock properties through biochemical and mechanical pretreatment
approaches. The goal of pretreatment is to remove or alter structural and compositional
impediments in order to improve the rate of enzymatic hydrolysis [55]. Approaches to improving
enzymatic hydrolysis yields can be classified into modifications of feedstock variables or
modification of pretreatment variables. In the case of pretreatment variables, approaches are
taken to improve either mass transfer, reaction kinetics, or a combination of both through
application of physiochemical conditions to improve feedstock accessible surface area, porosity,
or alter substrate affinity and accessibility for depolymerization [56].

13

Chemical pretreatments can be further classified based upon the general effect on cell
wall structure, with acidic pretreatments preferentially solubilizing hemicelluloses and relocalizing lignin and alkaline or oxidative based pretreatments solubilizing or depolymerizing
lignin. A third class of pretreatment grouping ionic liquid and organosolv pretreatments
physically fractionate cell wall components. Most pretreatments can be further categorized based
upon main chemical mode of action and reactant nucleophilicity, as described in the review by
Ong. et al. [56]. In general, the following pretreatments presented all are dependent upon similar
variables such as reaction temperature, time, pH, catalyst or reactant loading, and solids content,
although the importance of each variable may be significantly different depending upon
feedstock and pretreatment [55, 57].
Acid pretreatments such as dilute acid are well-studied techniques and have been used
commercially in the manufacture of furfural [55], with a main mode of action through
electrophilic hydrolysis of ester cross-links and β-ether bonds in lignin, as well as hydrolysis of
glycosidic linkages in hemicelluloses to form monomers and oligomers, which can further be
dehydrated to form furans [58]. Lignin has been observed to aggregate and localize along the cell
wall surface after dilute acid pretreatment [59, 60]. Dilute acid pretreatment may also impact
cellulose crystal structure by altering hydrogen bonding interactions between glucan chains [61].
Neutral chemistry hydrothermal pretreatments such as liquid hot water pretreatment
(LHW) and steam explosion pretreatment initially utilize hydronium ions derived from water
autoionization to liberate weak biomass-derived acids such as acetyl groups, which in turn act as
weak acids in solution [62]. Hemicellulose backbone and substitutions are hydrolyzed to a lesser
extent, generally released as oligomers, while only labile bonds are broken in lignin [63]. Similar
to dilute acid pretreatments, lignin is relocalized as globules on the outer cell walls at high LHW

14

pretreatment severity [64], although minimal lignin is solubilized and generally increases in total
mass fraction significantly after pretreatment [58]. LHW pretreatment may reduce cellulose
crystallinity at high temperatures [65], however low to moderate severity pretreatment has shown
no significant change in crystallinity [66].
Alkaline and oxidative pretreatments act through either saponification reactions of
available esters in lignin and hemicelluloses, as well nucleophilic substitution or oxidation of βether bonds [56]. At high alkali loadings, hemicelluloses may also be extracted through peeling
reactions [67], although side chain substitutions and acetate can still be extracted at low alkali
loadings [68], as well as significant fractions of lignin in grasses, which are particularly
susceptible [53] to moderate alkali pretreatment conditions.
Alkaline hydrogen peroxide (AHP) pretreatment is an oxidative pretreatment derived
from existing pulp and paper applications [69, 70]. AHP pretreatment is unique in that it is
carried out at room temperature [71], targeting mainly lignin bonds and solubilizing a large
fraction of lignin from the lignocellulosic matrix. β-ether bonds are cleaved at higher peroxide
loadings along with reductions in lignin molecular weight contributing to lignin solubility and
removal in grasses [72, 73]. Hemicellulose removal is present due to the caustic nature of
pretreatment causing saponification of acetate and ester bonds, increasing hemicellulose
solubility [74]. Cellulose content and crystallinity remains relatively unchanged in AHP
pretreatment, indicating that the major changes in biomass are associated with loosening of the
cell wall matrix [75].
Ammonia Fiber Expansion (AFEX) pretreatment falls generally under alkaline
pretreatments, and utilizes anhydrous ammonia under high pressure, followed by an explosive
decompression to disrupt cell wall structure [76]. AFEX results in changes to cell wall
15

ultrastructure [77] through targeting of ester linkages between lignin and hemicelluloses, as well
as deacetylation and partial depolymerization of hemicelluloses [78]. In addition, cellulose decrystallization is observed, forming cellulose III allomorphs, which has been shown to increases
digestibility significantly [79], and is the basis of newer Extractive Ammonia (EA) pretreatments
[80].
Pretreatment technologies that dissolve or fractionate lignocellulose include organosolv
pretreatments and ionic liquid pretreatments, which will be briefly mentioned for completeness.
The main mode of ionic liquid pretreatment, similar to EA pretreatment is the disruption of
cellulose hydrogen bonding and de-crystallization of cellulose and removal from the cell wall
[81]. Currently, there exist multiple ionic liquid pairs for application in lignocellulosic biomass
pretreatment, which are highlighted in a recent review [82]. Organosolv pretreatment utilizes a
combination of organic solvents and aqueous solutions to extract lignin and hemicelluloses from
lignocellulose, and similarly to ionic liquids, possess a wide range of organic solvent-aqueous
pairings [83]. The majority of organosolv pretreatments delignify through cleavage of β-ether
linkages in lignin, followed by reductions in lignin molecular weight. Hemicellulose glycosidic
linkages are also targeted, resulting in hemicellulose solubilization, while cellulose dedecrystalization also occurs [84], giving organosolv pretreatments significant similarity with
multiple other pretreatments mentioned earlier.
Enzymatic hydrolysis
Following pretreatment, lignocellulosic biomass is subjected to depolymerization to
break sugars down from polymers into monomers. The main mode of action in this process is the
breaking of glycosidic linkages between individual cellulose and hemicellulose subunits, which
can be performed either thermochemically via acid and heat, or enzymatically using enzymes to

16

target individual bonds. Enzymatic hydrolysis is favored for lignocellulosic processing, due in
part to the production of inhibitors in acid hydrolysis limiting fermentability [85], however the
presence of a multitude of bond configurations requires multiple enzymes. In nature,
microorganisms such as bacteria and fungi produce biomass-degrading enzymes, secreting
cellulolytic enzymes from classes of glycoside hydrolases (GH), which work together
synergistically to depolymerize polysaccharides [86]. The primary enzymes required for
lignocellulosic hydrolysis are cellulases and xylanases, however accessory enzymes play an
important role in improving both enzyme dosing and overall hydrolysis yields [71, 87]. In
addition, cellulose structure can impact enzymatic hydrolysis yields, with amorphous cellulose
having higher digestibility compared to crystalline cellulose [54]. Changes in crystalline
cellulose allomorph imparted by pretreatments such as EA [80] also have shown to increase
hydrolysis yields [79].
Enzymatic depolymerization effectiveness may be inhibited through either nonproductive adsorption to lignin or other cell wall polymers, product inhibition, limited enzyme
accessibility, or enzyme deactivation [88]. Non-productive adsorption and limited enzyme
accessibility are two factors directly related to feedstock properties, with lignin and to a lesser
extent xylan believed to act as a sheathe and physically occlude cellulase accessibility to
cellulose via steric hindrance [89, 90]. Delignification pretreatments alone have shown marginal
increases in cellulose accessibility, indicating lignin indirectly limits cellulose accessibility
through limiting accessibility to xylan, which in turn limits cellulose accessibility [88, 91]. There
also exists a limit for improving cellulase accessibility through pretreatment, with harsh
pretreatment conditions exhibiting a decrease in enzyme accessibility, likely due to cell wall
collapse or aggregation [92].

17

Non-productive adsorption is thought to occur due to hydrophobic and electrostatic
interactions between lignin and cellulase carbohydrate binding modules (CBM), promoting a
high affinity for lignin [93]. Increases in carboxylic acid content and therefore higher net
negative charge in lignin has shown decreases in non-productive binding of cellulases [94], while
adding negative charges to cellulases can significantly decrease lignin affinity [95, 96].
Approaches using proteins such as Bovine Serum Albumin (BSA) as lignin blockers prior to
enzymatic hydrolysis have also reduced non-productive adsorption of cellulases [97], with BSA
thought to reduce lignin affinity by interacting with hydrophobic charges. Pretreatment altering
lignin properties may also participate in changing non-productive adsorption, and may provide
avenues for tailoring pretreatment conditions to minimize non-productive adsorption [94, 98].

Plant cell wall properties contributing to recalcitrance
Cell wall composition
Lignin is often cited as the most common cell wall component to inhibit enzymatic
digestibility, with strong negative correlations observed before and after pretreatment in multiple
pretreatments [73, 99, 100]. Lignin monomer composition has been examined as a variaible
impacting enzymatic hydrolysis, with some studies showing increased S/G monolignol content
positively correlates with yield [99, 101]. This is not a universally observed trend however [56],
leading to the conclusion that other properties as well as pretreatment type may impact S/G ratio.
Esterified ferulate content in grasses has shown positive correlations with hydrolysis, and may be
due to the more labile ester linkages associated with ferulates enabling cell wall loosening [102,
103].
Because the contribution of lignin towards recalcitrance is substantial in most feedstocks,
deriving non-lignin correlations to digestibility can be difficult to deconvolute from lignin. The

18

substitution pattern of arabinan has recently been proposed to impact recalcitrance, with higher
abundances of GAX arabinose substitution correlating positively to digestibility [104].
Additionally, substituted xylan has been correlated to increased extractability, which may be an
indicated of a less tightly crosslinked cell wall matrix [105]. Similar conclusions have been
drawn in a study utilizing sequential extraction of switchgrass, which showed the greatest
increase in hydrolysis yield after alkaline extraction has removed significant fractions of
substituted xylan [106].
Morphological fractionation and tissue maturity
Biomass morphological fractions and varying tissue maturities can impart significant
differences in feedstock composition [105], and present an opportunity for either modifying
harvesting methods or employing physical fractionation prior to biomass pretreatment and
digestibility. Physical fraction of corn stover to stem pith, stem rind, and leaves resulted with the
highest yield in stem pith, followed by leaves, and stem rind for both no pretreatment and liquid
hot water pretreatment [107]. Similar conclusions have been observed in physical fractionation
of sugarcane stems [108]. In herbaceous feedstocks, multiple studies have all come to similar
conclusions that stem fractions are less digestible compared to leaf fractions [109, 110].
Tissue maturity can also impact enzymatic hydrolysis yields. Internode maturity in
switchgrass stems decreased in digestibility in more mature internodes [111], as did maize
digestibility with stem age [112]. As a general trend, structural carbohydrates and lignification
increase in proportion of total biomass with tissue maturity, while soluble sugars and other
components decrease [113].

19

Feedstock improvement through genetic manipulation
The past decade has seen significant strides in understanding the genes and transcription
factors involved in plant cell wall biosynthesis and regulation, and are the subject of numerous
reviews [24, 114]. Feedstock manipulation aims to improve feedstock sugar yields through
reductions in cell wall recalcitrance, while maintaining normal feedstock growth. General
approaches to feedstock improvement include modification of cell wall component structure,
abundance, or localized expression in tissues, with a few examples of each highlighted in the
proceeding text. Modification of cellulose abundance and crystallinity are two approaches that
have been taken, however over-expression of CESA complexes resulted in reduced cellulose
content due to silencing of other genes [115], while reductions in crystallinity were accompanied
by dwarfed phenotypes [116]. Given the essential role cellulose plays in maintaining cell wall
strength, approaches manipulating cellulose content and structure may not be advantageous.
Typically, down regulation of major hemicellulose components such as xylan backbone
synthesis [24] has resulted in improved sugar digestibility at the expense of severe growth
phenotypes; however, regulation of tissue specific expression of xylan biosynthesis to vessel
cells xylan deficient mutants irx7, irx8, and irx9 has shown rescued phenotypes coupled with
increased digestibility [117]. Alteration of xylan side chain substitution, specifically reduction of
uronic acid side chains on hemicellulose in the gux mutants showed increased digestibility with
no noticeable growth phenotypes [118], while reductions in other side chains such as acetylation
[38, 119] had dwarfed phenotypes.
Reductions in lignin content follow similar approaches to xylan modification, with severe
reductions in lignin causing phenotypic dwarfism. Vessel specific expression of cinnamate 4hydroxylase in Arabidopsis resulted in reductions in lignin content without sacrificing plant

20

fitness [120], while the creation of a competitive metabolic pathway reducing shikimate, a key
enzyme in the lignin pathway, also significantly reduced yields without dwarfism [121]. Changes
to lignin monomer content have also shown some success [122], as have incorporation of
chemically labile ester linkages via the design of a novel lignin monomer to improve
pretreatment effectiveness [123]. Engineering of lignin structure without sacrificing functionality
is an active research field, working to reduce lignin degree of polymerization, reduce lignin
hydrophobicity, reduce lignin-carbohydrate covalent bonds, increase chemically labile linkages,
and improve lignin valorization [124].

Characterization of biomass properties
Hydrophilicity
Water imbedded within the cell walls causes cell walls to swell and soften, acting as a
plasticizer and separating individual polymers through hydrogen bonding and electrostatic
interactions [125]. Other solvents can swell cell walls; however, hydrogen bond donors show
preferential swelling [126]. Individual cell wall components have different affinities for water,
with hydrophobic lignin generally having the lowest affinity, followed by cellulose,
hemicelluloses, and hydrophilic pectins [127, 128]. Available hydroxyl and carboxylic acid
groups found on polysaccharide chains are the predominant cell wall features water bonds with,
forming hydrogen bonded layers [129]. In cellulose, water may absorb within a cellulose
elementary fibril, with water molecules hydrogen bonded between individual elementary
microfibril chains [130].
Because of this phenomenon, cellulose is sometimes thought of as a hydrate, given the
native state of cellulose fibers can be expected to contain up to 8% moisture by weight, which is
equivalent to one water molecule per glucose unit [125]. Furthermore, initial water added to a

21

cellulosic fiber immediately adsorbs and hydrogen bonds to elementary fibrils [131]. Water
closest to cell wall surfaces has an altered chemical environment due to hydrogen bonding and
does not have a freezing point [132], while water beyond the first few molecular layers has a
depressed freezing point, which can be used to quantify pore size of nonporous materials [133].
The nature of available chemical bonds on polysaccharide chains also presents the ability
for polyelectrolytes to impact biomass-water interactions and impact cell wall swelling. In the
case of salts, conversion of hydroxyl groups and carboxyl groups to the salt form can impact the
pH of the surrounding solution, as well as induce osmotic diffusion gradients [134]. This
phenomenon is most prevalent in carboxymethylated super-absorbent pulps and cellulose gels,
however chlorite delignified pulps have exhibited similar changes in cell wall swelling,
indicating that unlignified cell walls behave similar to a hydrogel [135]. One hypothesis for
hydrolysis effectiveness has been proposed around sufficient delignification inducing loss of cell
wall rigidity and subsequent expansion [73], with cell wall swelling in response to
polyelectrolytes providing one proxy for measuring cell wall rigidity to osmotic pressure.
Fiber saturation point (FSP) is defined as the water weight where the cell wall of a
sample becomes saturated with bound water, and is used as a metric for measuring bulk
hydrophilicity. Multiple techniques including pore dilution by solute exclusion, vapor pressure
adsorption, or gravimetric filter pad techniques have been employed to precisely measure FSP,
with multiple techniques yielding similar values [136, 137].
Differences in feedstock, composition, or tissue type can impact water sorption behavior.
In addition, water sorption can be dramatically impacted by chemical pretreatment, depending on
the chemical and ultrastructural changes of the cell wall. Water retention value (WRV) is one
standardized way of determining the pseudo-FSP, through determination of bound water in a
22

sample via gravitational separation [138]. Cell wall compositions and response to pretreatment
can both contribute to cell wall hydrophilicity, enabling the usage of WRV to represent multiple
feedstock properties.
Recent work correlating compositional relationships to WRV in a genotypically diverse
group of maize samples concluded that individual contributions from both lignin components
and hemicellulose components can be related to WRV, limiting weighting of individual
contributions towards overall WRV [103].
WRV has been successfully employed to demonstrate increases in cell wall swelling of
AHP delignified and LHW pretreated grasses along with correlate subsequent increases in
enzymatic hydrolysis [139]. WRV has also been used to represent cellulose accessibility in fiber
sheets, and captured loss in cellulose accessible area imparted by hornification processes [140,
141]. From these studies, it was concluded that WRV represents cell wall response to diverse
structural and compositional features which likely impact cellulose accessibility and indirectly
impact enzymatic digestibility.
Nuclear magnetic resonance (NMR) relaxometry represents another avenue for
measuring biomass-water interactions, using time domain spin-spin (T2) NMR relaxometry
techniques. In addition to gauging general biomass recalcitrance, specific information on the
chemical environment of water may be extracted [142]. Water constraint has been proposed as a
general term to describe the extent of hydrogen bonding between water and the cell wall, with
increased water constraint correlating to increases in accessibility [143, 144]. Relaxation time as
a single metric similar to WRV has shown to correlate well with enzymatic hydrolysis yields in
model cellulose components [143], and has also been used to describe tissue-specific chemical

23

states of water in sugarcane [145], however to date has not been directly correlated to enzymatic
hydrolysis of lignocellulosic biomass.
Porosity
The idea of cell wall hydrophilicity can be extrapolated to ultrastructural features of the
cell wall, namely the presence of void spaces or pores contributing to total cell wall accessible
surface area. In the context of lignocellulosic biomass, pore formation may result from chemical
pretreatment, altering void spaces and available surface area for enzyme penetration and
subsequent hydrolysis. The lamellar structure of plant cell walls allows for the assumption of a
simple slit model to define cell wall pores, and has been used in prior studies to define pore
geometry [146, 147]. Solute exclusion is one technique to measure pore accessibility of various
sizes of probe molecules to obtain a pore size distribution [136].
The technique works through dilution of probe solution indicating interaction with pores,
and is based upon the assumptions of equal concentration of probe solution within the pore and
surrounding bulk fluid, as well as full accessibility of a probe to a pore. Solute exclusion is
limited by irregular pores under-contributing to total pore volumes, known as the ‘ink-bottle’
effect, or the presence of ionized groups excluding probes via electrostatic interaction [148]. The
solute exclusion technique has been successfully utilized to describe pore changes resulting from
delignification [149], sulfite pretreatment [150], and hornification [151].
Differential scanning calorimetry thermoporemetry (DSC-TP) is another technique
utilized to measure pore distributions, and is based upon changes in freezing point depression of
water associated with the cell wall discussed earlier. The relationship between specific melting
enthalpy of water associated with the cell wall is related to a specific pore geometry using the
Gibbs-Thompson equation. This technique has been successfully employed in observing pore
24

changes during drying [133], as well as recently to identify the impact of alkaline and
autohydrolysis pretreatment on porosity [65]. Cryoporismetry is not limited to DSC-TP, with 1H
NMR cryoporisemetry having been utilized following similar assumptions in dilute acid
pretreated corn stover [152]. One of the key advantages of solute exclusion and cryoporisemetry
is the ability to measure samples in a wet state, while older techniques for measuring pore
distributions such as nitrogen adsorption or mercury porismetry require dry samples, which
imparts altered pore properties [147].
Crystallinity
The amount of inter- and intra- molecular hydrogen bonds between cellulose elementary
microfibrils gives rise to ordered crystalline cellulose microfibril structures. To date, there are
four crystalline cellulose allomorphs, with cellulose I the most abundant and naturally occurring,
found as cellulose Iι in some bacteria and regenerated pulps, and Iβ in plant cell walls. Other
allomorphs are derived from either chemical treatment with alkali (II) or ammonia (III), coupled
with solubilization and regeneration (II), or through heating of cellulose III (IV) [28]. Cellulose
crystalline structure and abundance can be observed using X-ray diffraction techniques, or
through solid-state 13C NMR.
As described earlier, amorphous regions of cellulose have less ordered hydrogen bonding
and do not contribute to the crystallinity of cellulose, with a ratio of crystalline cellulose to
amorphous contribution called crystallinity index (CI) or relative crystallinity index in this text
(RCI). RCI is used to interpret changes to cellulose structure after physiochemical or biological
treatments, and is a non-destructive way to observe crystalline cellulose changes [29].
The relative amount of crystalline cellulose compared to amorphous cellulose has been
linked to reductions in hydrolysis rates, with higher RCI values corresponding to more
25

crystalline cellulose and slower initial hydrolysis yields [153, 154]. Altering the crystalline
hydrogen bond network as mentioned earlier has shown improvements to yield [79], along with
de-crystalization of cellulose to amorphous cellulose increasing hydrolysis rates by an order of
magnitude [155]. Crystallinity has also shown to have no correlation with overall hydrolysis
yields [156], however can still be used to gauge relative cellulose organization.
Native cell wall crystallinity can be affected by perturbations to components within the
cell wall during growth, although higher order interactions between individual cell wall
components are not well understood. A few notable examples of irregular cellulose include
cellulose deficient mutants from KORRIGAN1 [157] and CESA1 and CESA3 mutants [116],
reduced crystalline cellulose organization in lignin deficient mutants [158], and disruption in
microfibril angle observed in xylan modified mutants [159].
Microscopy
Direct observation of cell wall organization via microscopy techniques is another
approach towards characterizing biomass, and enables visualization a cell wall surface and
structural features at the micron or nanoscale.
Transmission electron microscopy (TEM) is a widely used microscopy technique that
utilizes electrons passing through a sample to derive images, and allows for high-resolution
views of the cell walk ultrastructure and of different cell wall layers [148]. One limitation of
TEM is sample thickness, limiting analysis of three-dimensional topography that may be present
in samples. Scanning electron microscopy (SEM) provides high-resolution topographic
information with minimal sample preparation aside from drying and coating. One advantage of
SEM over TEM is the ability to examine three-dimensional features of cells often overlooked
when taking small cross-sections using TEM.
26

Atomic force microscopy (AFM) utilizes direct contact between a sample and a probe tip
to produce a three-dimensional image. The probe tip is attached to an oscillating cantilever,
allowing changes in tip contact height to be observed as a change in cantilever oscillation [160].
AFM also has the advantage of capturing nanomechanical properties of the sample surface
through direct contact, allowing for direct measurement of properties such as modulus force,
deformation, adhesion force, and dissipation [161]. Alternatively, complex molecules such as a
cellulase CBM may be linked to an AFM tip to alter the chemical interaction between the tip and
surface, allowing for examination of a sample surface from the perspective of an external probe
interaction [162]. The major limitation of AFM is disruption of native sample state during
preparation. While AFM sampling can be performed in vacuum, fluid, or air, thin samples of
relative height uniformity are required to effectively image, requiring some cutting or sample
isolation prior. Additionally, in the case of imaging secondary cell walls, cellulose microfibril
layers are imbedded within coated lignin layers, often requiring a chemical pretreatment such as
acid-chlorite pretreatment prior to imaging, which will certainly impact native structure and
interpretation of results relative to a native perspective [163].
Confocal microscopy is another common microscopy technique, using laser
excitation to visualize cell wall surfaces. Often, microscopy is coupled with staining techniques
[164], or along with immunohistochemical techniques to highlight local abundance of certain
components [165]. Confocal microscopy may also be used to autofluorescence cell walls, and
obtain information about chemical makeup [166, 167], however is limited by some lack of
specificity between individual components [168]. More recent endeavors using confocal
microscopy coupled GFP-labelling of individual cell wall components to obtain special

27

distribution information [169], or GFP-labelling of proteins involved in cell wall biosynthesis to
obtain real-time visualization of cell wall synthesis and organization [170, 171].
Cell wall extractability
Biomass response to deconstruction can also be approached through investigation of cell
wall extractability of certain components. Glycome profiling, using a suite of plant cell wall
glycan-directed monocolonal antibodies (mAbs) represents one technique to identify cell wall
deconstruction patterns and understand cell wall organization [172, 173]. This technique is useful
in application for identifying structural differences between feedstocks [106], understanding
pleiotropic impacts of gene mutants on cell wall extractability [174], and evaluating pretreatment
effectiveness in different feedstocks [175, 176]. More recently, glycome profiling has been used
to identify structural changes during tissue maturity [105], as well as understand recalcitrant
unhydrolyzed carbohydrates in residual solids after enzymatic hydrolysis [87]. While a powerful
technique with multiple applications, glycome profiling is limited to recognizing individual
clades of epitopes in most cases, as the specific substitution patterns of many epitopes are not
known [175, 177].

