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This is to certify that the
dissertation entitled

PRODUCTION AND ANALYSIS OF BlOLOGICALLY—ACTIVE
CELLULASES FOR ETHANOL FUEL IN MAIZE BIOMASS

presented by

Callista B. Ransom

has been accepted towards fulfillment
of the requirements for the

Ph.D. degree in PLANT BREEDING AND
GENETICS

 

 

 

mortar” M

' Major Professor’s Signature
ll —~ 0 01 »— 0 Q?

Date

 

MSU is an afflnnative-action, equal-opportunity employer

PRODUCTION AND ANALYSIS OF BIOLOGICALLY-ACTIVE CELLULASES FOR
ETHANOL FUEL IN MAIZE BIOMASS

By

Callista B. Ransom

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

PLANT BREEDING AND GENETICS

2007

ABSTRACT

PRODUCTION AND ANALYSIS OF BIOLOGICALLY-ACTIVE CELLULASES FOR
ETHANOL FUEL IN MAIZE BIOMASS

By
Callista B. Ransom
The US needs a competitive substitute for fossil fuels; ethanol biofuel is an
attractive choice in a suite of alternatives. Production of ethanol involves fermentation of
sugars, which can come from plants as simple sugars, starches, or complex structural
polysaccharides of the plant cell wall.

Ethanol production from plant biomass requires pretreatment to disrupt the lignin;
addition of the hydrolysis enzymes to the pretreated feedstock; and fermentation of the
resulting sugars to ethanol, which must then be distilled. The enzymes necessary for cell
wall degradation include cellulases (endo- and exo-glucanases and B-glucosidases), and
hemicellulases (most importantly xylanases). Roadblocks stand in the way of this
technology becoming mature and economically feasible, including the high costs of
enzymes and pretreatment.

One possible solution to this problem is to use crops as enzyme biofactories. In
this work, the cellulases endoglucanase El from Acidothennus cellulolyticus and B-
glucosidase (BG) from Butyrivibriofibrisolvens Hl7c were produced in transgenic maize
and were shown to be enzymatically active and accumulated 0.01% to 1.16% and
approximately 0.15% to 3.11% of plant total soluble protein, respectively. These
enzymes were also able individually to convert cellulose (E1) or cellobiose (BG) to
fermentable sugars, and also biomass (AF EX-n'eated corn stover) when supplemented

with commercial enzyme. In addition, unsupplemented combinations of plant-produced

cellulases successfully converted AFEX-treated corn stover to fermentable sugars. Thus,
production of hydrolysis enzymes in crop plants is feasible and further developments will

make it even more attractive.

© COPYRIGHT BY
CALLISTA B. RANSOM
2007

DEDICATION

I dedicate this work to God, my husband Aous Abdo, and my parents, Carol and Rodger
Ransom.

ACKNOWLEDGEMENTS

I would like to first of all thank God for making this experience possible for me
and for giving me the patience and perseverance to finish.

I thank my parents, Carol and Rodger Ransom, for always believing in me and
encouraging me to succeed, and for always supporting me. I thank them for being there to
comfort me during the bad times and cheer me on during the good ones. I am who I am
today because of them. I thank my husband, Aous Abdo, for his continuous
encouragement and support. His constant nurture and inspiration gave me hope, courage
and determination and helped me survive grad school intact and emerge as a better
person. Without my family, this degree would not have been possible.

I would like to thank my committee members for their guidance. I thank Dr.
Sticklen for providing me with the opportunity to do research in her lab and the
experience she provided me with. I thank Dr. Freed for his career advice and other
counsel. I thank Dr. Sink for his knowledge and the use of his lab and equipment. I thank
Dr. Hancock for his advice and support. I thank Dr. Dale for his expertise and the
opportunity to utilize his lab and resources, and for taking over for Dr. Sink. I thank them
all for believing in me.