28

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Chapter 2 : Identification of development stage and anatomical
fraction contributions to cell wall recalcitrance in switchgrass

This work has been published as original research in July 2017, in Biotechnology for Biofuels,
Volume 10, Issue 1, Page 184.

Introduction
Development of renewable sources of fuels and chemicals is necessary to improve energy
security and mitigate climate change effects resulting from greenhouse gas emissions [1, 2].
Lignocellulosic biomass, which includes dedicated bioenergy crops and agricultural residues, is a
widely available and largely untapped source of reduced carbon in the production of renewable
fuels and chemicals [3]. A current major challenge in deconstructing plant cell walls are plantevolved mechanisms for resisting microbial and animal degradation of plant cell walls, termed
biomass recalcitrance [4]. Within a feedstock, abundance and density of cell type, structure and
substitution of cell wall polymers, and organization of polymers within the cell wall can all
contribute to biomass recalcitrance [4].
Plant cell walls account for the majority of lignocellulosic feedstock dry mass, and are
comprised of primary and in some cases secondary cell walls depending upon the tissue type.
Primary cell walls are found in growing cells, and contain cellulose microfibrils, pectins, and
hemicellulose polysaccharides. Secondary cell walls are significantly thicker than primary cell
walls, provide mechanical strength and structural reinforcement to the cells, and are found
predominantly in fiber and vascular cells post cell enlargement [5], and represent the majority of
the mass of lignocellulosic feedstocks. Secondary cell walls contain cellulose microfibils,
hemicelluloses, and lignin, an aromatic heteropolymer comprised of phenylpropanoid subunits.

45

In monocot grasses, the primary hemicellulose components are glucuronoarabinoxylans
(“xylans”) containing arabinosyl, acetyl, and uronosyl substitutions on a β-1,4 linked xylan
backbone [6]. The arabinofuranosyl residues in grass xylans can also be covalently linked via
ferulic acid ester crosslinks to lignin or other xylans, and this phenomenon has been
hypothesized to play an important role in promoting cell wall rigidity and decelerating expansion
[7]. Mixed-linkage glucan and xyloglucans are also present as minor hemicellulose components
in cell walls, and may account for a minor fraction of non-cellulosic glucan found in mature
tissues [6, 8]. Lignin content and composition is often cited as one of the predominant
contributors of cell wall recalcitrance [9], however there also exists a body of literature that has
examined the role of hemicelluloses [10], structural pectins [11], and polymer interactions within
the cell wall as other significant contributors to cell wall recalcitrance [12].
Switchgrass (Panicum virgatum L.) is a well-studied C4 perennial grass envisioned as a
promising herbaceous feedstock to supply North American cellulosic biorefineries along with
other monocot grasses such as miscanthus (Miscanthus x giganteus), and corn stover (Zea mays)
[13, 14]. Switchgrass is an attractive feedstock due to high production yields, low energy and
nutrient inputs, wide adaptability, and short growing time [15]. Switchgrass grows as a clonal
modular organism, with the tiller forming the main clonal module and phytomers consisting of
multiple shoot meristems. Shoots are comprised of leaf blades, sheath leaves, stem nodes and
internodes, auxiliary buds, and ligules [16]. Grasses such as switchgrass grow through elongation
of stem internodes, each of which is linked at the base node to a single leaf sheath and blade.
Analysis of individual internodes, as opposed to bulk plant material, is useful as internodes
represent sequential stages of maturity during plant development, affecting cell wall composition
and abundance of cell type within biomass [17]. Basal internodes tend to have greater lignin and

46

cellulose content coupled with lower digestibility compared to internodes near the top of a single
tiller [18]. Tissue maturity in relation to harvest times may impact switchgrass digestibility [19],
including time left on field post maturation [20]. In addition, there are significant differences in
compositions of stem, sheath, and leaf anatomical fractions, with the stem generally containing
more lignin and being less digestible compared to the leaf [21, 22]. The distribution, structure,
and extractability of non-cellulosic cell wall glycans in miscanthus stem and leaf organs sampled
at distinct maturities indicated that hemicellulose structure and extractability varied with tissue
maturity, and may account for significant non-lignin contributions to recalcitrance [23].
Within-plant heterogeneity for graminaceous feedstocks such as switchgrass has
important impacts on several important processing variables for a biorefining process. Ranges of
particle size distribution, composition, hygroscopicity and drainability, chemical pretreatment
effectiveness, and extent of enzymatic hydrolysis can all result from differing responses of
biomass to comminution. Physical fractionation, employed either on-field during harvest or at
the biorefinery, is one potential route to addressing feedstock challenges to potentially improve
agronomic, logistics, and processing-conversion outcomes. Limited work has been published on
biomass fractionation in the context of pretreatment and hydrolysis. In one study, disc refining
followed by air classification of corn stover yielded pith and depithed stover fractions that were
subjected to dilute acid pretreatment or mild base pretreatment, quantifying mass losses and
sugar release response during acid treatment [24]. Other work identified significant differences
in enzymatic hydrolysis response of corn stover stems compared to other fractions [25-27].
Studies investigating tissue-specific responses have been assessed for liquid hot water
pretreatment of corn stover [28] and wheat straw [29], although the only industrially-relevant
fractionation approach that have been studied are alteration of leaf-stem ratio in wheat straw

47

[30]. With AFEX pretreatment, the impact of knife-milling and sieving of corn stover [31] as
well manual separation of stem, sheath, leaf, and cob have been assessed [32], and found to yield
fractions with differing compositions and responses to deconstruction.
The objective of this study is to characterize within-plant heterogeneity by identifying
key compositional differences between switchgrass anatomical fractions at different internode
maturities and relating compositional differences to enzymatic digestibility response following
mild alkaline pretreatment. Structural differences are evaluated by localization of phenolics
within tissue cross-sections before and after alkaline pretreatment using confocal microscopy,
while hemicellulose and pectin structure and extractability are evaluated using glycome profiling
following chemical extraction.

Materials and methods
Field conditions and biomass preparation
Switchgrass (var. “Forestburg”, upland ecotype) was manually harvested post-anthesis
above either the 5th or 6th internode Figure 2.1 from a field at Michigan State University (East
Lansing, MI; 42°42’48.80” N by 84°28’1.41”W) between 17-19 September, 2014. During
harvest, intact harvested tillers were dried indoors at room temperature. After harvest was
completed all tillers were dried overnight in a 45°C oven. Internodes were then manually
separated into leaves, leaf sheaths, and stems, while the nodes and panicle were discarded
Figure 2.1. Whole fractions were air dried to a moisture content of ~5% (g H2O/g total), and
particle size was reduced using a Wiley Mini-Mill (Thomas Scientific, Swedesboro, NJ) to pass a
30-mesh screen. Samples were stored in dry, airtight bags until further use.

48

Figure 2.1 : Schematic of switchgrass tillers used for this study. Tillers were harvested above
either the 5th or 6th internode, and then separated into internodes, discarding the nodes and
panicle. Internodes were further manually subdivided into leaves, leaf sheaths, and stems. Notes:
diagram is representative and not to scale.

Pretreatment conditions
Alkaline pretreatment was performed in 8 mL borosilicate glass tubes (Thermo Fisher
Scientific, Waltham, MA) with rubber lined caps. Alkali loading was performed at a 10% (g
NaOH/g dry biomass) loading at 10% (g biomass/g solution) loading, giving a 0.25 M NaOH
concentration. Samples were vortexed and immersed in an 80°C water bath for 1 hr, with mixing
every 15 min. After the elapsed pretreatment time, solids were separated from the alkaline liquor
49

and washed with de-ionized water until pH neutral. Following washing, samples were rinsed
with absolute ethanol and allowed to air dry at 45°C to a moisture content <5%, with mass loss
following pretreatment determined gravimetrically.
Composition analysis
Extractives-free cell wall material (i.e., alcohol insoluble residue or “AIR”) was isolated
following extraction and de-starching according to Foster et al. [33]. All composition procedures
were performed on de-starched AIR, except for the determination of extractable sugars. Minor
polysaccharide content (rhamnan, mannan, galactan, fucan, non-cellulosic glucan) was
determined using the alditol acetate method with minor changes [33]. Cell wall acetate, glucan,
xylan, arabinan, and Klason lignin content were determined for AIR using the NREL/TP 51042618 protocol [34] with minor modifications [35]. Thioacidolysis was used to determine lignin
monolignol yields [36]. Extractable free glucose, sucrose, and starch were determined on the
original untreated biomass as described by Santoro et al. [37].
Alkaline saponification was used to determine the p-hydroxycinnimic acid content, as
described previously [38], with modifications to the quantification method. Briefly, 0.5 g of
biomass was treated with 25 mL of 3 M NaOH in a sealed pressure tube at 121°C for 1 hr to
release esterified p-coumarate (pCA) and esterified or etherified ferulate (FA). Liquid samples
were centrifuged at 13000 x g for 3 min. Next, 1 mL of alkaline liquor was taken, and pH
adjusted to pH 1.5 using concentrated H2SO4 (Sigma-Aldrich Corp., St. Louis, MO). Samples
were centrifuged at 13000 x g for 3 min to remove any precipitated solids, and subsequently
analyzed by HPLC (Agilent 1100 Series) with an Aminex HPX-87H column (Bio-Rad, Hercules,
CA, USA) using a mobile phase of 5 mM H2SO4 with a 10% (v/v) ACN organic modifier.
Quantification was performed using a diode-array detector with a UV wavelength of 210 nm.

50

Standards containing FA and pCA ranged from 0.05 mg/mL to 1 mg/mL. To account for
precipitation of pCA and FA with pH adjustment, solubility with respect to pH curves were
determined, with R2 linear coefficients of 0.999 for both FA and pCA observed from the range of
pH 1.4 - 1.8. Samples were analyzed in biological triplicate for reproducibility.
Enzymatic hydrolysis
Enzymatic hydrolysis was performed at 2.5% solids loading (g AIR biomass/g solution)
in 1.5 mL microcentrifuge tubes (Posi-Click, Denville Scientific, Holliston, MA). Enzymatic
hydrolysis was performed using 10 mg protein/g glucan for pretreated biomass and 30 mg
protein/g glucan for untreated, de-starched, extractives-free biomass using CTec3 and HTec3
(Novozymes A/S BagsvĂŚrd, Denmark), at a CTec3:HTec3 ratio of 2:1. A buffer solution of 50
mM citric acid (pH 5.20) was used to maintain pH, and samples were incubated at 50°C with
orbital mixing at 180 rpm for either 6 hr or 48 hr. Samples were centrifuged at 13000 x g for 3
min post-incubation and filtered using 22 Âľm mixed cellulose-ester filters (EMD Millipore,
Billerica, MA). Glucose and xylose were quantified by HPLC (Agilent 1100 Series) with an
Aminex HPX-87H column (Bio-Rad, Hercules, CA, USA) with a mobile phase of 5 mM H2SO4.
Glucose yield was determined based on quantified glucose released compared to total AIR cell
wall glucan available (as glucose), including non-cellulosic glucan.
Confocal microscopy
Air dried whole samples were sectioned by hand using a platinum tipped razor blade and
prepared for imaging by washing with 50 mL of de-ionized water prior to immersing in water
buffered at a pH of 7.0 by 20 mM Na2PO4 overnight. Pretreated samples were generated by
NaOH pretreatment at 0.10 g NaOH/g biomass of whole cross-sections, followed by washing
with de-ionized water until pH neutral, and immersion in water buffered at a pH of 7.0 by 20

51

mM Na2PO4 overnight. Confocal fluorescent images were collected on an Olympus FluoView
FV1000 Confocal Laser Scanning Microscope (Olympus America, Inc., Center Valley, PA),
configured on a fully automated Olympus IX81 inverted microscope. Images were recorded with
a 10x UPlanFLN objective (NA 0.30). Blue and red autofluorescence signals were sequentially
recorded, as shown Figure 2.10in Appendix A. Blue autofluorescence was excited using a 405
nm diode laser and emission was recorded using a 430-470 nm band pass emission filter. The red
autofluorescence was excited using a 543 nm helium neon laser and emission was recorded using
a 560 nm long pass emission filter. A series of confocal images (XYZ) were recorded through
the thickness of each sample (approximately 50 Âľm). Images were collected in 10 Âľm intervals
when using the 10x objective. Each confocal series was then compressed into a maximum
intensity projection image and recorded in a TIF format. The color look-up table (LUT) for all
images was modified with the blue display rescaled from 0-4095 to 350-800 and the red display
rescaled from 0-4095 to 0-1200 to brighten images. All further processing was performed in
ImageJ (http://imagej.nih.gov/ij/).
Glycome profiling
Sequential cell wall extractions and glycome profiling of switchgrass anatomical
fractions were carried out as described previously [39, 40] on AIR biomass. Plant glycandirected mAbs were from laboratory stocks (CCRC,JIM, and MAC series) available at the
Complex Carbohydrate Research Center [41]. A complete description of the mABs used in this
study can be found in our prior work [42].

Results
Switchgrass anatomical fractions sampled at different internodes throughout the plant
representing differing levels of maturity were utilized in this work to develop an improved
understanding of the impacts of within-plant heterogeneity in switchgrass on a potential
52

cellulosic biofuels conversion process. Switchgrass samples were characterized with respect to
composition and assessed for response to pretreatment and enzymatic hydrolysis with the goals
of (1) identifying cell wall properties contributing to observed differences in cell wall
recalcitrance, (2) understanding how differences in composition and recalcitrance are distributed
throughout the plant, and (3) potentially use this understanding to inform both plant breeding and
harvesting strategies to optimally couple the plant feedstock to a conversion process.
Anatomical fraction- and internode-specific recalcitrance
Enzymatic hydrolysis before or after mild NaOH pretreatment was performed on
extractives-free (AIR), de-starched cell wall material from three switchgrass anatomical fractions
(leaves, leaf sheath, stem) as a function of internode (Figure 2.2).

Figure 2.2 : Enzymatic hydrolysis glucose yields of untreated and alkaline pretreatment of
anatomical fractions. Enzymatic hydrolysis of untreated biomass was performed on AIR +
destarched biomass, while alkaline pretreatment was performed using 10% w/w NaOH loading
at 80°C for 1 hr. Glucose hydrolysis yields are presented as the percent of total cell wall glucan
released as monomeric glucose. Enzyme loading was 30 mg protein/g glucan for untreated
fractions and 10 mg protein/g glucan for alkaline pretreated fractions with a ratio of 2:1 CTec3 to
HTec3. Replicates (n = 3) are displayed as averages with standard deviations.
53

From the results in Figure 2.2, lower glucose yields for both untreated and NaOHpretreated cases of stem internodes indicate that stem fractions are significantly more recalcitrant
compared to leaf sheath (referred to as sheath) and leaf internodes. Previous work using liquid
hot water pretreatment of switchgrass also found that pretreated leaves were more digestible than
stem internodes [43]. Also of note, the mass of above-ground biomass in switchgrass is known to
be primarily leaves, with stems and leaf sheaths each comprising on the order of half the mass of
the leaves [43]. Stem internodes also demonstrated a clear decrease in enzymatic hydrolysis
yields with increasing tissue maturity (i.e., from I1 to I5) for both the untreated and NaOHpretreated cases [44], although similar trends were not as clearly observed in the untreated yields
of leaf sheath and leaf internodes and indicate cell wall recalcitrance increases with maturity in
stem only.
NaOH-pretreated leaf sheaths again showed significantly different trends compared to
stem, with a trend of increasing hydrolysis yields with increasing internode in both the 6-hr and
48-hr hydrolysis yield cases. Leaf hydrolysis yields became similar between internodes in the 48hr yields for the NaOH-pretreated case, contrasting the general trend of decreasing yields with
increasing internode seen in the untreated and 6-hr hydrolysis yield for the NaOH-pretreated
cases. From these observations, the application of alkaline pretreatment resulted in a change in
recalcitrance between internodes in both leaf sheath and leaf fractions, although differences in
the 6-hr and 48-hr hydrolysis yield trends in leaf internodes suggest pretreatment also influenced
hydrolysis rate as well. The contributions of composition and cell wall biopolymer structure to
these differences in recalcitrance are described in the subsequent sections.

54

Cell wall composition
The content of major cell wall biopolymers (cellulose, xylan, lignin), minor
hemicelluloses (non-cellulosic glucan) and minor hemicelluose sugars (Rha, Ara, Man, Gal,
Fuc), alkali-labile hydroxycinnamic acids (p-coumarate, ferulate), and lignin composition
(syringyl and guaiacyl monomer yields by thioacidolysis) were quantified for stem, sheath, and
leaf internodes. Key results that highlight observed trends found are presented in Figure 2.3,
while the complete data set can be found in Table 2.1 in Appendix A.

Figure 2.3 : Compositional highlights of (A) lignin (B) hydroxycinnamates (C) minor
polysaccharides, and (D) acetate and arabinan in switchgrass fractions with respect to
internode. Internodes I1-I5 are displayed in ascending order from left to right for each
switchgrass anatomical fraction. All composition values were quantified on an extractives free
basis (AIR).

55

The Klason lignin content demonstrated a clear increase with respect to internode for
stem, sheath, and leaf (Figure 2.3A), with results presumably corresponding to increasing
maturation and development of secondary cell wall tissues following cell elongation and
differentiation in older fractions. Overall, stem lignin content was higher compared to sheath and
leaf, which is expected due to the abundance of sclerenchyma and xylem cells providing
structural support for both upright growth and normal vascular transport [5]. It should be noted
that ash was not quantified independently in this study, and may represent a portion of the
quantified lignin [45]. However, ash content has been shown to be higher in abundance in leaf
fractions when compared to stem and sheath fractions [43], likely limiting ash impact on trends
observed in this study.
Alkali-labile hydroxycinnamic acids also varied significantly with internode, with both
pCA and FA content decreasing between internodes I1-I5 for all switchgrass anatomical fractions
(Figure 2.3B). Decreases in pCA content with increasing internode maturity ranged from a 19%
decrease from I1 to I5 in stem samples, to 53% and 45% in sheath and leaf samples, respectively.
FA content also demonstrated the same trend between internodes I1-I5 with decreases of 47%,
65%, and 48% for stem, sheath, and leaf, respectively. Combined with the trend for lignin
content (Figure 2.3A), the inverse relationship between FA content and lignin content suggest
that crosslinking of cell wall biopolymers via ferulates is an important feature of cell wall
organization in tissues with less secondary cell wall lignification [18]. Prior work has concluded
that total FA content decreases with increasing maturity (based on harvest date) concurrent with
increasing lignin content in switchgrass [45], and is in agreement with the results of the present
work in which maturity is based on internode rather than harvest date.

56

Composition analysis of polysaccharide content revealed a similar content of xylan
between internodes (average content of 24.4% for stem and sheath and 19.8% for leaf), while
galactan, mannan, rhamnan, and fucan, representing minor hemicellulose and structural pectin
polysaccharides were higher in sheath and leaf fractions compared to stem and showed clear
trends of increased abundance with tissue maturity in sheath and leaf internodes (Figure 2.3C).
Functionally, this indicates in maturing stem internodes, pectic and minor hemicellulosic
polysaccharides represent a smaller fraction of total cell wall polysaccharides, and the decreases
in total compositional fraction suggest minimal continued synthesis of these polysaccharides
during tissue maturation. Conversely, increased overall composition content in more mature
sheath and leaf internodes suggest continued synthesis and accumulation of minor
polysaccharides during maturation. The exception to this trend, non-cellulosic glucan, showed an
increase with respect to internode in all three anatomical fractions as well as similar overall
content of non-cellulosic glucan (2.5% overall for all samples), indicating increases in either βglucan or xyloglucan content with internode maturation (Table 2.1 in Appendix A).
Stem arabinan content, indicative of glucuronoarabinoxylan (GAX), arabinogalactan
(AG), and rhamnogalacturonan I (RG I) pectin side chains decreased slightly in stem internodes,
while increasing only in leaf sheaths (Figure 2.3D). Given the small decrease in overall xylan
content between internodes, it can be hypothesized that increases in arabinan content correspond
to an increase in GAX or RG I side chain abundance in leaf sheath cell walls [23]. Many cell
wall polysaccharides are known to be O-acetylated, including pectins, xyloglucans, and xylans.
Cell wall acetyl content (Figure 2.3D) decreased significantly with stem internode maturity, with
only minor decreases observed with respect to sheath and leaf internode maturity, indicating

57

similarity in acetylated cell wall components between sheath and leaf internodes compared to
stem.
Non-cellulosic glucose and extractable sugar content
The accumulation of soluble sugars in the stems of grasses is of paramount importance to
agriculturally important crops such as sugarcane (Saccharum spp.) and sweet sorghum (Sorghum
bicolor). Soluble sugar abundance in other grasses used as feedstocks for cellulosic biofuels can
be a significant fraction of total dry weight [50], with both genetic and environmental drivers
responsible for starch and extractable sugar accumulation [46]. For example, soluble sugar
synthesis in plants has been shown to be environment responsive, with significant accumulation
of soluble sugars accompanying reduced structural carbohydrate and lignin content in
switchgrass harvests subjected to drought conditions [47].
Notably these sugars are often overlooked in many biomass-to-fuels studies or analyses
and can present both processing and analytical challenges. Soluble sugars may mask structural
polysaccharides in cell walls [48] resulting in false positives for carbohydrate content in
screening techniques relating feedstock digestibility to biomass physical characteristics or high
throughput digestibility studies that do not remove soluble sugars and starches before analysis
[49]. As a processing challenge, contents of extractable sugars and starch can be degraded during
many pretreatments (e.g., dilute acid or AFEX) representing a potentially significant loss of
value sugars. Furthermore, sugar degradation products can present additional challenges to the
biological conversion of sugars to biofuels or bioproducts. For example, in recent work using
AFEX pretreatment on switchgrass subjected to drought environmental stress, imidazoles and
pyrazines were formed by the reaction of ammonia with soluble sugars in high enough

58

concentrations to significantly inhibit fermentation compared to switchgrass harvests not
subjected to drought [47].

Figure 2.4 : Native switchgrass non-cellulosic sugar composition with respect to internode.
Internodes I1-I5 are displayed in ascending order from left to right for each switchgrass
anatomical fraction. Compositions are represented as a percent of total dry weight. Replicates (n
= 3) displayed as average with standard deviations.
In the present work, non-cellulosic glucan (presumably as xyloglucan and β-glucan) and
extractable such as sucrose, fructose, free glucose, and starches sugar contents were determined
(Figure 2.4) and represent a non-trivial fraction of overall feedstock mass. Stem internodes were
observed to have significantly more soluble sugars compared to similar sheath and leaf internode
contents, with soluble sugars combined with non-cellulosic glucan constituting between 12% to
18% of total dry weight. Starch content was dramatically different in stem internodes,
accumulating to up to 8% of total biomass dry weight in internode I5. Comparatively, sheath and
leaf starch content accounted for up to 2% dry weight, and decreased with internode maturity.
One likely reason for such a high stem extractible sugar content was due to sample harvesting
during grain filling [50].
59

Non-cellulosic glucan, presumably in the form of xyloglucans, β-glucans, and
glucomannans, were observed to accumulate from internode I1 through I5 for all three anatomical
fractions. Although comprising a small percentage of total biomass, β-glucans are generally more
easily digested compared to cellulose, while xyloglucans may be closely associated to cellulose
in the primary cell wall [51]. It can also be observed that the differences in the non-cellulosic
glucan can be correlated to differences in most of the minor hemicellulose and pectin sugars (i.e.,
Fuc, Rha, Man, Gal as presented in
Figure 2.3C).
Cell wall autofluorescence
At both the macroscopic and microscopic scales as seen in Figure 2.5, switchgrass stem,
sheath, and leaf exhibit obvious significant differences in overall morphology, tissue
organization, and cell type abundance. Structural (and compositional) heterogeneity across
fractions has important implications for biomass recalcitrance. As one example, sclerenchyma
secondary cell wall types in stems have significantly thicker cell walls and contain a high ratio of
lignified secondary cell walls compared to partially lignified parenchyma type cells [45],
resulting in anatomical fractions containing high contents of sclerenchyma cells expecting to
have higher recalcitrance compared to parenchyma rich fractions.

60

Figure 2.5 : Examples of differences in macroscopic morphology, cell wall organization,
and aromatic abundance as shown by autofluorescence between switchgrass anatomical
fractions from internode I2. Confocal laser scanning microscopy carried out using laser
emission wavelengths of 405nm and 543 nm. Tissue types are denoted from the following
notation: Ep – epidermis, Fb – fiber bundles, Vb – vascular bundles, Ph – phloem, X – xylem, Pc
– Parenchyma cells. Scale bars represent 200 µm.
Analysis of cell wall phenolics using confocal microscopy often employs combinations
of fluorescence tagging, chemical labels, and autofluorescence techniques to characterize the cell
wall landscape at the micron-scale resolution [52, 53]. Using phenolic autofluorescence alone,
general trends in phenolic localization and abundance can be obtained, however the wide range
of emission spectrums of phenolic compounds can limit deconvolution of specific phenolic
components [54-56]. With a low-field excitation using a diode laser at 405 nm,
hydroxycinnimates have an emission spectrum of around 420 – 460 nm (shown and denoted as
blue) along with alkaloids, flavonoids, and other phenyl propanoids [56]. Using a helium neon
laser excitation at 543 nm, most phenolics fluoresce with an emission spectrum of 600-650 nm
(shown and denoted as red), however hydroxycinnimates do not [55]. One convoluting factor in
this technique resides in the autofluorescence of chlorophyll. With an emission spectrum in the

61

600-700 nm region [56], chlorophyll will certainly provide signal contribution to the red
emission spectrum from chlorophyll rich tissues such as epidermal and bundle sheath regions of
leaf (Figure 2.5 and Figure 2.6).
Overlay of both autofluorescence channels of whole biomass cross sections revealed
several noticeable features of each organ (Figure 2.5). Stem and sheath cross-sections displayed
similar architecture, with atactostele patterned vascular bundles, distinct regions of fiber bundles,
and similar overall thickness [57]. The only notable difference resided in distribution of fiber
cells between epidermal located vascular bundles in the sheath cross-section. Both displayed
blue and red emission patterns in all tissue types, however signal intensity was much stronger in
secondary cell wall abundant regions such as sclerenchyma cell fiber bundles, and vascular
bundles. Leaf cross-section architecture was significantly different, with vascular bundles
surrounded by a bundle sheath of parenchyma cells, and a highly lignified lower midrib region.
Leaf autofluorescence showed a blue emission pattern in the vascular bundles along with some
epidermal tissues, while the red emission spectrum was localized to the lower midrib and bundle
sheath, and was most likely due to chlorophyll autofluorescence.
The impact of NaOH pretreatment on autofluorescence signature was examined for
multiple internodes to gauge both organ and internode response to pretreatment (Figure 2.6). In
stem and leaf sheath cross-sections, the red emission spectrum was localized to secondary cell
wall tissue such as vascular tissue and fiber bundles, while the blue emission spectrum was
observed predominantly in parenchyma cells.

62

Figure 2.6 : Cell wall autofluorescence cross-sections of NaOH-pretreated switchgrass
anatomical fractions from internodes I2, I3, and I4. Confocal laser scanning microscopy
carried out using an overlay of laser excitation wavelengths of 405 nm and 543 nm. Scale bars
represent 200 Âľm.

Comparing cross-sections of stem internodes I2 to I4, there was a clear increase of the
blue emission spectrum in the fiber bundles, while in leaf sheaths cross-sections sampled from
internodes I2 to I4, there was a clear decrease of the blue emission spectrum in the parenchyma
cells. Signal intensity from the red emission spectrum decreased in fiber and vascular cells in
both stem and leaf sheaths from internode I2 to I4. In addition to changes in signal intensity, cell
wall thickening was also observed in the parenchyma and interfascicular tissues of stem

63

internode cross-sections, corresponding to a maturation of tissues [18]. Similar thickening was
not as apparent in sheath or leaf cross-sections.
These results suggest that following NaOH pretreatment, components that fluoresced red
in the stem cross-sections were either preferentially extracted due to pretreatment, or were
observed in lower abundance relative to other components in more mature internodes. This may
also suggest that following alkaline pretreatment, there were regions of hydroxycinnimates that
remained unextracted or were more recalcitrant in stem internodes, as evidenced by the change
in ratio of blue emission to red emission spectrum. The loss of both emission spectrum signals
with internode maturity in sheath cross-sections suggests either the removal of both components
with alkaline pretreatment, or the loss of accessibility of both components for autofluorescence.
From the perspective of secondary cell wall development, more mature internode crosssections may have hydroxycinnimates in all cell types, while in younger stem internode crosssections, potential locations of hydroxycinnimates might be restricted predominantly to
developing parenchyma cells. In sheath cross-sections, younger internodes had potential
locations of hydroxycinnimates in all cell types, with the eventual observance of the blue
emission spectrum only in the vascular bundle regions. Leaf internodes following NaOH
pretreatment had a noticeable decrease in both emission signal intensities, with the blue emission
spectrum localized to the outer parenchyma and epidermal cells, and the red emission spectrum
localized to the bundle sheath and epidermal cells. Given the autofluorescence of chlorophyll in
the red emission spectrum, the decrease in red signal can be largely attributed to either
chlorophyll removal from alkaline extraction, or a decrease in chlorophyll in more mature leaf
internodes following senescence.

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Glycome profiling
Glycome profiling was performed to assess organ-specific variations in the cell wall
glycan composition and extractability across individual internodes of each anatomical fraction,
I1-stem, I1-sheath, and I1-leaf. In Figure 2.7A, selected regions of glycome profiles highlight
abundancies of xylan and pectin epitopes in cell wall extracts, while the full glycome profile
results can be found in Figure 2.11 in Appendix A.

Figure 2.7 : Abundance of select major non-cellulosic glycan epitopes in chemical extracts
from stem, sheath, and leaf anatomical fractions of internodes I1 (A), and extracts from
stem internodes I1-I5 (B). Extracts were screened by ELISA using a comprehensive suite of cell
wall glycan-directed mAbs as described in materials and methods. Binding response values are
depicted as heat maps with black-red-bright yellow color scheme representing from no binding
to strongest binding. The amount of carbohydrate material recovered per gram of AIR is depicted
in the bar graphs (green) above the heat maps. The panel on the right-hand side of the heat map
shows the groups of mAbs based on the class of cell wall glycan they each recognize.

65

With the oxalate extracts, extractability of xylan epitopes varied across anatomical
fractions. For instance, both I1-stem and I1-sheath exhibited similar xylan epitope extractability
patterns with significant abundance of arabinosylated xylan epitopes recognized by mAb CCRCM154 of xylan-4 clade [58], unsubstituted xylan epitopes recognized by several mAbs from
xylan-6 clade, and unsubstituted xylan epitopes recognized by all mAbs from xylan-7 clades.
The I1-leaf oxalate extract was significantly different, showing an abundance of only
arabinosylated xylan epitopes recognized by mAb CCRC-M154 of xylan-4 clade along with
significantly reduced abundance of unsubstituted xylan epitopes recognized by mAbs from
xylan-7 clades. Differences across anatomical fractions were also observed in the carbonate
extract, with all three carbonate extracts containing most classes of xylan epitopes from
unsubstituted and substituted regions. The I1-leaf sample again exhibited differences in the
pattern of extractability with a marginal reduction in the overall abundancies of these epitopes,
namely the Me-GlcA substituted xylan epitopes (xylan-5) and complete absence of xylan-6
epitopes. Xylan-3 epitopes were observed only in the I1-sheath, while all xylan clades were
observed in the I1-stem extract.
Likely due to grass susceptibility to alkaline pretreatment [38], the 1M KOH extract
showed the largest portion of total carbohydrates released, with significant xylan epitope
abundance in all three sample extracts. I1-leaf exhibited the strongest binding intensity for xylan
epitopes 4 through 7 in the 1M KOH extract, and this trend was mirrored in the 4M KOH
fraction. Similar patterns of xylan epitopes composition and extractability were recognized in the
4M KOH extract, however at this point binding intensities across samples were largely similar
and were probably the residual alkaline susceptible xylans that remained after the 1M KOH
extract [42]. The chlorite and subsequent 4M KOH PC extracts were used to determine

66

carbohydrates more tightly bound with lignin. Interestingly, the chlorite extract showed overall
similar epitope recognition patterns for all three extracts as the oxalate extract, with the I1-leaf
containing only xylan-4 and xylan-7 epitopes. Subsequent 4M KOH PC extraction showed
similar overall epitope extractability patterns and binding intensity as the 1M KOH extract, with
I1-sheath and I1-leaf extracts having the most intense binding for xylan-4 to xylan-7 epitopes.
From these results, it can be hypothesized that xylan integration within the cell walls of
each anatomical fraction are not equivalent. Specifically, xylan association in the leaf fraction
appeared to be more tightly integrated within the cell wall, as leaf xylan required a more severe
basic 1M KOH extract to observe significant xylan epitopes. Recognition of xylan-5 epitopes in
the carbonate and 1M KOH fraction was unexpected, as more substituted xylans such as
arabinoxylan are thought to exist in primary cell walls, while less substituted and branched
xylans are associated with the secondary cell wall [59]. This result may indicate an abundance in
arabinosylation of leaf xylans compared to stem xylan in these switchgrass fractions [60].
Furthermore, higher xylan epitope recognition in the 4M KOH PC suggests that xylan is more
integrated with lignin in the leaf fraction, which may be a result of hemicelluloses in the leaf
contributing more towards leaf internode recalcitrance compared to higher lignin containing
fractions such as stem and sheath.
In terms of pectin epitope trends, while I1-stem and I1-sheath displayed marginal pectin
epitope abundancies in the oxalate extract, the majority of recognized epitopes and highest
abundancies was in the I1-leaf extract. Specifically, epitopes of the homogalacturonan backbone
(HG backbone 1 & 2) and pectic arabinogalactan epitopes (RG-I/AG) were broadly recognized
in the I1-leaf extract, with weak to moderate binding also occurring in the I1-sheath extract. More
intense binding and recognition for the pectic arabinogalactan epitopes was observed in the

67

carbonate extract for I1-sheath and I1-leaf, with a moderate amount of recognition in the I1-stem
extract. In addition, HG 1 epitopes were observed in high intensity for all three extracts, and
rhamnogalacturanan-I backbone (RG-I backbone) was recognized highest in the I1-sheath
extract. Overall, a higher abundance of pectin backbone and pectic arabinogalactan (AG)
epitopes was noted in I1-leaf samples in 1M, 4M KOH and 4M KOH PC extracts, while patterns
in chlorite extracts were largely similar across samples.
From the pectin epitope recognition patterns, it is clear the stem anatomical fraction
generally had lower overall pectin content that was also less recalcitrant. Sheath and leaf
anatomical fractions displayed similar epitope recognition, suggesting similar pectin structures,
however, differential epitope binding intensity suggests that each anatomical fraction has a
unique organization of pectin in the cell wall. Pectic AGs are the major galactose containing
pectin found in grasses [61], with the higher content of galactan found in the cell walls of leaf
extracts suggesting higher overall pectic AGs content compared to stem samples [23].
Glycome profiling for internode specific extractability
Stem internodes I1- I5 were subjected to glycome profiling to examine trends in the noncellulosic polysaccharide extractability and structure with respect to tissue maturity. As earlier,
selected epitope profiles for xylan and pectins are shown in Figure 2.7B, while the complete
glycome profile can be found in Figure 2.12 in Appendix A.
Oxalate and carbonate extracts showed binding of xylan-4 and xylan-7 epitopes in all
internodes, with the significantly enhanced appearance of xylan-6 epitopes in the carbonate
fraction. There was a trend of reduced xylan-5 epitope abundance in the more mature internodes,
corresponding to a decrease in extractable Me-GlcA substituted xylans in oxalate, carbonate, and
1M KOH extracts. The 1M KOH extract showed significant epitope extractability of xylan-4 and

68

xylan-7, indicating that a large fraction of the total polysaccharides released in the 1M KOH
extracts were unsubstituted xylans or arabinoxylated xylans. The 4M KOH fraction showed
similar abundance of xylan-4, xylan-5, xylan-6, and xylan-7 epitopes among all developmental
stages of internodes, while the 4M KOH PC extracts showed an increasing trend of xylan mAb
extractability in more mature internodes.
Taken together, these results indicate that internode maturity impacts xylan
deconstruction and extractability. Specifically, higher abundance of xylans were recognized in
less severe chemical extracts for younger internodes, while harsh extractions after chlorite
delignification showed the opposite trend, indicating lignification plays a greater role in xylan
recalcitrance in more mature internodes [62]. Pectin epitope abundance was largely observed in
the carbonate and 4M KOH PC extracts, with an increase in binding intensity of pectic AG
epitopes with maturing internodes. In addition, a clear trend in decreasing binding strength was
observed with the epitopes for the HG backbone 1, which is an indication of a transition from an
abundance of primary cell walls to the development of secondary cell wall thickening [63]. In the
4M KOH PC extracts, a similar trend was observed for the pectic AG as in the carbonate
fraction, however, the HG backbone 1 epitope abundance increased with internode maturity.
Pectin trends indicate that less severe extracts result in more extractable pectins from more
mature internodes, which would indicate that pectins more closely associate within the cell wall
in less mature internodes or are more easily liberated compared to xylans. At the same time,
however, the trend observed post chlorite delignification suggests that there is a second fraction
of pectins that are closely associated with lignin, as indicated by the increased pectin and pectin
backbone abundance in more mature internodes. Pectin-lignin associations have been proposed
in grasses in prior studies [23, 64].

69

Discussion
Results for plant cell wall composition together with cell wall susceptibility to
deconstruction by a coupled alkaline pretreatment and enzymatic hydrolysis demonstrate
different switchgrass anatomical fractions possess significant differences both within fractions of
varying maturity, and between different anatomical fractions. These trends are shown as a
summary of Pearson’s correlation coefficients between cell wall properties and hydrolysis yields
for either individual anatomical fractions or for all pooled fractions are presented in Figure 2.8,
while data from a few illustrative identified correlations are plotted in Figure 2.9.

Figure 2.8 : Summary of Pearson’s correlation coefficients between plant cell wall
properties and glucose hydrolysis yields for each anatomical fraction and for pooled
samples. Data demonstrate strong correlations between many properties and yields.

First, lignin content was found to exhibit a negative correlation with untreated and
pretreated in stems and leaves (Figure 2.8), while demonstrating a strong, significant negative
correlation when all anatomical fractions are pooled for 48 hr hydrolysis yields following
pretreatment (R = -0.68; p-value = 0.005; Figure 2.8 and Figure 2.9A). As similar trends are
observed for untreated hydrolysis yields and pretreated hydrolysis yields, indicating the trend is

70

related to anatomical fraction structure and maturation rather than pretreatment response. Stems
contain the highest content of lignin of any of the anatomical fractions (
Figure 2.3A), with tissue maturity-dependent lignification and cell wall thickening [18]
observed in confocal microscopy cross-sections (Figure 2.6). These results together suggest
lignin is a defining feature of recalcitrance in the cell walls of developing stems. Notably, lignin
is a well-known contributor to cell wall recalcitrance, and as one example, our previous work in
diverse maize lines demonstrated that initial lignin content was a strong predictor of glucose
hydrolysis yields for no pretreatment, while lignin content after mild NaOH pretreatment was a
strong predictor of glucose hydrolysis yields comparable to what was performed in the present
work [38]. Glycome profiling results do suggest that hemicelluloses may play a parallel role in
inhibiting cell wall deconstruction, as a noted decrease in xylan epitope extractability was
observed in maturing stem internodes until alkaline extraction post chlorite delignification
(Figure 2.7), however higher lignification reducing xylan accessibility to extraction may have
contributed to the observed trends.

71

Figure 2.9 : Select correlations between data relationships between either lignin content or
ferulic acid against glucose hydrolysis yield represented as scatter plots. (A) 48-hr glucose
hydrolysis yields for NaOH-pretreat biomass versus lignin content, (B) ferulate content in
relation to total lignin, and 48-hr glucose hydrolysis yield with respect to (C) fucan and (D)
arabinan content.
Variability in lignin composition may also play a significant role in anatomical fractionspecific differences of recalcitrance. A strong negative correlation was observed between lignin
and ferulate content for all fractions, shown in Figure 2.9B (R2 = 0.923; p-value = 10-8). Coupled
with the observed decrease in ferulate content with increasing internodes shown in
Figure 2.3B and as reported previously in switchgrass stem internodes [18], suggesting
that ferulate content within the cell wall is dependent upon both tissue maturity and lignin
content. Ester crosslinking of cell wall biopolymers via ferulates is considered to be one of the
major contributing features cell wall recalcitrance in low-lignin grass cell walls [7, 65]. Notably,
ester crosslinks are susceptible to saponification during mild alkaline pretreatment relative to

72

ether linkages between other monolignols, and may potentially contribute significantly to
enzymatic hydrolysis yields following alkaline pretreatment.
Similar to stem internode composition, lignification increased in maturing leaf sheath cell
walls, however, the positive correlation between 48-hr glucose hydrolysis yields following
NaOH pretreatment and lignin content in leaf sheath samples (Figure 2.9A) suggests that other
cell wall components, such as hemicellulose substitution, pectin abundance, and polysaccharidelignin associations may be a more significant contributor to cell wall recalcitrance. In the leaf
sheath, significant positive correlations were observed between minor polysaccharide sugars
(Rha, Ara, Fuc, Gal, Man) and hydrolysis yields following pretreatment (Figure 2.8; Figure
2.9C and Figure 2.9D), and considering the leaf sheath cell wall composition of each of these
minor sugars can be more than double that in stem internodes (
Figure 2.3C), indicating these polymers may play a more important role in cell wall
recalcitrance. Of note, arabinan, which is present as substitutions on the xylan backbone in GAX
and to a substantially lesser extent as arabinogalactan (AG), is substantially higher in leaf sheath
cell walls than in stem internodes and exhibits a strong positive correlation to glucose hydrolysis
yields in pretreated biomass (Figure 2.9D). The implications of this are that increased
substitution of xylan with arabinosyl substitutions suggests a less tightly crosslinked cell wall
matrix that may be more accessible to enzymatic deconstruction [21,66]. This finding is in
agreement with the work of Costa et al [62], who found that the Ara:Xyl ratio in different
anatomical fractions isolated from sugarcane stems was a strong predictor of enzymatic
hydrolysis yields in un-pretreated biomass.
Although possessing similar arabinan content, the same trend was not observed in leaf
internodes; lower cellulose, xylan, and lignin content in leaf internodes suggest that due to

73

limited lignification in leaf internodes compared to stem internodes (Figure 2.2), structural
pectins and minor hemicellulose components with lower overall compositions playing a more
significant role in cell wall recalcitrance. Results suggest that a relatively internode-independent,
uniform response was observed in leaf cross-section phenolic autofluorescence to alkaline
pretreatment (Figure 2.6), corresponding to similar response to lignin removal. The effectiveness
of enzymatic hydrolysis of leaf internodes following NaOH pretreatment further suggests that
alkaline susceptible cell wall components did not have a differential effect upon leaf cell wall
digestibility. Although displaying comparable yields to alkaline pretreated leaf sheaths, leaf cell
walls may be more susceptible to a pretreatment that predominantly removes more
polysaccharide components, such as liquid hot water pretreatment, which has shown to
dramatically improve leaf digestibility [67].
Technologies for biomass fractionation or extraction employed either on-field or at a
centralized processing facility offer the potential to simultaneously improve agronomic, logistics,
and processing outcomes. As an example, on-site field fractionation of switchgrass could
capitalize on more recalcitrant stem internodes residing at the bottom of the tiller, harvesting
upper less mature internodes while leaving mature internodes on-field for soil coverage and
nutrient retention. Given stem and sheath leaf fractions constitute half of leaf dry weight in prior
reporting [43], loss of biomass yield would have to be weighed against increases in downstream
sugar yields due to less recalcitrant materials representing a larger total fraction of biomass.
Additionally, grass stems can accumulate a substantial amount of soluble sugars in
storage parenchyma cells surrounding the vascular tissues [68]. Given that soluble sugars can
represent more than 10% of the switchgrass total dry weight in this study, water extraction may
be proposed as a route to yield a stream rich in fermentable sugars. Sugarcane undergoes

74

mechanical fractionation coupled with counter-current hot water extraction to remove soluble,
fermentable sugars. Flow-through liquid hot water pretreatments are well-established
pretreatment technologies [69]; counter-current processing configurations can be envisioned that
integrate soluble sugar extraction with liquid hot water pretreatment, each taking place at their
respective optimal temperatures.