I thank my colleagues in the Sticklen lab for their help and support: Anwaar
Ahmad, F arzaneh Teymouri, Hesham Oraby, Hassan Salehi, Rashid Ahmad, Chuansheng
Mei, Jason Miller, Chunfang Qi, Sarah Karinen, Abby Vannortwick, Roxy Nichols,

Yanfen Zhai, Kingdom Kwapata, Sang Park and Robab Sabzikar. I will miss them.

vi

I thank Dr. Renate Snider for helping me with my writing and giving me
confidence and encouragement. I thank Dr. Jennifer Grzegorek for her support. I thank
Dr. Sasha Kravchenko for her advice and help with the statistical analyses.

I thank Dr. Kelly for allowing me to use his lab’s equipment, and the personnel in
his lab for their friendship and support: Halima Awale, Karolyn Terpstra and Evan
Wright.

I thank Dr. Steve Van Nocker for allowing me to work and learn in his lab for
over a year; this experience allowed me to begin molecular work promptly in the Sticklen
lab. Also, I thank my colleagues there for their help and knowledge: Hua Zhang, Sang-
Dong Yoo, Philip Ludwig and Lingxia Sun.

I thank my friends and colleagues in the Departments of Crop and Soil Science
and Horticulture and the Plant Breeding and Genetics Program that I have not previously
mentioned: Veronica Vallejo, Ann Armenia, Cholani Weebadee, Dr. Karim Maredia, Dr.
Rebecca Grumet, Dr. Barb Sears, Dr. Eunice Foster, Dr. James Kells, Dr. Taylor
Johnston, Kaori Ando, Alejandra Ferenczi, Sue Hammar, Chrislyn Particka, Nat Hauck,
Clarice Mensah, Guozhu Tang, Pete Callow, Aubrey Sebolt, Janet Lewis, Jim Olmstead,
Guo-Qing Song, Donna Ellis, Darlene Johnson, Gina Centano, Sandie Litchfield, Debbie
Williams, Lyn Scrimger, Rita House, and Cal Bricker.

Last, but not least, I thank my colleagues in Dr. Dale’s lab for their expertise, help

and patience: Holly Gunter, Dr. Venkatesh Balan and Shishir Chundawat.

vii

TABLE OF CONTENTS

LIST OF TABLES ......................................................................................................... xi
LIST OF FIGURES ....................................................................................................... xii
KEY TO SYMBOLS AND ABBREVIATIONS ........................................................ xviii
1. LITERATURE REVIEW ............................................................................................. l
1. Introduction ............................................................................................................. l

2. The Plant Cell Wall ................................................................................................. 2
2.1 Cell wall components ......................................................................................... 3

2.2 Two major types of primary cell wall ................................................................. 5

3. Cell wall degradation ............................................................................................... 6
3.1 Microorganisms ................................................................................................. 6

3.2 Hydrolysis ......................................................................................................... 7

4. Ethanol production .................................................................................................. 9
4.1 Maize grain ethanol production .......................................................................... 9

4.2 The promise of cellulosic ethanol ..................................................................... 10

5. Production of Hydrolysis Enzymes in Biomass Crops ............................................ 13
5.1 Plants as molecular biofactories ....................................................................... 13

5.2 Successful plant-produced. hydrolysis enzymes ................................................ 14

5.3 TSP must be extracted prior to pretreatment ..................................................... 15

5.4 Thermostable enzymes are desirable ................................................................ 16

5.5 Subcellular targeting and sequestration ............................................................ l6

6. Other Approaches .................................................................................................. 19
6.1 Microbial engineering ...................................................................................... 19

6.2 Lignin pathway manipulation ........................................................................... 20

6.3 Up-regulation of cellulose pathway genes to increase sugar content ................. 21

6.4 Delayed flowering to increase biomass ............................................................ 22

6.5 Genetic manipulation to increase biomass ........................................................ 22

7. Conclusion ............................................................................................................ 23
11. MATERIALS AND METHODS .............................................................................. 26
1. Transformation Vectors ......................................................................................... 26
1.1 Genes of Interest .............................................................................................. 26