Conclusions
In this work, we were able to demonstrate that significant heterogeneity in cell wall
structure, composition, properties within switchgrass results in substantially different feedstocks
response to processing in a cellulosic biorefinery. Feedstock heterogeneity may offer the
potential to either tailor harvest approach, harvest time, or employ physical fractionation during
processing to optimize processing outcomes such as sugar yield. Using switchgrass physically
fractionated by anatomical fraction and internode, we identified compositional differences that
impact recalcitrance and related results to macroscopic observations. Specifically, we showed
that while all anatomical fractions experienced different extents of lignification between
internodes; stem internode, leaf sheath, and leaf blade fractions were shown to have different
structural features that dominate cell wall recalcitrance. Lignin content was identified as
exhibiting a strong correlation to cell wall recalcitrance in stem internodes and initial
recalcitrance in leaves. Leaf sheath and leaf cell wall recalcitrance were also shown to be
impacted significantly by hemicellulose content and substitution, as well as structural pectin
content. Ferulate and p-coumarate content were inversely correlated to lignin content in all
fractions and decreased significantly with increasing internode maturity. Additionally, soluble
sugars can account for a non-trivial fraction of total switchgrass dry weight, and represent a
potential avenue to generate an enriched stream of easily extracted sugars to improve process
economics.

75

Acknowledgements
The authors would like to thank Dr. Melinda Frame (Michigan State University) for assistance
with confocal microscopy sample preparation and image collection.

76

APPENDIX

77

APPENDIX A: Supplementary figures

Table 2.1 : Compiled AIR + destarched cell wall polysaccharides, acetate, and lignin
content for five internodes of three switchgrass anatomical fractions. Samples are presented
as an average (n = 3) with standard deviations following in Table 2.2.

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Table 2.2 : Standard deviations for compiled AIR + destarched cell wall polysaccharides,
acetate, and lignin content for five internodes of three switchgrass anatomical fractions.
Samples are presented as an average (n = 3) with standard deviations following.

79

Figure 2.10 : Cell wall autofluorescence of switchgrass organ cross-sections from internode
I2. Confocal laser scanning microscopy carried out using laser excitation wavelengths of 405 nm
and 543 nm. Tissue types are denoted from the following notation: Ep – epidermis, Fb – fiber
bundles, Vb – vascular bundles, Ph – phloem, X – xylem, Pc – Parenchyma cells. Scale bars
represent 200 Âľm.

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Figure 2.11 : Heat map abundance of major non-cellulosic glycan epitopes in oxalate,
carbonate, 1M KOH, 4M KOH, chlorite, and 4M KOH PC extracts from cell walls of stem,
sheath, and leaf samples of internodes I1. Extracts were screened by ELISA using a
comprehensive suite of cell wall glycan-directed mAbs as described in materials and methods.
Binding response values are depicted as heat maps with black-red-bright yellow color scheme
representing from no binding to strongest binding. The amount of carbohydrate material
recovered per gram of AIR is depicted in the bar graphs (purple) above the heat maps. The panel
on the right-hand side of the heat map shows the groups of mAbs based on the class of cell wall
glycan they each recognize.

81

Figure 2.12 : Heat map abundance of major non-cellulosic glycan epitopes in oxalate,
carbonate, 1M KOH, 4M KOH, chlorite, and 4M KOHPC extracts from stem internodes
I1-I5. Extracts were screened by ELISA using a comprehensive suite of cell wall glycan-directed
mAbs as described in materials and methods. Binding response values are depicted as heat maps
with black-red-bright yellow color scheme representing from no binding to strongest binding.
The amount of carbohydrate material recovered per gram of AIR is depicted in the bar graphs
(purple) above the heat maps. The panel on the right-hand side of the heat map shows the groups
of mAbs based on the class of cell wall glycan they each recognize.

82

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Chapter 3 : Relating nanoscale accessibility within plant cell walls to
improved enzyme hydrolysis yields in corn stover subjected to
diverse pretreatments
This work has been submitted as original research in July 2017 to the Journal of Agricultural and
Food Chemistry

Introduction
Lignocellulosic biomass is a promising feedstock for the production of renewable fuels and
chemicals. One key challenge limiting effective utilization of biomass resides in plant-evolved
cell wall recalcitrance [1], or resistance to degradation imparted by the higher order structure of
plant cell walls. The physical structure of the plant cell wall matrix can be considered a
nanostructured composite material, with networks of cellulose fibril bundles and matrix
polymers forming cell wall layers [2]. Networks of microfibrils are associated and potentially
cross-linked with amorphous hemicellulose polysaccharides through non-covalent interactions
[3]. In lignified secondary cell walls, this network can contain lignin, which is deposited to
impart structural rigidity, reduce water and solute permeability, and limit extracellular
interactions [4]. Thermochemical pretreatment can be applied to improve the deconstruction of
plant cell wall polysaccharides by cellulolytic enzymes [5], with the goals of pretreatment to
reduce recalcitrance by removing or redistributing hemicellulose and lignin, disrupting the
crystalline structure of cellulose, or disrupting the cell wall ultrastructure. Pretreatments often
have the net effect of increasing cellulose accessibility to cellulolytic enzymes [6,7].
While cellulose accessibility can be considered as the ability for an enzyme to bind to
cellulose and hydrolyze a glycosidic bond, this property is often indirectly assessed as accessible
surface area defined by a combination of particle size, porosity, and pore volume [8]. Porosity in
90

the context of the plant cell wall matrix can be considered to be due to a combination of
composition and structural features impacting the rigidity and internal surface area of the
composite polymer network. Lignin is a hydrophobic component when sufficiently removed
from the cell wall has been directly associated with loss of cell wall rigidity [9]. Xylan removal
is also associated with increases in cell wall porosity, but is also coupled to a certain level of
lignin removal depending on pretreatment [10]. The removal or relocalization of cell wall
components alters the structural integrity of the cell wall matrix, allowing swelling and
contraction in response to the solvent, exhibiting behavior like a hydrogel. It is with this model in
mind, that porosity is a dynamic term that defines both structural features, as well as the cell wall
response to environmental conditions.
Techniques to measures properties of porosity include metrics to measure bulk properties of
the cell wall such as water-cell wall associations using water retention value (WRV) [11],
measuring cell wall surface adsorption with dyes of known hydrodynamic radii to determine
accessible substrate area using the Simons’ Stain technique [12] or of fluorescent-labeled
enzymes for direct enzyme accessibility measurements [13]. Water constraint measured using T2
NMR relaxometry provides information on the extent of water-cell wall hydrogen bonding, with
increased water constraint associated with increased water accessibility within the cell wall [14].
Pore volume is a specific characteristic of porosity, defined by the porous nature of the cell wall
in an idealized slit model. In the idealized slit model, the cell wall thought of as a series of
variably spaced lamellae that contain distinct pore size, and therefore pore volume characteristics
[15]. The effect of cell wall component removal or re-localization during pretreatment can be
associated to changes in the cell wall ultrastructure inducing 3D topographical changes in the
form of induction of pore formation or voids [16]. Techniques to measure cell wall pore

91

distributions quantify the interaction of water within the pores to determine pore distributions.
Solute exclusion utilizes the availability of porous water to influence diffusion of a probe of
known hydrodynamic radii into a pore of equal or larger diameter [15], however is dependent
upon uniform pore sizes for accurate pore volume determination [17]. Differential scanning
calorimetry-thermoporometry quantifies heat endotherms associated with freezing point
depression of water closely associated with an interface [18], while 1H NMR cryoporosimetry
quantifies water content through similar thermochemical assumptions [19], however requires
assumed pore geometries to extract meaningful pore distributions.
The relationship between pore volume and enzymatic hydrolysis has been investigated in
multiple studies. Originally observed as differences in the initial rates of glucose hydrolysis for
acid-pretreated hardwoods and softwoods [20], a positive correlation between pore volume and
enzymatic hydrolysis has also been determined in other pretreatment and feedstock systems such
as chlorite-delignified sugarcane bagasse [21]. Other studies have not observed similar trends,
with dilute acid pretreatment of corn stover showing no discernable relationship between
pretreatment severity, hydrolysis yield, and pore volume [19]. Different methods of measuring
pore properties such as Simons’ Stain coupled with differential scanning calorimetrythermoporometry have been recently used to evaluate alkali, autohydrolysis, and multi-stage
pretreatments of wheat straw, concluding that delignification increases pore accessibility, while
autohydrolysis increased cellulose accessibility at the expense of restricted pore accessibility
[22]. Our prior work has also demonstrated relationships between water sorption, water retention
and settling volume as predictors for enzyme binding and enzymatic hydrolysis using alkaline
hydrogen peroxide and liquid hot water pretreatment of corn stover and switchgrass [11].

92

The goals of this study are to investigate corn stover subjected to a range of pretreatment
conditions that include alkaline hydrogen peroxide (AHP) delignification and liquid hot water
(LHW) pretreatment. These pretreatment-modified feedstocks are used to assess the impact of
pretreatment on cell wall structural properties and to highlight key relationships between cell
wall properties and enzymatic hydrolysis. The role of nanoscale porosity as a key structural
property is investigated in the context of both solute exclusion measured accessible pore
volumes, WRV determined accessible water-cell wall surface area, and solute-induced cell wall
swelling. Correlations derived from both these chemical and physical changes in the cell wall are
developed and these properties are then related to enzyme accessibility and enzymatic hydrolysis
of cell wall polysaccharides.

Materials and methods
Biomass and composition analysis
Corn stover (Zea mays L. Pioneer hybrid 36H56) harvested in 2012 was milled with a Wiley
Mini-mill (Thomas Scientific) to pass a 5 mm screen, and air-dried to a moisture content of
approximately 5% prior to any pretreatments. Milled material was sieved, and a particle size
distribution between 425 micron and 1 mm was selected for use in all pretreatments and
subsequent experiments. Cell wall lignocellulosic material was isolated following extraction and
de-starching procedure outlined [23] three sequential washes of 70% ethanol, 1:1 methanolchloroform, and acetone to obtain alcohol insoluble residue (AIR). AIR was de-starched using 50
Âľg Amylase /mL H2O (Bacillus sp., Catalog # A7595, Sigma-Aldrich St. Louis, MO) and 18.7
units of Pullulanase (Bacillus acidopullulyticus, Catalog # P9286 Sigma-Aldrich, St. Louis, MO)
in a 0.01% sodium azide solution, with rotary mixing (Intelli-Mixer, ELMI-tech, Newbury Park,
CA) at 37°C overnight. The non-cellulosic neutral monosaccharide content of the wall matrix
polysaccharides was determined following trifluoracetic acid (TFA) hydrolysis of the de93

starched AIR followed by derivatization of the monosaccharides as alditol acetate method by
GC/MS according to Foster et al [23]. Lignin content was determined on AIR biomass using the
NREL/TP 510-42618 protocol [24] with minor modifications as described in Li et al [9].
Composition analysis was performed in technical triplicate.
Pretreatments
AHP pretreatment was performed using four different H2O2 to biomass loadings, 0, 0.06,
0.12, and 0.25 g H2O2/g biomass, at an insoluble solid loading of 15% (mass biomass per total
liquid volume). Samples were prepared in 250-mL Erlenmeyer flasks, sealed with parafilm to
allow expansion due to O2 evolution, and placed in an incubator at 30°C with shaking at 180 rpm
for 24 hr. The pH was maintained at 11.5 during pretreatment, with adjustments at 3 and 6 hr
using 3 M NaOH [11]. LHW pretreatments were performed in a 5 L M/K Systems digester (M/K
Systems, Inc., Peabody, MA, USA). Using 100-mL pressure tubes (Ace Glass, Sigma-Aldrich,
St. Louis, MO), 6.0 g of biomass was loaded with 70 mL of de-ionized water, yielding a solids
content of 8.5% (mass biomass per total liquid volume). Pretreatment conditions were specified
at either 160°C for 30 minutes or 190°C for 5 minutes, with a heat-up rate of 1.0°C/min and a
cool-down rate of approximately 1.0°C/min. All biomass following pretreatment was washed
until clear, vacuum filtered with a 200 mesh Buchner filter to a moisture content of
approximately 85% measured gravimetrically, and stored at 4°C. AHP pretreatment on LHWpretreated material for two-stage pretreatments was performed under identical conditions as
described previously, using thoroughly washed LHW pretreated material dried to a moisture
content of about 5% and sieved to ensure similar particle sizes as the initial AIR material.
Material drying for hornification measurements was performed in a temperature-controlled oven
(Isotemp, Fisher Scientific) at 105°C.

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Enzymatic hydrolysis
Enzymatic hydrolysis was performed on never-dried pretreated material in 250-mL
Erlenmeyer flasks using 5% (insoluble mass biomass per total liquid volume) dry solids loading
with the moisture content of the never-dried material determined separately to achieve the target
solids loading. Reaction media was buffered pH 5.2 using 50 mM sodium citrate buffer, with pH
adjustments occurring after 6-hr and 24-hr hydrolysis times. Tetracycline and cycloheximide at a
total concentration of 40 Âľg/mL each were used as antimicrobials during hydrolysis. Enzyme
loading was performed using Cellic CTec3 and HTec3 (Novozymes A/S, BagsvĂŚrd, Denmark)
at 20 mg protein / g glucan for CTec3 loading, and 10 mg protein/ g glucan for HTec3 loading
when HTec3 was added. Hydrolysis was performed at 50°C with orbital shaking at 180 rpm and
sampling was taken at 6-hr and 72-hr total hydrolysis times. Samples were centrifuged at 13000
x g for 2 min post-incubation and filtered using 22 Âľm mixed cellulose-ester filters (EMD
Millipore, Billerica, MA). Samples were quantified by HPLC (Agilent 1100 Series) equipped
with an Aminex HPX-87H column (Bio-Rad, Hercules, CA, USA) using a mobile phase of 5
mM H2SO4 at 65°C. Glucose yields were determined based on quantified glucose (as glucan)
relative to total glucan available, including non-cellulosic glucan. All hydrolysis runs were
performed in technical triplicate.
Water retention value
Water retention values (WRVs) were determined according to a modified version of TAPPI
UM 256, as previously described [11], with a centrifuge speed of 4000 x g for 15 min. Briefly,
approximately 2.5 g of wet pre-washed biomass samples were soaked in about 150 mL of either
de-ionized water, or one of four NaCl solutions (0.25 M, 0.5 M, 0.75 M, and 1.0 M). After
soaking for 1 hr, samples were rinsed with around 200 mL of de-ionized water to remove salts

95

and vacuum filtered using a 200-mesh Buchner filter. Washed biomass was inserted into a spincolumn (Handee Spin Column Cs4, Thermo Scientific) with a 200-mesh stainless steel
membrane. Biomass was added to form a circular cake devoid of large spaces, with care taken to
avoid inducing drainage of biomass from pressing into column. Non-bound water was drained
via centrifugation at 4000 x g for 15 min. The mass of the drained biomass was determined using
tared aluminum trays, and dried in an oven at 105°C for at least 3 hr, then weighed again. The
WRV was determined as the ratio of the mass of water remaining in the biomass after
centrifuging divided by the mass of dry biomass. WRV was determined in technical triplicate.
Solute exclusion
Solute exclusion was performed following the original protocol outlined by Stone and
Scallon [25], using a series of neutral dextran probes of increasing molecular weight, with some
minor modifications to the procedure. Table 3.1 lists the probes used in this study, which were
obtained from Sigma-Aldrich. For this, approximately 5.0 g of wet, or 1.0 g of oven dried
biomass was added to tared 50-mL centrifuge tubes and re-weighed after biomass addition.
About 10 mL 0.4% (w/v) dextran solution by was added to each tube, and reweighed. Tubes
were vortexed for thorough mixing, and stored at 4°C for 48 hr, with periodic vortexing. A 1 mL
sample was taken, centrifuged at 13,000 x g for 2 min, and transferred to a sealed vial for HPLC
(Agilent 1100 Series) analysis. Biomass solids were thoroughly washed to remove the dextran
probes in centrifuge tubes through dilution with water followed by centrifugation and solidliquid separation, which was repeated three times. Centrifugation was performed at 900 x g for 3
min, with washed biomass dried overnight in a 105°C oven to determine oven dry biomass
weight. Probe concentration was quantified using refractive index on the HPLC, with no column
attached as described previously [19]. Blanks were used to determine any background or non-

96

dextran probe contributions to the refractive index signal. Inaccessible pore volume was
determined as previously outlined [17]. Samples were performed in technical triplicate.

Probe Molecules

Mol. Massa (Da)

Mol. Diameterb (Å)

D-(+)-Cellobiose

352

8

Dextran 6k

6000

39

Dextran 40k

40000

90

Dextran 70k

70000

110

Dextran 100k

100000

136

Dextran 450-650k

550000

310

Dextran 2000k

2000000

560

a

from the manufacturer. b Interpolated from Stone and Scallon 15.

Table 3.1 : Neutral dextran probes used in solute exclusion technique.

Enzyme binding
Enzyme binding experiments were performed on never-dried biomass using 0.2 g biomass
(dry basis)/ 10 mL solution buffer solution containing a cellulase cocktail (Cellic CTec3).
Binding isotherms were determined for protein concentrations over a range of 0 to 3.0 mg/mL
(protein content determined using nitrogen adsorption) in a 50 mM Na-citrate buffer solution at
pH 5.2. Samples were mixed overnight at 4°C using rotary mixing (Intelli-Mixer, ELMI-tech,
Newbury Park, CA). Bound protein mass was determined from the difference between
unbounded protein in free solution to total initial protein concentration determined using the
Bradford method [26]. Bound protein fraction was determined using a standard calibration curve
of initial protein concentrations. Bound enzyme fractions were determined from linear fit of
bound enzyme for the linear region of the Langmuir adsorption isotherm as described in our
previous work [11]. Samples were performed in technical duplicate.

97

Results and discussion
Composition analysis and enzymatic hydrolysis
Composition profiles of residual solids after AHP delignification, LHW pretreatment, and
combinations of the two as shown in Figure 3.1. These conditions were selected to generate
three distinct feedstock groups with significantly different residual solids compositions that
exhibit a wide range of materials for properties testing.

Figure 3.1 : Compositional profiles of corn stover subjected to either AHP pretreatment,
LHW, or LHW followed by AHP pretreatment. Compositional profiles for xylan, crystalline
cellulose, and Klason lignin are shown for alcohol insoluble residue (AIR) corn stover and
pretreated biomass. Composition analysis was performed in technical triplicate (n = 3), with
standard deviations shown on figure.
The results show that AHP-delignified biomass demonstrated a clear trend of increasing
lignin removal with increasing H2O2 loading (Figure 3.1) while retaining most of the xylan as
demonstrated in our previous work [27]. The LHW-pretreated biomass showed significant xylan
removal through solubilization, with the 160°C condition containing about 8% residual xylan by
mass, while the 190°C LHW condition had near complete removal of xylan. Residual biomass
was enriched in cellulose and lignin content for both conditions [28]. The combination of LHW
pretreatment followed by AHP delignification was performed at two AHP loadings (0.06 and

98

0.25 g H2O2 / g biomass) for both LHW pretreatment conditions to generate pretreated feedstocks
with modified xylan and lignin content. These pretreatment combinations represent a wide range
of altered cell wall composition, with cellulose accounting for up to 80% total mass for the most
severe pretreatment condition (Figure 3.1). Interestingly, the lowest H2O2 loading for AHP
delignification (0.06 H2O2 / g biomass) for both LHW-pretreated biomass at both 160°C and
190°C resulted in significantly higher removal of lignin compared to the same AHP
delignification conditions, and may be due to lignin relocalization associated with LHW
pretreatment above the glass transition temperature increasing lignin accessibility to removal by
AHP after LHW pretreatment.
Subsequent enzymatic hydrolysis was performed on never-dried pretreated biomass using
either a cellulase cocktail (Cellic CTec3) or a combined cellulase/hemicellulase cocktail (CTec3
+ HTec3) to measure the impact of cell wall composition and, indirectly, the modification of the
higher order structure of the cell wall matrix by pretreatment, to the rate and extent of cellulose
enzymatic hydrolysis. Results for 6-hr and 72-hr glucose hydrolysis yields, representing initial
rate and final extent of hydrolysis respectively, are shown in Figure 3.2.

99

Figure 3.2 : Enzymatic hydrolysis glucose yields of pretreated corn stover samples. Results
of never-dried hydrolysis of pretreated samples for 6 hr total hydrolysis time are plotted in blue
and dotted green, while 72 hr total hydrolysis times are plotted above in orange and striped gold.
Solid bars represented hydrolysis conditions using only 20 mg protein/g glucan cellulase loading,
while patterned bars represent hydrolysis conditions using both 20 mg protein/g glucan and 10
mg protein / g glucan hemicellulase loading. Analysis was performed in technical triplicate.
These results show that enzymatic hydrolysis using a cellulase and hemicellulase cocktail on
the AHP-only pretreated biomass resulted in high (94%) glucose hydrolysis yields at the highest
H2O2 loading, while LHW-pretreated biomass achieved only a 54% glucose yield at the highest
severity. Interestingly, a glucose yield of only 83% was observed for the highest severity twostage pretreatment, which can be attributed to cell wall structural changes from the two-stage
pretreatment limiting enzymatic hydrolysis. Reductions in cellulose accessibility resulting from
cellulose aggregation induced by sufficient removal of non-cellulosic components in the cell wall
would explain both reduced hydrolysis kinetics, as well as limited cellulose hydrolysis [29].
The cellulase enzyme cocktail used in this study (i.e., CTec3) has been shown to contain a
suite of accessory enzymes including xylanase and pectinase activities [30]. Furthermore, this
cocktail is capable of effectively hydrolyzing pretreated biomass with low to moderate

100

hemicellulose content without any additional accessory enzymes [30]; however, significantly
lower yields are observed in pretreated feedstocks containing a substantial fraction of the original
xylan, such as the AHP-only samples [31]. In the AHP-only case, delignification may increase
enzymatic hydrolysis effectiveness, however as reported in prior studies, cellulose accessibility
is likely inhibited by both lignin and xylan content [32], with lignin limiting accessibility to
xylan-coated cellulose microfibrils. Therefore, for AHP-only pretreated biomass, we can
hypothesize that without removal of xylan, significant regions of the cell wall matrix remain
partially inaccessible to cellulase binding. This result is further exemplified by the similar yields
observed between the cellulase only and cellulase plus hemicellulase for the 160°C LHW twostage pretreatments (Figure 3.2), which contain significantly lower xylan content compared to
the AHP-only pretreatments. Enzymatic hydrolysis using only the cellulase cocktail also resulted
in notable lower 6-hr glucose hydrolysis yields for all samples, indicating that lack of xylanase
and pectinase hinders initial hydrolytic rate of cellulases due to xylan limiting cellulose
accessibility. This conclusion is further supported by higher initial yields associated with highly
delignified samples, with delignification acting as an alternative avenue to increase enzyme
accessibility [33].
Non-productive adsorption may also play a role in limiting enzymatic hydrolysis
effectiveness, as there was a noted increase in enzymatic hydrolysis yields between the cellulase
only and cellulase plus hemicellulose cases for the two-stage 190°C LHW-pretreated biomass, as
xylan content accounted for less than 2% of total cell wall mass. This result indicates higher
overall enzyme loading may limit the effect of non-specific binding of enzymes to lignin by
providing more protein for adsorption to the charged surface of lignin, effectively neutralizing
the lignin surface and promoting cellulase adsorption to cellulose.

101

Cell wall swelling in response to increasing solvent ionic strength
Water retention value is a property that can be used to capture a number of different
structural features of the cell wall matrix as they relate to cell wall association with water. As
demonstrated in our recent work, WRV can be used to predict enzymatic hydrolysis in two
different alkaline pretreatments under a range of conditions [34], with WRV able to capture
higher order changes in the cell wall associated with nanoscale porosity and cellulose
accessibility in compositionally diverse pretreated samples. WRV is used similarly in this study
to understand changes in higher order structure, however WRV are lower in this work due to
changes in methodology increasing the amount of dewatering compared to prior work [11].

Figure 3.3 : Cell wall swelling measured using water retention in the absence and presence
of a salt. Water retention value (WRV) of never-dried corn stover at varying pretreatments
measured after incubation in water only (blue bars) or 1 M NaCl (orange bars). Samples were
performed in technical triplicate (n = 3), with standard deviation shown on figure.

In the present work, the WRV without an added solute was found to increase with
increasing H2O2 loading in the AHP-only pretreatment case, as well as increasing severity for
LHW pretreatment (Figure 3.3). Increases in WRV were also observed between low and high
H2O2 loading for individual two-stage pretreatments. These results indicate delignification by
102

AHP and xylan removal by LHW pretreatment can increase the amount of water associated with
the cell wall matrix. A noted decrease in WRV in the 190°C LHW two-stage pretreatments may
be attributed to two contributions; the first suggesting delignification in low-xylan cell walls
removes sufficient spacer components that could be hypothesized to result in cellulose
aggregation and collapse of the cell wall matrix, limiting available sites for water entrapment.
The second contribution may be due to lignin content making up a larger composition fraction of
the 190°C LHW two-stage pretreatments compared to the AHP-only and 160°C LHW two-stage
pretreatment, and subsequently limiting the amount of cell wall swelling.
The values for WRV in the presence of an added solute showed noted decreases for the high
H2O2 loading AHP-only samples, as well as 160°C LHW two-stage pretreatments, while no
decrease in WRV was observed in the 0 g/g H2O2 AHP-only sample or the LHW only samples.
The impact of increasing solvent ionic strength on cell wall water retention has been investigated
largely in the context of pulp and paper applications [35]. From our results, delignification
appears to be the primary contributor to solute-induced changes in WRV, and can be interpreted
as a change in cell wall rigidity. Within the secondary cell walls of grasses, lignin forms a rigid
matrix, with ferulate-mediated cross-links between within lignin and between lignin and xylans
preventing physical cell wall expansion in response to the solvent environment [36.37]. Lignin
content within the cell wall correlates to the extent of enzymatic hydrolysis in grasses, with
greater than 50% cell wall delignification associated with high (>80%) hydrolysis yields [9].
Disruption of lignin structure through sufficient cleavage of cross-links and solubilization can be
proposed to contribute to a loss of cell wall rigidity, allowing the cell wall to exhibit hydrogellike behavior in the presence of solutes.

103

The equilibrium swelling of hydrogels is known to depend on factors that include the crosslink and charge densities of the polymer networks, while increases in the number of ionic groups
in hydrogels can increase the swelling capacity by increasing osmotic pressure that driving the
infiltration of solvent into the polymer network [38]. This behavior has been observed in
carboxymethyl cellulose gels and to a lesser extent sulfite pulps, which exhibited changes in
WRV corresponding roughly to the content of acidic groups [35]. AHP pretreatment has been
shown to increase the cell wall carboxylate content [11], which would explain the swelling
behavior of AHP-delignified biomass, although solute-induced changes in WRVs are not as
pronounced as prior work using chlorite-delignified pulps [39]. The extent of non-covalent
physical association of cell wall components contributes to the ability of the cell wall to swell, as
cell wall swelling not as pronounced in the two-stage 190°C LHW-pretreated biomass, which
can be hypothesized to be due to partial cell wall coalescence.
Accessible volume distributions determined by solute exclusion
A set of dextran probes was next employed to assess the accessible nanoscale volume within
the pretreated biomass. Accessible volume distributions were determined by subtraction of
inaccessible volumes from the inaccessible volume at the largest probe size (560Å), representing
the total volume for a sample [15]. It should be noted that these volumes are not direct
measurements of cell wall pores, rather effective pore diameter based upon idealized probe
diameter in solution [40]; and thus, will be referred to as probe diameter in this work, shown in
Figure 3.4.

104

Figure 3.4 : Accessible volume distributions measured by solute exclusion. Accessible
volumes of never dried corn stover were determined using neutral dextran probes of average
molecular weight range (see Table 3.1), giving a range of effective hydrodynamic radiuses. (A)
AHP-only pretreatment conditions and (B) LHW and the two-stage pretreatments are shown on
the right figure. Samples were performed in technical triplicate (n = 3), with standard deviation
shown on figure.

From these results, there are clear differences in accessible volumes between AHP-only,
LHW-only, and two-stage pretreatments. In the AHP-only samples, accessible volume was
restricted to only smaller (<12 Å) probe sizes for the 0 g/g H2O2 pretreated sample, while
increasing H2O2 loading resulted in significant increases in accessible volumes for both small
and large probes, indicating the formation of larger (>39 Å) porous voids [15]. Studies using
chlorite-delignified sugarcane bagasse have yielded highly similar accessible volume
distributions to the AHP-only pretreated samples in this work, indicating that delignification of
diverse graminaceous biomass feedstocks results in similar cell wall responses, and that lignin
removal may result in the formation of water-accessible surfaces within the cell wall [21].
Accessible volume distributions in the LHW samples showed some noted increases
accessible volumes for probes between 39-136 Å between the 160°C LHW and 190°C LHW
condition, however larger probe volumes were similar to the 0 g/g AHP-only pretreated sample.
In addition, overall accessible volumes were considerably lower compared to AHP-only

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pretreated samples, indicating limited accessible volume formation in LHW-only samples. One
explanation for the limited accessible volumes observed in LHW only samples may be due
melting and relocalization of lignin during pretreatment from spaces between cellulose
microfibrils to the cell wall surface [41] together with xylan solubilization that may result in
coalescence of cellulose microfibrils, decreasing the enzyme accessibility of the cellulose
(Figure 3.5C).
For sequential two-stage pretreatment, low H2O2 loading coupled with the 160°C LHW
pretreatment condition resulted in accessible volume distributions comparable to the highest
H2O2 loading for AHP-only pretreatment condition. Increased H2O2 loading of the 160°C LHW
condition resulted in higher accessible volumes for the larger probes coupled to minimal
increases in smaller probe accessible volumes. This trend was also observed between the low and
high H2O2 loading for the 190°C LHW condition, and likely indicates the formation of larger
porous regions with increased H2O2 loading rather than the formation of more porous regions in
the cell wall as observed in the AHP-only conditions. Smaller probe (<136 Å) accessible
volumes were also considerably lower in the 190°C LHW two-stage pretreatments compared to
the 160°C LHW conditions, and support the previously proposed mechanism of either a collapse
of inter-lamellar layers within the cell wall at high overall pretreatment severity, or through
cellulose aggregation reducing cell wall porosity due to the lack of xylan to act as a spacer [29].
Based upon results from enzymatic hydrolysis, WRV, and solute exclusion, the following model
(Figure 3.5) was derived for cell wall changes resulting from pretreatment.

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Figure 3.5 : Structural model for pretreatment effect on cell wall organization. Transverse
section of cellulose microfibrils for (A) native cell wall, (B) AHP-pretreated cell wall, (C) LHWpretreated cell wall, and (D) two-stage AHP pretreated LHW cell wall. AHP pretreatment results
primarily in reduction in lignin content, promoting the formation of porous regions and inducing
cell wall swelling. LHW pretreatment primarily removes xylan and relocalizes lignin, increasing
accessible surface area. Two-stage pretreatments contain significant reductions in both xylan and
lignin content, increasing porous regions, and in the case of 190°C LHW, sufficient noncellulosic component removal results in cell wall aggregation. Orange squares representing
cellulose microfibrils, green lines representing xylan and gray clouds representing lignin.

Drying-induced pore collapse on cell wall properties and enzymatic hydrolysis
Drying-induced hornification was investigated for select pretreatment conditions to
determine the impact that changes in higher order structure of pretreatment-modified cell walls
107

have on cell wall swelling, accessible volumes, and enzymatic hydrolysis yields in
compositionally diverse samples.

Figure 3.6 : Impact of oven drying on cell wall properties and enzymatic hydrolysis. (A)
Accessible volumes of oven-dried and rehydrated corn stover were performed similarly to
Figure 3.4. (B) Water retention value (WRV) of oven dried corn stover at varying pretreatments.
(C) Hydrolysis yields for oven-dried (OD) samples with 6-hr total hydrolysis time are plotted in
blue, and 72-hr total hydrolysis time are plotted in orange. Comparative hydrolysis yields for
never-dried (ND) 6-hr and 72-hr total hydrolysis time from Figure 3.2 are shown as green dotted
and yellow striped bars respectively. Hydrolysis conditions used 20 mg protein/g glucan CTec3
and 10 mg protein / g glucan HTec3 loading. Experiments were all performed in technical
triplicate, with standard deviations are shown on each figure.

The results show that oven-drying resulted in significant changes in accessible volume
distributions (Figure 3.6A), with decreases in accessible volume observed to some extent for all
samples relative to the never-dried samples. There was however, a stark contrast in how
significantly accessible volumes decreased, with samples subjected to AHP delignification,
demonstrating highly altered accessible volumes compared to 0 g H2O2 / g biomass AHP and
190°C LHW only pretreated biomass, which demonstrated only minimal loss of accessible

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volumes [42]. Furthermore, there was a noted decrease in volumes accessible to larger probes in
samples subjected to AHP delignification. These results can be related to the structural model
presented in Figure 3.5, with drying resulting in the collapse of larger porous regions within the
cell wall (Figure 3.5B), while samples containing higher fractions of lignin (Figure 3.5C)
maintained their ability to sorb water within nanoscale pores [43].
WRVs were also observed to substantially decrease (Figure 3.6B), with all samples
experiencing at least a 40% decrease in WRV after drying and exhibited a closer distribution of
WRV between pretreated biomass compared to the never-dried WRV. Hornification has been
demonstrated to reduce accessible internal surface area within delignified wood fibers that limits
water sorption [44,45]. This can be visualized in Figure 3.5D, with oven-drying removing water
and resulting in the irreversible coalescence of some cell wall components [46]. For the
enzymatic hydrolysis yields after oven-drying, the biomass delignified at high H2O2 loadings
demonstrated the largest decrease in 6-hr hydrolysis yields (Figure 3.6C), followed by less
significant decreases for the other pretreated samples. The 72-hr hydrolysis yields were only
significantly lower in the 0.25 g/g AHP-only pretreated biomass. As the AHP-delignified
biomass is the sample most impacted by drying-induced hornification, this indicates that the
hydrated spaces within the cell wall of these samples, presumably containing mostly xylan, are
most susceptible to irreversible coalescence. Interestingly, prior work using 0.50 g/g H2O2 AHPdelignification followed by lyophilization resulted in no differences in final hydrolysis yields
[31]. Different methods of drying have been shown to influence extent of hornification, and have
been explored in the context of altered cell wall properties [47].

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Enzyme binding
As noted earlier, the commercial cellulase cocktail contains a number of accessory enzyme
activities other than cellulase. As a result, the observed bound protein curves represent general
protein binding rather than for binding of a specific cellulase. In addition, contributions from cell
wall biopolymers other than cellulose (i.e., hemicellulose and lignin) can contribute to the
observed adsorption behavior through non-specific or non-productive binding [48]. However,
binding isotherms are useful for assessing the impact of pretreatment on enzyme-accessible
surfaces.

Figure 3.7 : Bound enzyme fractions of CTec3 on pretreated biomass. Bound enzyme
fractions are shown as a percent of bound enzyme to total enzyme loading. Binding
concentrations were performed in technical duplicate, with standard deviations shown on figure.

The results in Figure 3.7 show the percent of bound enzyme measured as a fraction of total
enzyme in solution for a linear range of enzyme loadings representing the linear, lowconcentration region of the binding isotherm. The percentage of bound enzyme increased with
increasing H2O2 loading for AHP-only pretreated biomass, comparable to results published in
our prior work [11]. The LHW-only pretreated biomass had a lower percentage of bound enzyme

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compared to the higher severity AHP-only pretreated biomass, indicating that structural changes
induced by LHW pretreatment alone do not impact enzyme adsorption the same extent as AHPdelignification. The two-stage pretreatments showed increases in the fraction of bound enzyme
with both H2O2 loading, and to a lesser extent with LHW pretreatment severity, however low
H2O2 loadings were still comparable or higher than all but the highest H2O2 loading AHP-only
pretreated biomass.
Correlation of properties
By correlating compositional and structural properties of pretreatment-modified plant cell
walls to hydrolysis yields, a number of important trends can be identified that provide insight
into the system.

Figure 3.8 : Comparisons between glucose hydrolysis yields, solute exclusion, water
retention value, and lignin content. (A,B) Glucose hydrolysis yields taken from Figure 3.2
were from 72 hr total hydrolysis time using an enzyme solution of CTec3 and HTec3. Xylan
content is displayed as a range for each linear trend in orange on (A). Accessible volume
distributions (C, D) were used from Figure 3.4, and lignin content was taken from Figure 3.1.
111

The first major trend that can be observed is that comparing 72-hr glucose hydrolysis yields
to lignin content (Figure 3.8A) shows three distinct linear relationships between AHP-only, and
each LHW condition. From this, lignin content can be shown correlated to hydrolysis in the
context of increased H2O2 loading, and therefore extent of delignification result in increased
enzymatic hydrolysis yields. The major compositional difference between each grouping in
Figure 3.8A is the amount of xylan present (shown as a range for each grouping on Figure
3.8A), which can be taken as a proxy for the extent of cell wall modification during LHW
pretreatment.
Next it was observed was that glucose hydrolysis yields were positively correlated with 90
Å probe accessible volumes (Figure 3.8B). Cell wall volumes accessible to the 90 Å dextran
probe were selected to correlate with hydrolysis yields, as 90 Å corresponds to a slightly larger
probe size than necessary for reasonable estimation of typical cellulase (e.g., TrCel7A) [49].
Notably, as the accessible volumes increased, the 72-hr hydrolysis yields were observed to
approach saturation. This trend is reasonable, because although accessible volumes may indicate
increased accessibility within the cell wall for cellulases, there is likely a limit to the extent
accessibility plays in increased enzymatic hydrolysis relative to other intrinsic factors [50].
Interestingly, for this data set the accessible volumes across multiple probe sizes were shown to
directly correlate to lignin content, with AHP-delignified biomass exhibiting a strong negative
correlation with accessible volumes at 39 Å, 90 Å, and 136 Å probe sizes (Figure 3.8C), while
LHW-only and two-stage pretreated biomass accessible volumes correlated only to the 136 Å
probe size (Figure 3.8D). These correlations indicate that for biomass subjected to AHP
delignification, increases in the nanoscale probe-accessible volumes can be directly related to the
extent of delignification as proposed in the schematic in Figure 3.5B. Accessible volumes

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observed in AHP-delignified biomass also likely contribute to enzyme penetration within the cell
wall, with prior correlations between enzyme binding and accessible pore volumes shown in
sulfite pretreated hardwoods [49] and alkali-delignified hardwoods [51].

Figure 3.9 : Comparisons between glucose hydrolysis yields and (A) bound enzyme
fractions and (B) water retention value. Glucose hydrolysis yields taken from Figure 3.2 were
from 72 hr total hydrolysis time using an enzyme solution of CTec3 and HTec3. WRV were
taken from Figure 3.3, and bound enzyme fractions were taken from Figure 3.7.

The fraction of bound enzyme was demonstrated to exhibit a strong positive correlation to
glucose hydrolysis yields irrespective of pretreatment conditions or composition (Figure 3.9A).
This result is not surprising as cellulase binding and glucose hydrolysis yields are well-known to
be correlated in diverse feedstocks subjected to diverse compositional changes resulting from
pretreatment [11,32]. Correlation plots of WRV versus 72-hr hydrolysis yields two distinct
trends (Figure 3.9B), with the AHP-only pretreatment resulting in a clear linear trend between
WRV and hydrolysis yields as demonstrated in our prior work [11,34], while a second trend with
a different slope was observed for the two-stage 160°C LHW-pretreated biomass. No trend was
observed for the two-stage 190°C LHW-pretreated biomass followed by AHP delignification
(Figure 3.9B), and based upon these results, there appears to be some relationship between
glucose hydrolysis yields with water-accessible surface area in pretreated biomass not exhibiting
cell wall coalescence due to significant non-cellulosic component removal. In addition, these

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distinct, pretreatment-dependent trends are comparable to what we identified in our prior work
with AFEX-pretreated corn stover or switchgrass exhibiting distinct trends for at different
ammonia loadings [34].
As a final correlation between properties, WRV has also been proposed as a proxy for
measuring fiber saturation point (FSP) [52], which is generally defined as the total inaccessible
volume at 560 Å [15]. Additionally, WRV and FSP are hypothesized to measure similar
properties in pulps [53]. Linear comparisons between the WRVs and FSPs for never-dried and
oven-dried pretreated biomass measured show distinct linear trends (Figure 3.10).

Figure 3.10 : Comparison of inaccessible pore volume to water retention value. Total
inaccessible pore volume was determined using solute exclusion at the effective pore size of 560
Å and compared to WRV (0 M NaCl) from Figure 3.3.

In the never-dried samples, the slope of the linear relationship was less than one, indicating
WRV measurements were higher than the equivalent inaccessible volume measurement. This is
important, because one of the limitations of the solute exclusion method resides in the inability
for water in porous regions with non-uniform or narrow openings to interact with the dextran
probes during the solute exclusion technique [17], while WRV accounts for water localized
within porous regions of irregular geometries [25]. After oven drying, the slope between

114

inaccessible volume and WRV was effectively unity, indicating that the solute exclusion and
WRV techniques quantify similar cell wall properties after oven drying. The substantial drop in
these properties is clearly attributed to hornification, resulting in the collapse of certain cell wall
porous regions and reductions in accessible surfaces shown previously (Figure 3.6). Based upon
this comparison, WRV likely over-measures total accessible surface area compared to the FSP in
the never-dried case, which may be due to WRV also incorporating water-cell wall binding in the
form of intra-lamellar water association within cellulose microfibrils or between microfibrils not
be quantified by the FSP. Based upon the two-stage LHW pretreated biomass exhibiting higher
WRV relative to FSP (Figure 3.10) compared to the AHP-delignified only biomass, this may be
an appropriate evaluation, as xylan removal could increase water-cellulose microfibril
interactions [54], while still being inaccessible to quantification by the solute exclusion method.

Conclusions
In summary, AHP delignification and LHW pretreatment of corn stover in this study yielded
a series of compositionally diverse feedstocks exhibiting significantly different cell wall
properties that were used to assess how both compositional and structural features impact cell
wall recalcitrance to enzymatic hydrolysis. Specifically, we were clearly able to demonstrate
that, while the mechanisms by which AHP and LHW pretreatment improve cellulose
accessibility are dramatically different, both pretreatment types were shown to increase
hydrolysis yields, enzyme sorption, and WRV. AHP delignification was shown to result in the
substantial increase in the accessible volume to a series dextran probes, while LHW pretreatment
resulted in only changes to smaller pore distributions. Solute-induced cell wall swelling
measured by WRV showed AHP-delignified pretreatments displayed partial loss of cell wall
rigidity, which may explain in part larger accessible volume distributions and increased porosity
in AHP-delignified biomass. Overall, this study highlights the importance that cell wall

115

organization and its modification during processing during biorefining can have on feedstock
response to enzymatic hydrolysis.

116

APPENDIX

117

APPENDIX B: Supplementary figures and additional results

Figure 3.11 : Solute-induced swelling of biomass at multiple salt concentrations. Water
retention value (WRV) of never dried and oven dried corn stover at varying pretreatments
measured without salt (0 M NaCl), and after soaking in salt solutions (0.25 -1 M NaCl). (A)
shows AHP-only pretreatment conditions, (B) shows LHW-only pretreatment conditions, (C)
shows two-stage pretreatment conditions, and (D) shows oven-dried pretreatments tested.
Samples were performed in technical triplicate (n = 3), with standard deviation shown on figure.

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Figure 3.12 : Enzyme binding curves of CTec3 on never-dried biomass and linear fits for
bound enzyme fractions. Bound CTec3 concentrations are displayed as individual data points,
with a linear best fit representing the relative ratio of bound enzyme to total enzyme loading. (A)
shows AHP-only pretreatment conditions, (B) shows LHW-only pretreatment conditions, and
(C) shows two-stage pretreatment conditions. Binding concentrations were performed in
technical duplicate (n = 2), with standard deviations shown on figure.

Figure 3.13 : T2 NMR relaxation curves of water interacting with pretreated biomass. (A)
Relaxation curves of AHP-only pretreatment samples, (B) relaxation curves of 160°C LHW and
two-stage pretreatments, and (C) relaxation curves of 190°C LHW and two-stage pretreatments.
Samples were performed and fitted in technical duplicate (n = 2), with standard deviations shown
on figure.

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Figure 3.14 : T2 NMR relaxation curves monocomponent T2 relaxation times. Relaxation
times were determined from a mono-exponential component fit of each relaxation curve.
Samples were performed and fitted in technical duplicate (n = 2), with standard deviations shown
on figure.

Figure 3.15 : 1H-NMR relaxometry of AHP-delignified corn stover showing (A) spin-spin
(T2) relaxation curves and (B) correlation of monocomponent exponential fits of these
relaxation curves to the corresponding WRV and glucose hydrolysis yield.

The following was published as part of an original research paper submitted to Bioresource
Technology. It is of note that the data used in this study is not the same data presented earlier in

120

this supplementary, rather from earlier work performed. However, the methods and objectives
were similar, and therefore included in the Appendix.
1

H spin-spin (T2) NMR measurements were carried out in a Bruker static probe at

frequencies of 300.103MHz on a Varian 300 MHz NMR interfaced with a Dell Precision T3500
desktop running CentOS 5.6 with VnmrJ 3.2A. Spin-spin relation times were determined using a
standard 2D Carr-Purcell-Meiborn-Gill (CPMG) sequence with a 5 ¾s (90°) 1H pulse, 10 ¾s
(180°) 1H pulses, 16 scans, 10 s recycle delay and 𝜏 = 0.0002. Eight data points were recorded
with 4 to 1024 echoes, and data was analyzed and reported as a monocomponent exponential fit
of the data [6]. Technical replicates were performed at a constant temperature (25 °C) and a
sample total solids content of 25% (wt sample per wt total).
Another approach to measure physical and chemical environment involves using T2 NMR
relaxometry to assess water constraint. Similar to DSC, T2 NMR relaxometry provides a
quantifiable relationship between biomass-water interactions, and results are dependent on both
the physical and chemical environment experienced by the solvent, with more solvent constraint
resulting in shorter T2 relaxation curves. Monocomponent exponential decay curves from CGMG
T2 relaxation studies for AHP-delignified corn stover (Figure 3.15A) showed a general trend of
increased chemical constraint with increasing extent of delignification. Relaxometry curves were
similar for low and moderate lignin removal, however become increasingly more constrained as
more lignin was removed.
Increased water constraint is an indication of increased hydrogen bonding associated with
biomass, and may be due to delignification increasing available hydroxyl content and carboxyl
content [11] or due to increased accessibility to cellulose for hydrogen bonding rather than
changes in surface area or particle size affecting porosity [14]. These conclusions are further

121

supported by the correlation between T2 monocomponent exponent fits and enzymatic hydrolysis
yields (Figure 3.15B), indicating that systems with more constrained water, have greater
cellulose accessibility and result in higher enzymatic hydrolysis yields. The linearity between
WRV and T2 monocomponent exponent fits (Figure 3.15B) indicates that WRV and T2 NMR
relaxometry likely quantify the same biomass properties in AHP-delignified samples.