1.2 Selectable Markers ........................................................................................... 27

2. Maize Transformation, Acclimation and Care ........................................................ 29
3. DNA Analyses ....................................................................................................... 30
3.1 Extraction ........................................................................................................ 30

3.2 PCR ............ , ..................................................................................................... 30

. 3.3 Southern blots .................................................................................................. 31

4. RNA extraction ...................................................................................................... 32
5. Northern blots ........................................................................................................ 32

viii

6. Labeling, hybridization and detection for Southern and northern blots ................... 32

7. Extraction of TSP .................................................................................................. 33

8. Activity Assays ...................................................................................................... 34
8.1 MUCase Activity Assay for El ........................................................................ 34

8.2 IUPAC Assay for BG ....................................................................................... 34

8.3 p-Nitrophenol (pNP) Assay for BG .................................................................. 35

8.4 DNS Assay for Reducing Sugars ...................................................................... 35

8.5 Glucose analyzer ............ . .................................................................................. 36

9. Western Analysis ................................................................................................... 36
9.1 General Procedure ............................................................................................ 36

9.2 Specific conditions ........................................................................................... 36

10. Pretreatment of Biomass ...................................................................................... 37
11. Conversion Analyses ........................................................................................... 37
11.1 E1 .................................................................................................................. 37

11.2 Microplate Hydrolysis .................................................................................... 38

12. Progeny Analyses ................................................................................................ 39
III. RESULTS. FOR El .................................................................................................. 4O
1. Transformation ...................................................................................................... 40

2. Molecular and enzymatic analyses ......................................................................... 40

3. Conversion analyses .............................................................................................. 42
.4. Second generation ................................................................................................. 43
IV. RESULTS FOR BG ................................................................................................. 46
1. Maize plants are transformed with BG and FLC ..................................................... 46

2. Maize plants are transgenic and express BG ........................................................... 47

3. TSP from transgenic BG plants converts cellobiose to glucose ............................... 55
V. RESULTS OF EXPERIMENTS RELATED TO OPTIMIZING RATIOS OF PLANT-
PRODUCED HYDROLYSIS ENZYMES FOR CONVERSION .................................. 61
1. Introduction. .......................................................................................................... 61

2. Results and Discussion .......................................................................................... 62
2.1. Preliminary Results ......................................................................................... 62

2.2. Optimal concentrations of transgenic plant TSP .............................................. 69

VI. DISCUSSION ......................................................................................................... 84
l. Plant-produced El and BG .................................................................................. 84

2. Limitations/problems ............................................................................................. 87
2.1 Problems Related to Molecular Analyses ......................................................... 87

2.2 E] Enzyme Activity Assays ............................................................................. 90

2.3 Problems Related to Materials and Recordkeeping ........................................... 91

3. Conclusions ........................................................................................................... 93
APPENDIX A. RESULTS FOR FLC ............................................................................ 94
1. Introduction ........................................................................................................... 94

2. Materials and Methods ................... 94

ix

3. Results and Discussion .......................................................................................... 95

APPENDIX B. XYLANASE TRANSFORMATION OF MAIZE (ALONG WITH FLC),
ANALYSES OF TOBACCO AND MAIZE TRANSFORMED WITH XYLI, AND

CLONING OF XYLI AND A NEW XYL ....................................................................... 99
1. Introduction ........................................................................................................... 99

2. Materials and Methods ........................................................................................... 99

3. Results and Discussion ........................................................................................ 101
3.1 Transformation of maize with XY L1 and F LC ............................................... 101

3.2 Molecular and enzyme assays for XYL on maize and tobacco ........................ 102

3.3 Cloning of new XYL ..................................................................................... 109
APPENDIX C. CBHI AND FLC IN MAIZE .............................................................. 111
1. Introduction ......................................................................................................... 111