122

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Chapter 4 : Impact of impaired xylan biosynthesis on cellulose
microfibril organization in Arabidopsis thaliana secondary cell
walls
Introduction
Secondary cell walls represent a key biological and biomechanical component integral to
vasculature, strength, and development in land plants. With the main role of primary cell walls to
control cell elongation and determine cell shape, secondary cell walls are deposited after cell
wall growth to impart mechanical strength and promote upright growth of the plant [1]. The
secondary cell wall is generally comprised of three major classes of polymers, cellulose,
hemicelluloses, and lignin, which together form a three-dimensional cell wall matrix.
Cellulose is the major biopolymer present in both primary and secondary cell walls, and
is composed of linear chains of (1,4)-linked-β-D-glucosyl residues, which stack together with
other cellulose chains via inter- and intra- hydrogen bond networks and van der Waals forces to
form cellulose elementary microfibrils [2]. Cellulose is synthesized at the plasma membrane by
cellulose synthase enzymes that are organized into multi-protein cellulose synthase complexes
(CESA), with 10 CESA isoforms in Arabidopsis thaliana. Three distinct CESA isoforms are
required for normal synthesis, with CESA1, CESA3, and CESA6 involved in primary cell wall
synthesis [3], while CESA4, CESA7, and CESA8 are required for secondary cell wall synthesis
[4]. Specific cellulose orientation in parallel arrays within the cell wall is thought to be for
structural strength, imparted by cortical microtubules [5], however recent evidence has shown
cortical microtubules are not necessary to maintain directionality [6]. Live cell imaging of CSCs
during secondary cell wall formation indicate that dense formations of CSC complexes
contribute to the assembly of closely associated cellulose microfibril sheets and move in

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unidirectional concerted fashion [6] in epidermal cells transdifferentiated into secondary cell
wall-synthesizing xylem-like cells [7].
In higher plants, the structure of xylans are comprised of a β-(1-4)-linked xylosyl
backbone with a multitude of side chain modifications, resulting in arabinoxylans,
glucuronoxylans, (arabino)glucuronoxylan, (glucurono)arabinoxylan, and heteroxylans [8]. In
dicots such as Arabidopsis, glucuronoxylan (GX) is the predominant xylan in the secondary cell
wall, containing glucuronic (GlcA) or 4-O-methyl glucuronic acid (MeGlcA) substituted at the
Xyl O-2 position on a xylosyl backbone [9], as well as acetylation at some O-2 and O-3 positions
[10]. A reducing end terminal tetrasaccharide group of xylan comprised of a 4-β -D-Xyl-(1,4)-β D-Xyl-(1,3)-ι -L-Rha-(1,2)-ι -D-Gal-(1,4)-D-Xyl

is found in some dicots including Arabidopsis,

and is thought to play a terminating role in xylan chain length during biosynthesis [11, 12].
Recent advances have identified many proteins involved in xylan biosynthesis and
functional group addition, however the exact function of many of these genes is still unknown.
Notably, synthesis of the xylosyl backbone is impaired when proteins of the glycosyltransferase
family GT43 (IRX9, IRX9-L, IRX14, IRX14-L) or GT47 (IRX10, IRX10-L) gene families are
mutated [13-15]. While the inferred function of these genes is xylan synthase activity, to date
only IRX10, IRX10-L, and homologs of these exhibited xylan synthase activity in vitro by
heterologous expression and purification [16, 17]. IRX9 function has been hypothesized as a
non-catalytic role as a membrane anchor for a hypothetical larger xylan synthase complex,
analogous to some pectin biosynthesis proteins [18]. In addition, likely methyl transferases from
the DUF579 family of genes have been shown to affect xylan biosynthesis, with IRX15 IRX15-L
genes identified to significantly reduce both xylan content, and average chain length [19, 20].
Together with irx9 and irx10 mutants, reductions in xylan chain length correspond to parallel

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observations of higher than wild type ratios of MeGlcA/GlcA in all three mutants, and has been
hypothesized to relate to GlcA methylation rate keeping pace with reduced glucuronoxylan
backbone synthesis rate, resulting in the methylation of nearly all GlcA units in GX backbone
mutants in irx9 [11]. Glycosyl transferases from the families GT47 (FRA8 and FRA8H) and
GT8 (IRX8 and PARVUS) mutations have been shown to impact the terminating tetrasaccharide
group occurrence, resulting in dispersed distribution of glucuronoxylan chain lengths, and a
reduction in amount of glucuronoxylan chains [11, 21, 22]. GlcA is substituted to the xylan
backbone by the GT8 family GUX proteins (GUX1-5) [23]. Acetylation of xylan occurs via
members of the ESKIMO1/TRICHOME BIREFRINGENCE-LIKE (ESK1/TBL29) protein
family including ESK1 [17]. Knockout of xylan acetyltransferses result in reductions in xylan
corresponding to collapsed xylem and dwarfed phenotype [24]. Methylation of glucuronic acid
side units is controlled by GXMT proteins [25].
Many xylan mutants are categorized as part of a larger group of irregular xylem (irx)
mutants, that represents a category of Arabidopsis mutants that have collapsed and deformed
xylem vessels [26], and are attributed to perturbations of genes involved in secondary cell wall
biosynthesis. To date, there are fifteen irx mutant phenotypes identified and excluding a
collapsed xylem phenotype, a wide range of enzymes or transcription factors involved in the
synthesis of all secondary cell wall components, giving irx mutants a diverse range of
characteristics [27]. In addition, some mutants display pleiotropic effects, which convolute
identification of cell wall response to gene perturbations. One hypothesis for irregular or
collapsed xylem observed in irx mutants is due to the inability of the cell wall to organize
spatially in a way conducive resist compressive forces during stem development [26], which may

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be a result of reductions in cell wall components, or alterations to the molecular architecture of
components.
The role of xylan structure and interactions with other polymers within the secondary cell
wall matrix is not fully understood, however as the main hemicellulose in secondary cell walls of
dicots, the loss of plant fitness with perturbations to both xylan structure and abundance suggests
that xylan plays a crucial role in promoting normal plant growth and development. Xylan has
been proposed to non-covalently coat cellulose microfibrils and crosslink cellulose microfibrils
and covalently bind to other polymers [18]. Recent elucidations of substitution patterns [28, 29]
coupled with molecular simulations [30] and direct NMR evidence [31] indicate that xylan
hydrogen bonds to cellulose in a twofold helical screw confirmation (one 360° twist every two
glycosidic bonds) in the secondary cell wall. Both cellulose structure [31] and xylan substitution
patterns [32] impact xylan-cellulose binding patterns, as well as the availability for covalent
interactions with lignin [27]. Electrostatic charge of xylan substitution may also impact hydrogen
bonding interactions, as glucurosylation was shown to rescue the growth phenotype of acetyl
deficient tbl29 mutants [33].
Model systems such as bacterial cellulose synthesis have been employed to understand
cellulose-polymer interactions during cellulose synthesis, with prior studies indicating that both
xylan [34] and xyloglucan [35] can impact cellulose crystallization. Substituted mannans can
also induce differential levels of cellulose fibril organization, with varying extent of microfibril
non-covalent cross-links between individual fibrils depending upon mannan substitution
frequency [36]. Mixed-linked glucans likely play a similar role to xyloglucans in primary cell
walls as evidenced by mixed-linked glucan deficient rice mutants exhibiting fewer cellulose
microfibril cross-links, likely resulting in a weakened cell wall [37]. Lignin perturbations in

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Arabidopsis resulted in varied degrees of cellulose disorganization related to the severity of the
lignin phenotype [38], and indicated cellulose microfibril organization within the secondary cell
wall is likely dependent in part on electrostatic interactions imparted by non-cellulosic
components. Furthermore, disrupted cell wall structure has been associated with reductions in
stem mechanical strength, and indicates cell wall strength is directly related to secondary cell
wall organization and regularity to maintain cell wall fitness [23].
While notable strides have been made in characterizing many genes and proteins
associated with secondary cell wall synthesis, less focus has been placed on identifying the
underlying structural mechanisms maintain cell wall regularity, with descriptors for cell wall
phenotypes often limited to cell specific descriptors (i.e. irx - irregular xylem mutants). One
major reason for limited information on the structural mechanisms resides in the difficulty of
directly observing higher order interactions within the secondary cell wall, requiring indirect
metrics to characterize cell wall organization through microscopy techniques or chemical
analysis. In the case of xylan integration within the cell wall, molecular simulations and NMR
evidence have implicated xylan side chain structure significantly impacts xylan-cellulose
interactions, however the importance of xylan structure in promoting normal cellulose
organization within the cell wall has not been directly observed to date. As such, there currently
exists a barrier in the literature between understanding fundamental cell wall biosynthesis from
the molecular perspective and translation of knowledge to observed macroscopic ultrastructural
features.
The goals of this study are to characterize secondary cell wall organization in a series of
altered xylan mutants to (1) demonstrate synergy between non-direct techniques of cell wall
characterization and direct cell wall visualization to identify phenotypes and (2) explore the

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fundamental role that xylan-cellulose interactions play in secondary cell wall organization and
translate nanoscale cell wall observations to macroscopic properties; relating cell wall
architecture and organization to function at the cellular and whole plant scale.

Materials and methods
Plant lines and growth conditions
Plant genotypes used were Arabidopsis wild type (Col-0) and T-DNA insertion mutants
irx9 (Salk_057033) [11], irx10 (Salk_055673) [39], irx15 irx15-L (GK_735E12,FLAG_532A08)
[20] and irx15 irx15-L complementation lines, irx15 irx15-L IRX15-L (Line #1) [20] and irx15
irx15-L OE-IRX15-L (Line #6) [20]. Seeds were planted in wet peat pellets and cold treated for
48 hours at 4 °C, then transfer to growth chamber. Light/dark period was 16/8 hours at light
intensity 150 mE with light/dark temperature set at 23/20 °C. Humidity was not controlled. After
three weeks in the peat, pellets plants and pellets were transferred to individual pots 8x8x12 cm
with SureMix soil (Surefill, http://www.surefill.com/) and 3-4 plants per pot. Stems were
harvested post senesce and lyophilized to moisture content 5% (g H2O/g biomass). Samples postmilling were stored in air-tight containers for further use. Particle size reduction for water
retention, wide angle x-ray scattering, and enzymatic hydrolysis was performed using a Wiley
Mini-Mill (Thomas Scientific, Swedesboro, NJ) with a 30-mesh screen.
Composition analysis
Cell wall lignocellulosic material was isolated following extraction and de-starching
procedure outlined [40] using three sequential washes of 70% ethanol, 1:1 methanol-chloroform,
and acetone to obtain alcohol insoluble residue (AIR). AIR was destarched using 50 Âľg Amylase
/ml H2O (Bacillus species, Sigma-Aldrich, St. Louis, MO) and 18.7 units of Pullinase (Bacillus
acidopullulyticus, Sigma-Aldrich, St. Louis, MO) in a 0.01% sodium azide solution, with rotary
mixing at 37°C overnight.
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Non-cellulosic neutral monosaccharide content of the wall matrix polysaccharides was
obtained by treating de-starched AIR with trifluoracetic acid (TFA) followed by derivatization
using the alditol acetate method with minor changes [40]. Crystalline cellulose content was
isolated following TFA extraction using Updegraff reagent, followed by cellulose hydrolysis
with 72% sulfuric acid and quantification of the subsequent monosaccharide with the anthrone
colormetric assay. Acetyl bromide soluble lignin (ABSL) was determined as described
previously [41]. Samples were performed in technical triplicate and biological quadruplicate (n =
12) unless otherwise specified.
Glycome profiling
Previously milled stem samples were further ball-milled with a TissueLyser II (Qiagen
Inc., Germantown, MD) in preparation for sequential extraction. Sequential cell wall extractions
and glycome profiling of switchgrass organs were carried out as described previously [42, 43] on
AIR biomass. Plant glycan-directed monoclonal antibodies (mAbs) were from laboratory stocks
(CCRC,JIM, and MAC series) available at the Complex Carbohydrate Research Center [44]. A
description of the mABs used in this study can be found in the supplementary materials of prior
work [45].
Wide angle x-ray scattering
XRD measurement were performed on a Bruker Davinci Diffractometer system (Bruker,
Camarillo, CA). The diffracted intensity of Cu Kα radiation (with Nickel filter; λ = 1.5418 Å; 40
kV and 40 mA) was measured in a 2 Theta range between 5° and 50° with Global Mirror method
(Primary optics: 1.2 mm). The detector is in LYNXEYE ID mode and with a 3 mm slit. Milled
sample mass for XRD measurement was about 1.5 g of total lower stem for each run. Sample
collection was performed for 5 hours total time at 0.0128° step sizes with 10 seconds of exposure

135

at each step. Samples were performed in biological quadruplicate (n = 4). Extraction of cell wall
components were performed using the same methodology as described in the Glycome profiling
results. Relative crystallinity index (RCI) was determined following the peak difference method
and peak area methods outlined [46] using Matlab v2012a (Mathworks, Natick, MA) for peak
fitting with gaussian distributions for crystalline and amorphous peak contributions.
Tensile test
Air dried nine-week-old stem samples were selected from the first internode from the
bottom for tensile testing. Diameters were measured using a digital micrometer at the bottom and
top of each stem sample, giving an overall average. Tensile testing was performed on a United
SFM-20 load frame (United Testing Systems Inc., Huntington Beach, California) using a 9.07-kg
load cell. Samples were immobilized with an opposite twist 2.54 cm grip lined with 0.127 cm of
self-adhesive, 0.127 cm of flexible poly-vinyl carbide, and self-adhesive 320 grit sandpaper, with
a 2.54 cm rip separation. Testing to failure was performed at 0.127 cm/min using Datum 5i
(United Testing Systems Inc., Huntington Beach, California) software to plot force versus
elongation. Samples were performed with 20 biological replicates of stems (n = 20).
Scanning electron microscopy
Fresh nine-week stems were cut with a double-edged razor blade and fixed at 4C for 1-2
hours in 4% glutaraldehyde buffered with 0.1M sodium phosphate at pH 7.4. Following a brief
rinse in the buffer, samples were dehydrated in an ethanol series (25%,50%,75%,95%) for 15
minutes at each gradation and with three 15 minutes changes in 100% ethanol. Samples were
critical point dried in a Leica Microsystems model EM CPD300 critical point dryer (Leica
Microsystems, Vienna, Austria) using carbon dioxide as the transitional fluid. Samples were
mounted on aluminum stubs using high vacuum carbon tabs (SPI Supplies, West Chester, PA),

136

and System Three Quick Cure 5 epoxy glue (System Three Resins, Inc. Auburn, WA). Samples
were coated with osmium (~10 nm thickness) in a NEOC-AT osmium CVD (chemical vapour
deposition) coater (Meiwafosis Co., Ltd., Osaka, Japan). Samples were examined in a JEOL
7500F (field emission emitter) scanning electron microscope (JEOL Ltd., Tokyo, Japan).
Samples were prepared using three biological stems for each sample, and images were acquired
in triplicate (n = 3) at 200x, 14000x and 22000x magnification.
Atomic force microscopy
Air dried nine-week old stem samples were selected from the first internode from the
bottom for analysis. Longitudinal samples were prepared by hand-cutting using a platinum
tipped razor blade in a petri dish, and delignified using a solution of 0.1 N HCl with 10%
NaClO2 at 1% (w/v) biomass loading. Delignification occurred overnight at room temperature.
After delignification, samples were thoroughly washed with double distilled H2O until pH
neutral. Samples were transferred wet to clean mica surface and allowed to air dry at room
temperature.
Image collection was performed using a Bruker Dimension FastScan (Bruker, Camarillo,
CA) equipped with an Acoustic and Vibration Isolation Enclosure, FastScan Scanner, UltraStable High-Resonance Microscope Base, and a Nanoscope V Stage Controller and HV
Amplifier. Scanning was performed using the PeakForce Mapping mode in air, with
SCANASYST-AIR probes (Bruker, Camarillo, CA). Tips were silicon nitride with a nominal tip
radius of 2 nm, a resonant frequency of 70 kHz, and a spring constant of 0.4 N/m, which was
appropriate for the range of modulus tested in this study.
AFM operation and image pre-processing was performed in Nanoscope Analysis V1.5.
All images were taken at 512 x 512 pixels, with four different scan sizes (2.5 Âľm, 1.0 Âľm, 0.5

137

Âľm, and 0.25 Âľm) for each scan area, and at least three different scan areas measured for each
sample (n = 3). All images were fitted with a third order flatten prior to analysis to center data,
remove tilt, and remove bow caused by an uneven cell surface. Image roughness, fibril width
measurements, and height distribution profiles were all calculated in NanoScope Analysis V1.5
software using the Roughness, Section, and Particle Analysis tools respectively.
Microfibril orientation and dispersion were measured using Directionality analysis based
upon local gradient orientation on grayscale images using the Fiji is just ImageJ version of
software (https://fiji.sc/). Nanomechanical properties (elastic modulus, adhesion, deformation,
dissipation) were standardized to the same z-axis properties for direct comparison of image
intensities. Elastic modulus is calculated based upon the Derjaguin-Muller-Toporov model [47].
Deformation represents to maximum penetration depth during tip-cell wall contact, and can be
related to the elastic modulus [48]. The adhesion force represents the absolute value of the
negative force during tip release from the surface [49]. Dissipation is calculated by integrating
the area between the extension and retraction curves, and represents the difference in energy
imparted from the AFM tip to the sample [50].

Results
Irx mutant phenotype, composition, and extractability
The mutants chosen for this study were three xylan irx mutants with T-DNA insertions in
different genes with known specific expression in the secondary cell wall of xylem and
interfascicular fibers. Individual mutants were selected to frame extent of mutants affecting
xylan backbone chain elongation; irx9 was used to represent a xylan synthase mutant with a
strong growth phenotype, while irx10 was used to represent a mutant with a weak phenotype.
Irx15 irx15-L is a proposed methyl transferase that impacts xylan content and structure, as well
as displays similar phenotypes to xylan synthase deficient mutants. Two IRX15-L
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complementation lines of the irx15 irx15-L double mutant were also included to represent a
single mutant irx15 phenotype, which has shown wild type structure in prior work [20].
Complementation expressions had one line with 1-5 times wild type IRX15-L expression level,
irx15 irx15-L IRX15-L (line 1, [20]), and one over-expressing IRX15-L 5-30 times, irx15 irx15-L
OE-IRX15-L (line 6, [20]). All three mutants and complementation line have well-characterized
reductions in xylan backbone chain length (Figure 4.2B) and altered ratios of MeGlcA/ GlcA;
both features which may impact xylan interaction within the secondary cell wall.

Figure 4.1 : Growth phenotypes of xylan irx mutants. Growth phenotypes were documented
at week 6 and 9 as indicated. Arrows in SEM cross-section (lower row, irx9 and irx15 irx15-L)
indicate the irregular xylem (irx) phenotype of collapsed xylem vessels. Scale bars for overhead
growth photos are 1.8 cm, side growth photos are 8 cm, and SEM images are 100Âľm.
Together with the wild type, these five genotypes were grown in parallel in several
batches (Figure 4.1), and examined using SEM for phenotypic analysis. SEM analysis of stem
cross-sections revealed collapsed xylem in irx9, along with significant reduction of secondary
cell wall thickness, corresponding to prior reporting of up to 60% reduction in thickness
139

observed via TEM [11]. Reductions in secondary cell wall thickness were observed to a lesser
extent in irx15 irx15-L, along with normal xylem cell shape and irregular interfascicular cell
surface as reported previously [20]. Irx10 is reported to exhibit minor phenotypic dwarfism and a
moderate irregular xylem phenotype [39] which was not observed in our growth batches. This
discrepancy may be due to IRX10 and IRX10-L acting as functionally redundant genes, with
IRX10-L compensating for a loss of IRX10 and suppressing the phenotype to an extent [15].
Complementation line irx15 irx15-L IRX15-L displayed a reversion to wild type interfascicular
cell shape and size, as did irx15 irx15-L OE-IRX15-L at the SEM scale shown. No noticeable
differences were observed in the cell structure of parenchyma-type secondary cell walls,
indicating localization of mutant phenotype to vascular and interfascicular tissue (Figure 4.12)
as previously reported [19, 20].

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Figure 4.2 : Cell wall polysaccharide composition and xylan chain length. (A) Non-cellulosic
cell wall neutral monosaccharide content of irx mutant stems determined from TFA hydrolysis.
Standard deviations are shown (n = 12, four biological replicates with three technical replicates
for wild type, irx9, irx10, and irx15 irx15-L; n = 6, two biological replicates with three technical
replicates for irx15 irx15-L IRX15-L and irx15 irx15-L IRX15-L). * indicates statistical
significance from wild type (p < 0.05). (B) Xylan degree of polymerization (DP) percent
compared to wild type as reported by prior studies.
Chemical composition analysis was also used to confirm the phenotype (Figure 4.2A),
with reductions in xylan content corresponding to 61% xylan content in irx9 and 85% in irx15
irx15-L compared to wild type, as documented prior in irx9 [11], and irx15 irx15-L [19, 20]. In
irx9, increases in rhamnose, arabinose, mannose and galactose suggest a higher presence of
pectin or arabinogalactopectins (AGP). This could be a consequence of increased ratios of
primary cell wall compared to secondary cell wall, or a compensation mechanism by the plant to
supplement for lost function of xylan. Increased rhamnose and galactose content observed in
irx15 irx15-L likely are due to increased proportion of reducing end tetrasaccharide structure
resulting from shortened xylan chains. Overexpression of IRX15-L in irx15 irx15-L OE-IRX15-L

141

resulted in a 7% increase in xylan content compared to wild type, which has not been reported
previously.
Another characteristic of irx mutants is the altered xylan chain length, or degree of
polymerization (DP) presumably due to altered xylan synthase activity. Xylan DPs have been
reported in individual mutants and complementation lines in previous studies, and were used in
this study based upon prior reporting. Figure2B highlights the percent decrease in chain length
reported in prior literature, along with the literature reference. To supplement visual cell wall
characterizations as well as investigate the pleotropic effects of these mutants, crystalline
cellulose (Figure 4.3A) and lignin contents (Figure 4.3B) were determined. To more
comprehensively document the effect of irx mutants on cell wall deconstruction and
extractability, glycome profiling was performed on irx9, irx10, and irx15 irx15-L (Figure 4.3C).