2. Materials and Methods ......................................................................................... 111

3. Results and Discussion ........................................................................................ 112
3.1 Transformation of maize with Syn-CBHl and FLC ........................................ 112

3.2 Molecular analyses for Syn-CBHI on maize .................................................. 112
REFERENCES ............................................................................................................ 117

LIST OF TABLES

Table 1. Subcellular targeting of various hydrolysis enzymes. ....................................... 18

Table 2. Mean enzymatic activity and percentage E1 of total plant soluble proteins

produced by transgenic maize plants (To). N=number of replicates; S.D.=standard
deviation. ...................................................................................................................... 41

Table 3. Percentage of E1 in second generation (T1) plants as determined by activity
assay. The first four columns show the parents and % E1, while the last two columns
show the same information for the second generation. 1-15: Second generation E1 plants;
- : Activity showed less than negative (nontransgenic) control; HNPC: Nontransgenic
variety ‘Honey ‘11 Pearl’ negative control; N/A: Not applicable; * PCR negative for

pMZ766-E1CAT ............................................................................................................ 45

Table 4. Transformation events and regeneration of To maize plants co-transformed with
a combination of pUC18 13 (containing BG) and pDM302 (event 11 only; containing bar)
or pGreen (all other events; containing FLC and bar). ................................................... 47

Table 5. Plant lines (To) and apparent copy numbers based on hands on Southern blots. 55

Table 6. Glucose (g/L) released from 0.015M cellobiose after 30 minutes using 100 pg
maize TSP from plants transformed with 30. Hill: Untransformed maize negative
control. The difference between 0 and 30 min was not significant at a=0.05 (t=-2.9l7,

df=12). .......................................................................................................................... 56

Table 7. Glucose (g/L) released from 1% cellobiose at 0 min, 1 h, 7.25 h and 26.5 h,
using 570 pl maize TSP from plants expressing 36. Data have been adjusted by
subtracting the substrate blank. Hill: Untransformed maize negative control. The
differences between 0 min and 1 h, 0 min and 7 .25 h, and 0 min and 26.5 h were all
significant at a=0.05 (t=2.667, 6.733 and 8.338 respectively, df=6) ............................... 57

Table 8. Mean activity in units pNP (pNPU) and estimated %BG in transgenic plant TSP.
=3. ............................................................................................................................... 60

Table 9. Maize plants tested for XYL activity using PABAH assay for reducing sugars.
Absorbance taken at 410 nm for 0 min, 30 min and 4 h are shown, along with the
adjusted values (0 min subtracted). .............................................................................. 108

Table 10. Tobacco plants total protein concentrate and extracellular fluid wash tested for
XYL activity using PABAH assay for reducing sugars. Absorbance taken at 410 nm for 0

min, 30 min and overnight are shown, along with the adjusted values (0 min subtracted).
.................................................................................................................................... 108

xi

LIST OF FIGURES

Figure 1. pMZ766-E1CAT. CaMV 358: Cauliflower Mosaic Virus (CaMV) 35S
promoter; 0: tobacco mosaic virus (TMV) translational enhancer; PrlaSP: tobacco
pathogenesis-related protein (apoplast targeting signal); E 1 -cat: coding sequence of the
catalytic domain of endo- 1 ,4-B-gluccanase E1 from A. cellulolyticus; nos:
polyadenylation signal from the nopaline synthase gene. ............................................... 26

Figure 2. pUC1813. CaMV 358: Cauliflower Mosaic Virus (CaMV) 35S promoter; ER:
endoplasmic-reticulum leading sequence; bglA: gene encoding B-glucosidase; VT:
vacuole-targeting sequence; 35S-t: CaMV 35S terminator. ............................................ 27

Figure 3. pBY520 (Xu et al. 1996). Actl-S’: rice actin 5’ region (promoter); HVAI :
barley H VAI gene; pinII: potato proteinase inhibitor 11 terminator; 358 5’: Cauliflower
Mosaic Virus 35S promoter; bar: bar gene encoding Bialaphos herbicide resistance; nos-
3’: nos terminator. ......................................................................................................... 27