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Figure 4.3 : Crystalline cellulose, lignin, and glycome profile analysis of cell wall material
of irx mutant stems. (A) chemical quantification of crystalline cellulose content (B) and ABSL
lignin content of cell wall material. Standard deviations are shown (n = 12, four biological
replicates in three technical replicates each for wild type, irx9, irx10, and irx15 irx15-L; n = 6,
two biological replicates with three technical replicates for irx15 irx15-L IRX15-L and irx15
irx15-L OE-IRX15-L). * indicates statistical significance compared to wild type (t-test, p < 0.05).
(C) Cell wall material of wild type, irx9, irx10, and irx15 irx15-L inflorescence stems was
extracted sequentially with increasing chemical severity using glycome profiling. A selected
subset of antibodies and extracts are shown, while the full profile can be found in Figure 4.13
Crystalline cellulose content and lignin content were both significantly lower in irx9
(90% of wild type cellulose and 65% of wild type lignin) compared to wild type, while only
lignin content was moderately lower in irx15 irx15-L (91% of wild type lignin). Decreased lignin
content corresponded to the observed reductions in secondary cell wall thickness, while
reduction crystalline cellulose in irx9 correspond to stunted growth and is not a direct
143

consequence of lost gene function [51]. Irx10 displayed a small, but statistically significant
increase in crystalline cellulose content (105% of wild type), indicating some level of phenotypic
deviation from the wild type.
Glycome profiling was employed on sequentially extracted cell wall material to identify
cell wall polysaccharide structure using a series of carbohydrate-specific antibodies. Glycome
profiling is useful in identifying cell wall polysaccharide deconstruction patterns when subjected
to increasing harsh chemical extraction, and can elucidate features relating to cell wall matrix
organization and integration of cell wall polysaccharides. Glycome profiling results from
carbonate and 1M KOH extracts revealed an overrepresentation of pectin and xylan epitopes in
irx9 and to a lesser extent irx15 irx15-L fractions, while irx10 extractability was nearly identical
to wild type. Specifically, xylan epitope extractability varied between irx9 and irx15 irx15-L,
with a higher presence of mAbs for Xylan4-7 clades in the carbonate fraction of irx9, while only
Xylan-7 clade were significantly higher in the carbonate fraction of irx15 irx15-L compared to
wild type. Extractability of Xylan4-7 clades increased for irx15 irx15-L in the 1M KOH extract
compared to wild type, indicating xylan may be more integrated into the cell wall, requiring
harsher chemical extraction to remove from irx15 irx15-L cell walls compared to irx9, but more
easily extracted compared to wild type or irx10 xylan.
RG-I/AG pectic arabinogalactan epitopes were significantly higher in irx9 compared to
wild type in both carbonate and 1M KOH extracts, while noticeable differences between irx15
irx15-L and wild type pectins were identified only in the 1M KOH fraction. Xyloglucan epitope
recognition was shown to differ only in irx9, and interestingly, the lack of individual xyloglucan
epitope clades in irx9, indicating irx9 most likely has structurally different xyloglucan compared
to the wild type.

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Relative crystallinity index measured by wide angle x-ray scattering
Differences in composition and extractability suggest changes to the cell wall
organization that may result from deficiencies in cell wall components. Examining cellulose
microfibril crystallinity using wide angle x-ray scattering (WAXS) is one non-destructive
technique for probing the structure of cellulose and quantifying the relative crystalline order
within a sample [52], and has been used previously to quantify changes in cellulose crystal order
and size [53].

Figure 4.4 : WAXS diffraction pattern of irx mutant stems and relative crystallinity index.
(A), representative WAXS diffraction patterns of wild type, irx9, irx10, and irx15 irx15-L stems
showing the diffraction peaks for the [110], [1-10], [200], and [400] crystal faces. (B), relative
crystallinity indexes (RCI) of the [200] peak face calculated from the WAXS diffraction pattern,
such as the ones in (A). Standard deviations are shown (n = 4, biological replicates for wild type,
irx9, irx10, and irx15 irx15-L; n = 4, two biological and two technical replicates for irx15 irx15L IRX15-L and irx15 irx15-L OE-IRX15-L). * indicates statistical significance compared to wild
type (t-test, p < 0.05).
WAXS diffraction patterns (Figure 4.4A) show qualitatively distinct tends in irx9 and
irx15 irx15-L compared to wild type, with a clear decrease in the [200] crystal face is observed
relative to the [1-10] and [110] crystal faces, as well as a reduction in prominence of the [1-10]
and [110] crystal faces compared to the amorphous baseline in only irx9. Quantification of
relative crystallinity index (RCI) (Figure 4.4B) by the peak difference method [46] revealed a
lower calculated RCI of the [200] crystal face in irx9 (22% lower) and irx15 irx15-L (9% lower)
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compared to wild type. Reductions in RCI can be attributed to either a reduction in crystalline
chains, or an increase in crystallite size [5, 54, 55]. In the case of irx9, reduced crystalline
cellulose content measured chemically (Figure 4.3A) could account for the reduction in RCI,
however cellulose content does not account for the RCI reduction observed in irx15 irx15-L.
Crystallinity measurements made in other mutant systems have indicated KORRIGAN1
Aspen Homolog PttCel9A1 impacts cellulose crystallinity through either increasing the relative
amount of amorphous cellulose able to interact with hemicelluloses due to cellulose chains under
tension during endo-hydrolysis, or through reduction of crystallite size [54]. Arabidopsis
knockout studies of domains for CESAs such as, rsw2 [56], any1 [57], and CESA1aegeus/
CESA3ixr1-2 [53] have all shown reduced crystallinities attributed to reduced CESA glucan
polymerization velocities, and would be expected to result in deficient cellulose due to
modification of the CESA synthase. Studies examining higher order cellulose crystallinity due to
non-cellulosic cell wall component modification have received significantly less attention, likely
due to the limited knowledge of higher order interactions within the cell wall. Modifications in
poplar to lignin syringyl content showed no changes to cellulose crystallinity, however inducing
the formation of tension wood did significantly increase cellulose crystallinity, with increased
crystallization likely due in part as a stress response [58]. Hemicellulose-cellulose crystallinity
relationships have been observed in bacterial celluloses as mentioned earlier, however to date no
study has observed cellulose crystallinity impacted by xylan within the secondary cell wall, and
have mostly focused on cellulose orientation in primary cell walls due to multiple hemicelluloses
[37, 48].
One of the hypothesized roles of xylan within the cell wall matrix is to act as an
electrostatic spacer, associating via hydrogen bonding along the linear regions of the

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polysaccharide chain with other hydrophilic polysaccharides such as cellulose and other xylans.
Alterations to xylan structure and content may result in changes to inter-hydrogen bonding
between cellulose microfibrils, resulting in more amorphous regions of cellulose compared to
normal xylan. Conversely, sufficient removal of xylan may promote increases in crystallinity due
to an absence of steric hindrance between cellulose microfibrils normally imparted by xylan.
Hemicellulose chain length can impact hemicellulose-cellulose affinity, with shorter xylan
chains have lower equilibrium concentration binding with bacterial celluloses compared to
longer chain xylan [59]. This is important, as shorter xylan chain length is characteristic of irx9
[11] and irx15 irx15-L [20], and may implicate altered xylan-cellulose interactions which
correspond to altered cellulose crystallinity. However, lower xylan DP may only result in noted
changes in cellulose crystallinity if accompanied by reduced xylan content, as no noted
reductions in crystallinity were observed in irx10, or the irx15 irx15-L complementation lines.
For completeness and transparency, relative crystallinity index using the peak area
method was calculated as well, yielding similar conclusions for irx9 and irx15 irx15-L (Figure
4.14), albeit the difficulty of fitting all peaks resulted higher errors associated with the technique.
Relative crystallite size of the [200] face determined using the Sherrer correlation [60] were
found to be around 4.3 nm on average for all samples, which was comparable to prior studies
[61, 62] and indicated no significant change in crystallite size (Figure 4.14C). Similar calculated
crystallite sizes indicate that changes in cellulose were limited to changes in cellulose crystalline
chain abundance rather than crystallite size, which is reasonable given the ability for changes in
hydrogen bonding networks to impact cellulose crystallinity [63].

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Interfascicular ultrastructure is altered in irx mutants
To more thoroughly investigate the ultrastructural organization of the secondary cell wall,
SEM and AFM were performed as complimentary microscopy techniques [64]. SEM was
performed on native, freshly harvested stems that were dehydrated and coated with osmium (~10
nm), providing information on native cell wall shape and surface features, while AFM was
performed on chlorite-delignified, air dried samples to directly examine the cellulose microfibril
layers of the secondary cell wall. Metal coating in SEM likely renders features observed at the
nm scale smaller or unresolved, while AFM requires chemical modification of the surface, but
provides nanoscale resolution. Interfascicular cells were examined as model cells for the impact
of irx mutants on secondary cell wall biosynthesis [65]. Direct comparison of SEM and AFM
images at similar scale are shown in Figure 4.5, highlighting comparable ultrastructural scales
observed by two different techniques.

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Figure 4.5 : Ultrastructure of irx mutants observed with SEM and AFM. Scanning electron
microscopy images of sclerenchyma cell surface texture in native wild type Arabidopsis (A,G,S),
irx9 (B,H,T), irx10 (C,I,U), irx15 irx15-L (D,J,V), irx15 irx15-L IRX15-L (E,K,W), and irx15
irx15-L OE-IRX15-L (F,L,X) stems. Atomic force microscopy images of sclerenchyma cell wall
surface texture after delignification in wild type Arabidopsis (M,Y), irx9 (N,Z), irx10 (O,AA),
irx15 irx15-L (P,BB), irx15 irx15-L IRX15-L (Q,CC), and irx15 irx15-L OE-IRX15-L (R,DD)
stems. Scale bars are 2 Âľm for low magnification SEM (A-F), 1 Âľm for intermediate
magnification SEM (G-L), and 200 nm for high magnification SEM (S-X), 500 nm for the 2.5
Âľm AFM images (M-R), and 200 nm for the 1.0 Âľm AFM images (Y-DD).

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Consistent with previous literature, irregular cell shape was observed in SEM at the
whole cell cross-sections (Figure 4.5A-F) in irx9 [11] and irx15 irx15-L, with a return to wild
type cell shape in irx15 irx15-L IRX15-L [20], while no discernable difference in cell wall
thickness was observed for irx10. Interestingly, the overexpressed complementation line irx15
irx15-L OE-IRX15-L had similar cell thickness to wild type, but exhibited irregular features
similar to irx15 irx15-L. Examination of individual cell wall cross-sections with SEM at the 2.5
Âľm scale (Figure 4.5G-L) revealed normal secondary cell wall and warty layer organization
with smooth cell inner cell wall surfaces in wild type (Figure 4.5G) and irx10 (Figure 4.5I)
only, while rough, non-uniform features were observed in the other irx mutants (Figure
4.5H,J,K,L). These rough features were also accompanied by fibril-like features coating the
inner secondary cell wall and in some cases peeling away from the cell wall, shown
predominantly in irx9 (Figure 4.5T), irx15 irx15-L (Figure 4.5V), irx15 irx15-L OE-IRX15-L
(Figure 4.5X), and to a lesser extent in irx15 irx15-L IRX15-L (Figure 4.5W), indicating an
incomplete phenotypic compliment at the microscale not observed in prior literature.
Interestingly, irx10 also showed slight rough features at the high magnification scale (Figure
4.5U), indicating that all mutants expressed some ultrastructural difference compared to wild
type (Figure 4.5S) at the highest magnification.

Similar to SEM, comparative AFM images revealed nanoscale differences in the cell wall
ultrastructure for all mutants compared to wild type. After chlorite delignification, cellulose
microfibril layers in the secondary cell wall were exposed, revealing cellulose microfibril
organization and orientation with respect to an individual cell wall layer [66]. Qualitatively, at
the 2.5 Âľm scale (Figure 4.5M-R), the cellulose microfibril surface in wild type (Figure 4.5M)

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resembled a smooth sheet, with uniform orientation similar to prior work with maize secondary
cell walls [66]. Contrary to wild type smoothness, large clusters of microfibril bundles were
observed in irx9 and irx15 irx15-L (Figure 4.5N,P), and an uneven smoothness was observed in
the irx15 irx15-L complementation lines (Figure 4.5Q,R). To some degree, the microfibril
organization and uniformity was disrupted in irx10 (Figure 4.5O), however the features were not
as dramatic as the other mutants. At higher resolution, AFM images in irx9 and irx15 irx15-L
(Figure 4.5Z, BB) show the formation of microfibril bundles originating from the microfibril
sheet layer, indicating irregular microfibril features are associated in some part to microfibril
layer synthesis during secondary cell wall thickening, however, may also be imparted by chlorite
delignification inducing aggregation. A summary of qualitative observation, including AFM
image scales not shown in Figure 4.5 are described in Table 4.1. Furthermore, the images in
Figure 4.5 were chosen as representative samples for describing interfascicular cell shape and
AFM surface features, with additional replicates for SEM and AFM images of each sample
found in Figure 4.15-9.

Table 4.1 : Summary features observed in SEM and AFM height images for irx mutants.

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While features at the 2.5 Âľm SEM and AFM images were generally comparative, chlorite
delignification certainly impacts microfibril surfaces observed via AFM to some extent.
However, based upon the stark contrast between wild type and mutants, there is a likely a strong
connection between observed features and native microfibril organization, with chlorite
delignification only partially contributing to observed differences.
Visual inspection of microfibril size at the 2.5 Âľm AFM scale (Figure 4.5M-R) show
clear formations of large macrofibril-like features in irx9 and irx15 irx15-L originating from the
microfibril sheet. To capture the variability in microfibril bundling, fiber widths of well resolved
features were calculated and presented as a distribution (Figure 4.6), similar to prior studies of
primary and secondary cell walls [48, 62]. Example calculation of fibril diameters can be found
in Figure 4.23.

Figure 4.6 : Fiber widths measurements of delignified irx mutant secondary cell walls using
AFM. Fibril widths were determined from well-resolved features in 2.5 Âľm image sizes, using
five measurements for each of three individual images for each sample (n = 15). Averages of
fiber measurements are displayed as a horizontal bar for each sample on figure.
Fibril widths down to individual elementary fibrils were generally not resolved due to the
proximity of fibers within the secondary cell wall as well as the hydration state during sampling
(air dried) impacting elementary fibril size [62], however larger features were captured. From the

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distribution, irx9 and irx15 irx15-L showed similar patterns of macrofibril size distributions, with
some bundles reaching 100 nm in width. Irx10 and the complementation line irx15 irx15-L OEIRX15-L had less distinct bundling, indicating limited aggregation aside from visual microfibril
breaks from the microfibril surface. Irx15 irx15-L IRX15-L showed similar distribution compared
to wild type. Wild type average microfibril size was slightly higher than prior fibril
measurements made on wild type primary cell wall in Arabidopsis [48], however the distribution
was somewhat similar with the exception of single fibril measurements (3-4 nm) not observed in
Figure 4.6.
Roughness analysis was performed as well as microfibril orientation to assess cellulose
microfibril sheet spatial distributions (Figure 4.7). Roughness is a term used to describe
irregularities of surface features from a mean plane, with larger irregularities corresponding to
higher roughness, and is dependent upon both sample scale and measurement size [67].
Microfibril orientation is used in this context to determine directionality of microfibrils within an
image, with dispersion measuring the deviation from the mean angle of orientation.

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Figure 4.7 : Roughness analysis, fiber orientation, and 3D topography measurements of
delignified irx mutant cell walls using AFM. (A), Roughness parameters for average roughness
(Ra) and root-mean-square roughness (Rq) measured on AFM height images at 0.5 Âľm image
scale. P values indicate statistical significance compared to wild type from t-test. (B), Microfibril
orientations measured by dispersion of fibers distributed throughout an entire image at the 1.0
Âľm image scale. Standard deviations for (A) and (B) are shown on figure for three different
images (n = 3). (C), 3D surface profiles of wild type and irx9 demonstrating fiber alignment in
wild type observed at three different image angles compared to irx9. Scale bar represents 100 nm
and image scale is 0.5 Âľm.
The average roughness (Ra) and root-mean-square roughness (Rq) were used to quantify
average image roughness, and deviation from mean roughness respectively [68], with roughness
analysis (Figure 4.7A, full analysis in Table 4.2) indicating to varying degrees of statistical
significance that all mutant microfibril surfaces were rougher compared to the wild type. In
particular, irx9 and irx15 irx15-L grouped similarly, as did the complementation lines, whose
roughness was most likely determined based upon the uneven wavy surfaces (Figure
4.5CC,DD) and not irregular microfibril deviation. Irx10 was also statistically different from the

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wild type, although was less pronounced in roughness and had the lowest deviation from the
mean.
Microfibril orientations were revealed in wild type images, fibril orientation dispersion
was less than 10 degrees for a single microfibril layer, indicating a narrow range of directionality
changes within the microfibril plane. Comparatively, all mutants showed some extent of
dispersion deviating from the wild type, with irx9 showing the largest deviation. Factors
contributing to dispersion were attributed to microfibril bends, breaks, out of plane features [69],
or microfibril weaving between an individual layer which is generally attributed to primary cell
wall tissue [48]. An example plot of the distribution of all microfibril orientations within a single
image can be found in Figure 4.21. A combined visual approach using 3D surface topography
(Figure 4.7C), clearly shows the combination of roughness distribution and microfibril
orientation.
Both the wild type and irx9 clearly show rough features, with the intensity of wild type
appearing more evenly distributed (Figure 4.7C Tilt1). Analysis of the same images at different
orientations (Figure 4.7C Tilt2-3) reveal patterned fiber alignments in the wild type that are
likely not fully captured in the roughness analysis, but are captured in the microfibril orientation
analysis. Likewise, microfibril orientation may not accurately capture fiber orientation in the
presence of wide range of changes in 3D topography, such as in the irx15 irx15-L
complementation lines, highlighted in the height distribution profiles (Figure 4.8).

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Figure 4.8 : Height distribution profiles of delignified irx mutant cell walls using AFM.
Gaussian distributions highlighting the distribution of height topography of each sample at 1 Âľm
scale. Distributions of each image triplicates (n = 3) are shown on figure. Gaussian distributions
of all image scales can be found in Figure 4.22.
Height distributions on an ideal flat surface represent the frequency of particle diameters
[70], however in the case of secondary cell wall surfaces, cellulose microfibril layers may not be
flat and normally distributed. As a result, the height distribution likely represents the range of
observed diameters corresponding to a general understanding of surface flatness, and will be
used in this context to understand variations on microfibril surface topography. From Figure 4.8,
a few conclusions can be drawn about the height profile distributions for each sample. In
correspondence with roughness calculations, irx9 and irx15 irx15-L were similarly grouped, as
were the complementation lines. Wild type and irx10 exhibited somewhat similar Gaussian
distributions, while significant deviations in distribution were shown in the other irx mutants.
Specifically, the complementation lines highlight the heterogeneity in height distributions, which
are likely due to a different mechanism of disorganization causing non-flat height profiles, and
are captured by the other AFM characterization techniques (Figure 4.5Q,R).
Mechanical properties of irx mutants
One of the major functions of the secondary cell wall is to provide mechanical strength to
the individual cell and at the level of tissue and organ of the plant. Irx mutants are characterized

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by mechanical failure at the individual cell level in the form of irregular xylem vessels in the
stem, which directly impact nutrient transport and resistance to osmotic pressures [26, 27]. In
parallel, interfascicular cells which are generally associated with stem strengthening, and from
the prior analysis performed in this study, have documented altered cellulose microfibril
organization at multiple levels. The altered ultrastructure of the secondary cell wall in these cells
are therefore likely to impact physical properties of the stem related to maintaining an erect
position, e.g. bending, breaking, and tensile strength.
Stem tensile strength was measured for lower stems as a metric for overall stem
mechanical strength, with results indicating lower tensile strength in irx9, irx15 irx15-L (p<
0.0005), and to a lesser extent irx15 irx15-L OE-IRX15-L (p<0.1) compared to wild type, shown
in Figure 4.9.

Figure 4.9 : Mechanical strength of xylan irx mutant stems. Tensile stress to breaking of stem
tissue measured by universal tensile test. Box and whiskers plot is the result of twenty replicates
of one biological batch of stems (n = 20). * indicate statistical significance from wild type
(p<0.1), ** indicates statistical significance from wild type (p<0.0005).
From the tensile strength test, results indicate that cell wall structure impacts stem
strength. The role of xylan within the cell wall has not been directly implicated in providing
mechanical strength within the cell wall, with cellulose microfibrils providing a significantly

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higher contribution to cell wall strength [1], however in pulp and paper applications, xylan is a
desirable property in wood pulp for producing paper with higher tensile properties, providing
pulp fracture stiffness [71]. Molecular dynamic simulation work has shown one reason for
improved fiber strength resides in hydrogen bonding between xylan and cellulose stabilizing the
cellulose chain by limiting disruption of hydrogen bonding between cellulose chains [72]. Xylancellulose films have shown improved mechanical properties can result from the incorporation of
cellulose into xylan films, and cited similar conclusions [73].
The severe stem mechanical phenotype observed in irx15 irx15-L was comparable to
mechanical strength observed in irx9, however, significant differences in xylan content between
the two suggests that in the case of mutants exhibiting both reduced xylan content and altered
structure, xylan structure likely plays a stronger role in dictating phenotype. This is supported by
somewhat similar observations made for irx9 and irx15 irx15-L for all techniques, however a
fundamental examination of xylan chain length with respect to cellulose microfibril strength
would need to be explored to confirm this hypothesis.
Altered xylan as observed in the SEM and AFM images does result in irregular cell shape
and at the nanoscale level disruptions in cellulose microfibril organization. Therefore, irregular
cell wall organization observed in irx9 and irx15 irx15-L, and irx15 irx15-L OE-IRX15-L
specifically, and may be the underlying reason for reductions in stem strength. To further probe
the role of cell wall organization in mechanical properties, qualitative nanomechanical mapping
of modulus, deformation, adhesion and dissipation was recorded for each pixel along each of the
topology AFM scans (Figure 4.10), generating spatial distribution maps of each property (full
figure can be found in Figure 4.24). In the context of nanomechanical properties, elastic
modulus is a force based upon contact deformation between the AFM tip and cellulose

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microfibril surface [47], with high modulus regions representing stiff surfaces and corresponding
to lower deformation distances of surfaces. Adhesion force represents the absolute value of the
negative force during tip release from the surface [49]. Dissipation is measured based upon
electrostatic discharge, and may be representative of electrochemical surface heterogeneity [74]
of soft matter.

Figure 4.10 : Merged nanomechanical AFM imaging of irx mutants. Nanomechanical
mapping of modulus (MPa), adhesion (nN), deformation (nm), and dissipation (eV) were scaled
to the same absolute intensities for direct comparison of distributions. Groups were designated
based upon qualitative features, with irx9, irx10, and irx15 irx15-L placed in group 1, and irx15
irx15-L IRX15-L and irx15 irx15-L OE-IRX15-L placed in group 2. Scale bar represents 200 nm,
with all images corresponding to a 1.0 Âľm size.
Uniform features were generally observed in the wild type for all four properties,
suggesting that an organized cellulose-hemicellulose network post chlorite delignification can
induce mechanical uniformity. Nanomechanical images of irx9, irx10, and irx15 irx15-L (Figure
4.10 Group1) exhibited similar uneven distributions, specifically in modulus, adhesion, and
dissipation. Spatial uniformity of adhesion and dissipation mostly restored in the irx15 irx15-L
IRX15-L complementation lines (Figure 4.10 Group2), with uneven modulus and deformation
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distributions likely corresponding to the wavy features observed in the height distribution
profiles (Figure 4.8) limiting AFM tip interaction with the surface. The over-expressed
complementation line (Figure 4.10 Group2) resembled a mix of features between irx15 irx15-L
and the complementation lines, with similar modulus and deformation to the complementation
line, while exhibiting minor heterogeneity in adhesion and dissipation.
Taken together, there is an apparent change in the electrostatic and surface strength
environment of the irx mutants compared to wild type. Specifically, non-uniform modulus and
deformation suggests microfibril defects induce reductions in cellulose microfibril surface
strength, which extrapolated to the macroscale could result in reductions in compressive strength
[26]. Stark contrasts of adhesion and dissipation profiles in the irx mutants also suggests that the
chemical environment of the microfibril surface may also be altered, which could be a function
of cellulose microfibril organization or of post-chlorite delignification hemicellulose localization
along the microfibril surface [48]. In addition, the observation of a nanomechanical phenotype in
irx10 and to a lesser extent in irx15 irx15-L IRX15-L suggests altered xylan structure (DP) can
induce altered xylan-cellulose interactions in the absence of lignin, however lignin likely acts as
a stabilizer and maintains a normal cellulose microfibril structure as observed in the native SEM
images (Figure 4.5) and the absence of a stem mechanical phenotype (Figure 4.9).

Discussion
The secondary cell wall is a complex system, with multiple interdependent components
synthesized and coordinating in unison for precise control of nanoscale architecture promoting
cell wall strength integrity. Xylan is an abundant component found in the secondary cell wall that
plays an important role in promoting normal secondary cell wall organization, however, due to
technological limitations as well as the complexity of cell wall biosynthesis, limited knowledge

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is still known about specific interactions between cell wall components during both primary and
secondary cell wall synthesis [75].
The root cause of irregular organization of the secondary cell wall in this study has been
proposed to be directly linked to alterations in xylan content (Figure 4.2A), as evidenced by
irx9, irx15 irx15-L, and irx15 irx15-L OE-IRX15-L, and to alterations in xylan structure,
specifically xylan backbone DP (Figure 4.2B). In order to understand the impact of altered xylan
on the cell wall, multiple indirect chemical and structural methods were used to elucidate
structural changes imposed by altered xylan. Increased xylan epitopes recognition observed in
the carbonate and 1M KOH extracts of irx9 and irx15 irx15-L via glycome profiling (Figure
4.3C) revealed xylan was not as strongly integrated within the cell wall, and in the case of irx9,
significantly higher pectin epitope recognition suggests pectin content may be increased as a cell
wall compensation mechanism for altered xylan. Normal non-cellulosic carbohydrate extraction
has been observed in irx15 irx15-L IRX15-L, suggesting a similar response to irx10 for the
complementation line [20].
RCIs determined from WAXS (Figure 4.4) provide direct evidence of irregular cellulose
microfibril organization, with lower RCI in irx9 and irx15 irx15-L suggesting less crystalline
cellulose within the wall. Chemical analysis of crystalline cellulose content ruled out reductions
in crystalline cellulose content contributing to a reduced RCI signal in irx15 irx15-L, and
crystallite size ruled out changes in microfibril diameter, indicating that cellulose microfibril
crystallization must be impacted [54]. Xylan chain length has been implicated in xylan
adsorption to cellulose, which could alter the extent of xylan-cellulose hydrogen bonding, and
impact hydrogen bonding between individual cellulose microfibrils [59]. Xylan structure (DP) in
all the mutants observed are different, which implicates that a reduction in xylan content and

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well as structure likely results in changes in cellulose crystallinity, however the specific
mechanism altering cellulose crystallinity can only be speculated with the given results.
Ultrastructural characterization using SEM and AFM revealed overlapping, but distinct
information supporting chemical analysis. In SEM, irx9, irx15 irx15-L, and irx15 irx15-L OEIRX15-L displayed apparent alterations to inner cell wall shape at the whole cell level, while
nuanced differences were observed in irx10 and irx15 irx15-L IRX15-L upon closer inspection.
The severity of cell wall irregularity observed in SEM was roughly correlated to extent of
difference in chemical extractability observed in chemical analysis (Figure 4.3), as well as more
importantly, related to irx mutants containing altered cell wall compositions. AFM images of
delignified secondary cell wall microfibril surfaces showed irregular features in all genotypes,
which implies phenotypes for irx mutants displaying altered xylan structure (irx10, irx15 irx15-L
IRX15-L), but not altered xylan content are suppressed until the cell wall is delignified.
Lignification occurs during SCW synthesis along with xylan deposition and cellulose microfibril
layer synthesis, and with similar transcription factors governing expression of both xylan and
lignin in interfascicular cells during SCW formation [76]. Lignin may play a parallel role to
xylan within the cell wall, supplementing deficiencies in non-cellulosic components to maintain
microfibril organization, which would explain altered lignin mutants also exhibiting irregular
cellulose organization [38].
The complexity of features observed in AFM images (Figure 4.5) made a single metric
to encapsulate microfibril surface irregularities unfeasible, however by measuring cell wall fibril
bundle size (Figure 4.6), roughness (Figure 4.7A,C), microfibril dispersion (Figure 4.7B), and
height topography (Figure 4.8) a few distinctions can be made between genotype microfibril
surfaces. Irx9 and irx15 irx15-L displayed aggregation of microfibrils originating from the

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microfibril surface, into larger macrofibrils, traversing individual microfibril layers. This cell
wall feature cannot be directly confirmed to exist in the native cell wall and may be the result of
delignification and drying inducing aggregation [77], however is likely due to removal of lignin
inducing cellulose aggregation in the absence of sufficient xylan to maintain microfibril
organization. Roughness analysis was likely impacted by the height distribution profiles,
resulting in the complementation lines having the highest roughness values, but were useful in
demonstrating microfibril disorganization impacts surface uniformity in irx9, irx15 irx15-L, and
to a lesser extent irx10. Similar deviations in microfibril orientation between all mutants
compared to wild type does indicate that any change to xylan structure or content results in
similar loss of orientation.
In addition to characterizing fundamental xylan-cellulose interaction, another major
outcome of this study is the observation of previously invisible phenotypes using SEM and AFM
to examine 3D topography. Identification of precise gene functions has been accelerated by the
parallel development of analytical techniques to analyze cell wall structure [78], such as 3D
electron tomography [79] and neutron scattering [80]. SEM and TEM are standard microscopy
techniques for nanoscale characterization of plants [81], however TEM is usually employed to
view changes within cell wall layers. From this work, we show that SEM can provide unique
information on the integration of cell wall structures to observe phenotypes not obtainable using
standard TEM alone, and propose broadening the application of SEM for plant cell wall
phenotyping. AFM provides a broad range of tools to quantitatively characterize the cell wall
landscape to supplement the observations from SEM, as well as provides the ability to probe
nanomechanical properties of the cell wall surface. This is important and novel, as these
microscopy in conjunction with one another, provide a route of direct measurements towards

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connecting the role of nanoscale cell wall organization in macroscopic phenotypes observed
(Figure 4.11).

Figure 4.11 : Schematic of analytical techniques utilized in this study, and proposed
connections between individual techniques.

The loss of Arabidopsis stem mechanical integrity observed in irx9, irx15 irx15-L and
irx15 irx15-L OE-IRX15-L (Figure 4.9) was comparable to similar mechanical phenotypes
observed in xylan modified irx9 mutants of rice properties [82], and of IRX8 homologs in poplar
[83]. Mechanical phenotypes are not limited to xylan-deficient mutants, with mechanical
deficiencies in cellulose deficient brittle-culm1 [84] rice mutants that functions to crystallize
cellulose [85]. Lignin deficient mutants such as irx4 in Arabidopsis exhibit mechanical
phenotypes [86], however reductions in other cell wall components convolute individual
contributions to whole stem mechanical phenotypes. As an extended application of the results
and techniques presented here, analysis of other Arabidopsis mutants may yield novel
information relating to understanding cell wall polymer interactions. Specifically, investigation
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of cellulose, and lignin deficient mutants, or combination mutants could help understand the role
of lignin in maintaining normal cell wall structure with or without altered xylan structure.
Additional invisible phenotypes, such as altered xylan side chain substitution in the case of the
gux mutants [23] or glucuronic acid methylation gxmt mutants [25] may be elucidated, and
demonstrate the dynamic nature of the secondary cell wall in maintaining normal cell wall
structure.
In summary, AFM height profiles and nanomechanical mapping demonstrate that cell
wall topographical and organizational heterogeneity impact nanoscale mechanical properties, and
together with SEM, tensile strength measurements, chemical analysis, and WAXS crystallinity
measurements provide significant evidence that altered cell wall polymer abundance and
structure can significantly impact cell wall organization at the nano and macroscale.

Conclusions
The secondary cell wall is a precise biological matrix that imparts cell wall rigidity, with
loss of a single component resulting in dramatic phenotypic changes. This study investigated the
effect of xylan deficient mutants in Arabidopsis on secondary cell wall organization to
understand the role xylan plays in normal development. Reductions in relative crystallinity from
WAXS diffraction patterns, coupled to increased extractability within the cell wall of irx9 and
irx15 irx15-L suggest altered hydrogen bonding between xylan and cellulose, and reduced cell
wall integration of xylan. Nanoscale cell wall organization was impacted to some degree in all
genotypes tested, and demonstrated altered xylan structure resulted in phenotypic changes that
may have been masked by other cell wall components such as lignin.
Quantification of microfibril surface roughness, heterogeneity and orientation observed in
AFM suggest normal cellulose deposition and organization is dependent in part upon noncovalent interactions with non-cellulosic polymers in the cell wall matrix. Cell wall defects

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corresponded to nanomechanical heterogeneity in microfibril surface strength and chemical
environment, with parallels between nanoscale and macroscale mechanical deficiencies in irx9,
irx15 irx15-L, and irx15 irx15-L OE-IRX15-L displaying both altered xylan composition and
structure. The analytical techniques presented demonstrate novel ways of characterizing invisible
phenotypes to better understand the role of gene function impacting cell wall structure. Overall,
irregular cell wall features observed from the irx mutants in this study are likely attributed to
weakened cellulose-hemicellulose secondary cell wall matrix interfaces due to altered xylan
content and structure, and highlight the fine-tuned complexity of the plant cell wall.

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APPENDICES

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APPENDIX C: Supplementary figures

Figure 4.12 : SEM images of parenchyma cells surface texture in irx mutants. Wild type
(A,G,M), irx9 (B,H,N), irx10 (C,I,O), irx15 irx15-L (D,J,P), irx15 irx15-L IRX15-L (E,K,Q),
and irx15 irx15-L OE-IRX15-L (F,L,R) stems labeled. Scale bars are 2 Âľm for low magnification
(A-E), 1 Âľm for intermediate magnification (G-L), and 200 nm for high magnification (M-R).

168

Figure 4.13 : Extended dataset of the glycome profile of irx mutants. Glycome profile of
Arabidopsis cell wall material extracted sequentially with increasing chemical severity using
oxalate, carbonate, 1M KOH, 4M KOH, acidic chlorite, and 4M KOHPC pretreatments as
defined in materials and methods. Antibody binding strength is based upon optical density, with
binding intensity presented as a color gradient from black (low binding) to red (intermediate
binding), and yellow (strongest binding). Green bars on the top indicate the amount of
carbohydrate recovered per gram of cell wall AIR for each extraction.

169

Figure 4.14 : WAXS diffraction curve processing for crystallinity and microfibril diameter
calculation. (A), WAXS diffraction curve fitting for quantification of relative crystallinity index,
with ten Gaussian fits used to fit the major crystal faces and account for amorphous
contributions. (B), Relative crystallinity index determined using the peak area method as
described in the methods. (C), Sherrer microfibril diameter determined using the width at peak
half height of the [200] crystal face.

170

Figure 4.15 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis wild type. Scale bars for SEM are 2 Âľm for low magnification (left column), 1 Âľm
for intermediate magnification (middle column), and 200 nm for high magnification (right
column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the
1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the
0.25 Âľm images (fourth column).

Figure 4.16 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx9. Scale bars for SEM are 2 Âľm for low magnification (left column), 1 Âľm for
intermediate magnification (middle column), and 200 nm for high magnification (right column).
Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the 1.0 Âľm
images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the 0.25
Âľm images (fourth column).

171

Figure 4.17 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx10. Scale bars for SEM are 2 Âľm for low magnification (left column), 1 Âľm for
intermediate magnification (middle column), and 200 nm for high magnification (right column).
Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the 1.0 Âľm
images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the 0.25
Âľm images (fourth column).

Figure 4.18 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx15 irx15-L. Scale bars for SEM are 2 Âľm for low magnification (left column), 1
Âľm for intermediate magnification (middle column), and 200 nm for high magnification (right
column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first column), 200 nm for the
1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third column), and 50 nm for the
0.25 Âľm images (fourth column).

172

Figure 4.19 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx15 irx15-L IRX15-L. Scale bars for SEM are 2 Âľm for low magnification (left
column), 1 Âľm for intermediate magnification (middle column), and 200 nm for high
magnification (right column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first
column), 200 nm for the 1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third
column), and 50 nm for the 0.25 Âľm images (fourth column).

Figure 4.20 : Additional SEM and AFM images of sclerenchyma cell wall surface texture in
Arabidopsis irx15 irx15-L OE-IRX15-L. Scale bars for SEM are 2 Âľm for low magnification
(left column), 1 Âľm for intermediate magnification (middle column), and 200 nm for high
magnification (right column). Scale bars for AFM are 500 nm for the 2.5 Âľm images (first
column), 200 nm for the 1.0 Âľm images (second column),100 nm for the 0.5 Âľm images (third
column), and 50 nm for the 0.25 Âľm images (fourth column).

173

Table 4.2 : Roughness analysis and fiber width measurements of delignified irx mutant cell
walls using AFM. Roughness parameters for average roughness (Ra) and root-mean-square
roughness (Rq) measured on AFM height images at multiple image scales. Roughness is utilized
in this case to measure cell wall uniformity. Roughness calculations were performed on three
individual images for each scale (n = 3), with standard deviations shown in table. P values
indicate statistical significance compared to wild type from t-test.

Figure 4.21 : Distribution profiles of microfibril orientations for an entire image. Sample
image distribution profiles for each irx mutant and wild type. Dispersion was calculated based
upon the width of the gaussian distribution.

174

Figure 4.22 : Height distribution profiles of delignified irx mutant cell walls using AFM.
Gaussian distributions of height ranges at each image scale are displayed as individual plots of
three images for each sample.

175

Figure 4.23 : Example calculation of microfibril width. Microfibril widths were calculated
using the build in section Tool in Nanoscope. Well resolved microfibril features were used to
determine horizontal fibril widths.

176

Figure 4.24 : Merged nanomechanical AFM imaging of all irx mutants. Nanomechanical
mapping of modulus (MPa), adhesion (nN), deformation (nm), and dissipation (eV) were scaled
to the same absolute intensities for direct comparison of distributions. Scale bar represents 200
nm, with all images corresponding to a 1.0 Âľm size.

177

APPENDIX D: Additional irx experiments

The following experiments were performed alongside other irx work, and were determined to not
fit in the context of Chapter 4.
Additional methods
Water retention value
Water retention value (WRV) was determined according to a modified version of TAPPI
UM 256, as previously described [87], with minor modifications. Briefly, 0.5g of milled AIR
stems were soaked in 150ml of de-ionized water for 1 hour. After soaking for 1 hour, samples
were rinsed with around 200 mL of de-ionized water and vacuum filtered using a 200 mesh
Buchner filter. Washed biomass was inserted into a spin-column (Handee Spin Column Cs4,
Thermo Scientific) with a 200 mesh stainless steel membrane, and centrifuged at 4000 x g for 15
minutes. Drained biomass was weighed on pre-massed aluminum trays, and dried in an oven at
105°C for at least 3 hours, then weighed again. The WRV is the ratio of the mass of water
remaining in the biomass after centrifuging divided by the mass of dry biomass. Sequential
extraction was performed using the same methodology as Glycome profiling. Water retention
value was performed in technical triplicate of biological duplicates (n = 6) for each extraction.
Enzymatic hydrolysis
Enzymatic hydrolysis was performed at 1.0% solids loading (g AIR biomass/g solution)
in 1.5 mL Posi-Click Tubes (Denville Scientific, Holliston, MA). Enzyme solutions were made
at 30 mg protein/g glucan content using CTec3 (Novozymes A/S. BagsvĂŚrd, Denmark) and
HTec3 (Novozymes A/S. BagsvĂŚrd, Denmark), with a ratio of 2:1 CTec3:HTec3. A buffer
solution of 50 mM citric acid (pH 5.20) was used to maintain pH, and samples were incubated at
50°C with horizontal mixing at 180 rpm for 48 hr. Samples were centrifuged at 13000 x g for 3
min post-incubation and filtered using 22 Âľm mixed cellulose-ester filters (EMD Millipore,

178

Billerica, MA). Samples were quantified on the HPLC (Agilent 1100 Series) with an Aminex
HPX-87H column (Bio-Rad, Hercules, CA, USA) with a mobile phase of 5 mM H2SO4. Glucose
yield was determined based upon quantified glucose observed compared to total AIR cell wall
glucan available (as glucose) including non-cellulosic glucan, while xylose yield was determined
based upon total AIR cell wall xylan available (as xylose). Samples were performed in technical
triplicate of biological duplicates (n = 6).
Additional results
Cell wall deconstruction measured by water retention value
Water retention value is metric that can be used quantify cell wall hydrophilicity, is often
correlated with cell wall digestibility, and is dependent upon a cell wall composition as well as
structure [87]. In the case of irx mutants, hydrophobic xylan, lignin and hydrophilic pectin
content all contribute towards a sample bulk hydrophilicity. Sequential extraction can alter the
impact of individual components and elucidate how residual cell wall components interact with
an external variable. Water retention value was determined for initial AIR cell wall material, and
after two increasingly harsher chemical extractions (Fig. 4A), along with chemical composition
quantification of residue after all sequential extractions (EIC) (Fig. 4B-D).

179

Figure 4.25 : Water retention values and extractive insoluble residue of cell wall material of
irx mutants. (A), Water retention values (WRV) of alcohol insoluble residue (AIR) with and
without chemical extraction with 1M KOH and extractive insoluble components (EIC) represents
solid residue after sequential extraction with 4M KOH after chlorite delignification. (B-D),
Composition of neutral monosaccharides, crystalline cellulose, and ABSL lignin content of
residue remaining after 4M KOHPC extraction. Standard deviations are shown with two
biological replicates with three technical replicates (n = 6).

Initial WRV of AIR samples showed higher values for irx9 and irx15 irx15-L, which
were most likely due to the decreased xylan (Figure 4.2) and lignin content (Figure 4.3). After
1M KOH, glycome profiles indicated similar extractions in the 4M KOH fraction for all but irx9,
meaning that differences in lignin content most likely were the dominant composition variable
dictating water retention, and the lower lignin in irx9 and irx15 irx15-L resulted in higher WRV.
WRV of the EIC residue resulted in a different trend however, with the complementation lines
exhibiting increased WRV compared to wild type. Composition profiles (Figure 4.25B-D)
indicated that the predominant difference between components in EIC is the residual xylan (0.3%
total mass) and differences in lignin content. The likely explanation for the increases in water

180

retention observed after extractions in the complementation lines arises from the lower lignin
content relative to wild type. This result indicates that the complementation lines do not fully
revert to wild type, displaying similar characteristics to irx15 irx15-L after harsh extraction and
may have subtle phenotypic differences compared to wild type that were not observed in initial
phenotyping.
Enzymatic hydrolysis monomer sugar yields
After extensive investigation of the secondary cell wall ultrastructure and physical
properties, enzymatic hydrolysis was performed to relate structural and compositional features to
cell wall deconstruction.

Figure 4.26 : Enzymatic digestibility of cell wall material of irx mutant stems. Enzymatic
hydrolysis monomeric sugar yields of glucose (A) and xylose (B) of unpretreated AIR on a basis
of % monomeric sugar observed per total available polysaccharide (g/g*100). Standard
deviations are shown (n = 6, biological duplicate with technical triplicate).

From the time course enzymatic hydrolysis results (Figure 4.26), glucose and xylose are
more easily released for all the genotypes. Sugar yields irx9 [51], irx15 irx15-L, and irx15 irx15L IRX15-L [15] roughly correlate with prior studies utilizing pretreatments before hydrolysis,
indicating increases in hydrolysis yields can be extrapolated outside of no pretreatment only.
While the observed trend of glucose yields makes it hard to pinpoint the specific key contributor
181

to the improved yields, modifying xylan biosynthesis readily affects glucose yields in the
positive direction. This compositional alteration, coupled with the observed mechanical changes
argues for altered xylan plants as a viable route to improve feedstocks for biofuels production.
Understanding changes in cellulose RCI though alteration of xylan content
To further probe the hypothesis of xylan content impacting cellulose microfibril
crystallinity and organization, sequential extraction was performed on wild type (Fig. 5) to
observe composition and RCI changes in response to chemical extractions.

Figure 4.27 : Cell wall compositions of Arabidopsis wild type and WAXS relative
crystallinity index after chemical extraction. (A) cell wall neutral monosaccharide content
determined from TFA hydrolysis, (B) crystalline cellulose content, and (C) ABSL lignin content
of wild type AIR, and sequentially extracted with carbonate and 1M KOH. (D) Relative
crystallinity indexes of the [200] peak face calculated from the WAXS diffraction patterns in
wild type native, AIR, and sequentially extracted with carbonate and 1M KOH. Standard
deviations are shown (n = 3, biological replicates. * indicates statistical significance compared to
wild type (t-test, p < 0.05).
From the composition results (Figure 4.27A-C), carbonate (50 mM) removed
hemicellulose and pectin components, with decreases in xylose (78% of AIR), rhamnose, and
galactose content resulting in a small but statistical mass fraction increase in crystalline cellulose

182

content (7% increase from AIR). The 1M KOH extract was a significantly higher in
concentration, and after pectin removal in the carbonate fraction, predominately resulted in the
removal of xylans (xylose content was 47% of AIR) and xyloglucans, as observed in the
composition profile (Figure 4.27A) and glycome profiling results (Figure 4.13), with an
increase in the proportion of crystalline cellulose remaining (Figure 4.27B) in the residue (25%
increase from AIR). From these chemical changes, WAXS was performed and RCIs were
calculated, with a decrease in RCI after carbonate extraction, followed by a significant increase
in RCI after 1M KOH extraction (Figure 4.27D). AIR RCI values were shown to be statistically
equivalent to the native RCI, demonstrating that removal of non-structural components did not
impact RCI.
While not definitive, changes in relative crystallinity observed following carbonate and
1M KOH extraction may be attributed to changes in cellulose microfibril hydrogen bonding
within the cell wall. Specifically, chemical extraction would result in the removal of less
structurally integrated hemicelluloses first, with the remaining hemicelluloses experiencing an
altered chemical environment, as well as potentially altered structures due to chemical cleavage
of some bonds. One potential outcome of this would be increased xylan-cellulose hydrogen
bonding, resulting in lower observed crystallinities. Sufficient removal of non-cellulosic
components (1M KOH) would result in the opposite observation, with increased inter-cellulose
hydrogen bonding in the absence of spacer components. In more severe extractions such as 1M
KOH however, RCI gets convoluted as cellulose constitutes a larger fraction of the total residue,
which would also contribute to the proportional increase in observed RCI.

183

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