Figure 4. pDM302. Act1-5’: rice actin (Actl-5’) 5’ region (promoter); bar: bar gene
conferring Bialaphos herbicide resistance; nos: nos terminator. ..................................... 28

Figure 5. pGreen. LB: T—DNA left border; 35S: Cauliflower Mosaic Virus (CaMV) 358
promoter; bar". bar gene conferring resistance to Bialaphos; nos: nos terminator; FLC:
FLOWERING LOCUS C (FLC) gene; RB: T-DNA right border; nptII: gene conferring
bacterial resistance to kanamycin ................................................................................... 28

Figure 6. Western blot of 1 pg TSP from transgenic maize plants expressing E1 (To).
Lanes: +: positive tobacco control; -C: negative maize control (untransformed); 1-9:

transgenic maize plants. Invitrogen Magic MarkTM Western Standard used for size
markings. Percentages El as determined by enzyme activity assay are displayed above
bands (Table 2) .............................................................................................................. 42

Figure 7. Average conversion of cellulose to glucan using E1 produced from transgenic
maize. The substrates used in the experiment were Avicel, carboxymethyl cellulose
(CMC) and AF EX-treated corn stover (ACS). The enzymatic hydrolysis was done for a
period of 72 h, at 50°C at 90 rpm. T=TSP from transgenic plants; NT=TSP from non-
transgenic control plants. Error bars represent standard deviation from the mean. .......... 43

Figure 8. PCR analysis of T1 maize plants for E]. W: water; -C: non-transgenic maize

control DNA; 1-15: T1 plants from known crosses; +C: plasmid DNA (pMZ766-E1CAT).
...................................................................................................................................... 44

xii

Figure 9. Results of PCR for 30. Individual plants indicated above the lanes are
represented by line—plant number. W: reaction containing only water; 1 kb: 1 kb marker
(NEB); P: pUC1813. ..................................................................................................... 48

Figure 10. Results of PCR of maize transformed with 30 for the 36 gene. Individual
plants indicated above the lanes are represented by line-plant number. W: reaction
containing only water; -C: Non-transformed negative control; 1 kb: 1 kb marker (N EB);
P: pUC1813 ................................................................................................................... 48

Figure 11. Northern blots of maize expressing 30. Individual plants indicated above the
lanes are represented by line-plant number. Ethidium-bromide-stained RNA bands
corresponding to each of the plants fromthe gel before are displayed below the blots to
show relative loaded amounts. +C: Positive control (plant lO-l); —C: Untransformed
maize negative control ................................................................................................... 50

Figure 12. Southern blots of maize transformed with BC. Genomic DNA (30 pg) and
pUCl8l3 plasmid DNA (1 ng) were digested overnight with [A] BglII (left panel) or
Ncol (right panel) or [B-E] NcoI and fractionated on a 1% w/v agarose gel. Individual
plants indicated above the lanes are represented by line-plant number. 1 kb: 1 kb
molecular weight marker (NEB); P: pUC1813 positive control; -C: Untransformed maize
inbred line Hill. Panels D and E represent the same samples but different exposure times
(8 h and ‘/2 h, respectively) ............................................................................................. 53

Figure 13. Top panel: Glucose (g/L) released from 1% cellobiose at 0 min, 1 h, 7.25 h
and 26.5 h, using 570 pl maize TSP from plants expressing 80. Data have been adjusted
by subtracting the substrate blank. Bottom panel: Glucose (g/L) released per mg protein
in TSP. Numbers below the bars represent maize plants expressing 80. Hill:
Untransformed maize negative control. ......................................................................... 58

Figure 14. Glucose (g/L) released from 1% cellobiose at 0 min, 1 h, 7.25 h and 26.5 h,
using 570 pl maize TSP from plants expressing 30. Data have been adjusted by
subtracting the substrate blank. Top panel: Lines represent different maize plants
expressing BG; numbers below the lines represent sample collection time points; Hill:
Untransformed maize negative control. Bottom panel: T: transgenic plants; NT: non-
transgenic plants. Dashed lines represent 95% confidence intervals; solid lines represent
polynomial regression lines; dots represent individual data points. ................................ 59

Figure 15. Sugar determination via DNS on a 0.5% CMC plate. SPC: Spezyme CP;
MFX: Multifactorial xylanase; BG: Novozyme 188 ....................................................... 63
Figure 16. Sugar determination via DNS on a 1% cellobiose plate. Bottom panel:
Undiluted Novozyme 188 is not included to show the other dilutions in more detail ...... 64

Figure 17. Difference between glucose released in commercial enzyme (Novozyme 188)
vs. substrate background (blank) .................................................................................... 65

xiii

Figure 18. Sugar determination via glucose analyzer on a 1% cellobiose plate after
hydrolysis ...................................................................................................................... 66

Figure 19. Glucose released from 1% CMC after 1 h hydrolysis using TSP from plants
expressing E1 or CBHI .................................................................................................. 67

Figure 20. Glucose released from 1% CMC or Avicel after 24 h hydrolysis using TSP
from plants expressing E1 or CBHI. Numbers above 24 h bars indicate times higher than
0 h data .......................................................................................................................... 68

Figure 21. Sugar release from 1% Avicel (AV) or CMC after 24 h by application of
various combinations of plant TSP containing hydrolysis enzymes, determined by DNS
assay. E: El (30 pg); C: CBHl (120 pg); B0.1: BG (15 pg); B0.5: BG (75 pg); E:C: El
and CBHl (1:4 ratio); E:C:BO.1: E1, CBHl and BG (15 pg) (1:4:0.1 ratio); E:C:BO.5:
E1, CBHl and BG (75 pg) (1:4:0.5 ratio); E:C:Bl: El, CBHl and BG (150 pg) (1:4:1
ratio); AV: 1% Avicel; CMC: 1% CMC. ....................................................................... 71

Figure 22. Sugar release from 1% Avicel (AV) or CMC after 24 h by application of
various combinations of plant TSP containing hydrolysis enzymes, determined by
glucose analyzer. E: El (30 pg); C: CBHl (120 pg); BO. 1: BG (15 pg); B0.5: BG (75
pg); E:C: El and CBHl (1:4 ratio); E:C:B0.1: E1, CBHl and BG (15 pg) (1:4:0.1 ratio);
E:C:BO.5: El, CBHl and BG (75 pg) (1:4:0.5 ratio); E:C:Bl: El, CBHl and BG (150
pg) (1:4:1 ratio); AV: 1% Avicel; CMC: 1% CMC ........................................................ 72

Figure 23. Average sugar release after subtracting enzyme blanks determined by DNS
assay. E:C: E1 and CBHl (1:4 ratio); E:C:B0.1: El, CBHl and BG (15 pg) (1:4:0.1
ratio); E:C:BO.5: E1, CBHl and BG (75 pg) (1:4:0.5 ratio); E:C:Bl: El, CBHl and BG
(150 pg) (1:4:1 ratio); AV: 1% Avicel; CMC: 1% CMC. E:C:BO.1 for CMC is likely
higher than shown; E:C:B0.5 for AV is likely lower than shown; E:C:Bl for both is likely
lower than shown. ......................................................................................................... 74

Figure 24. Glucose release after subtracting enzyme blanks determined by glucose
analyzer. E:C: El and CBHl (1 :4 ratio); E:C:BO. 1: El, CBHl and BG (15 pg) (1 :4:0.1
ratio); E:C:BO.5: E1, CBHl and BG (75 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG
(150 pg) (1:4:1 ratio); AV: 1% Avicel; CMC: 1% CMC. E:C:BO.1 for CMC is likely
higher than shown; E:C:B0.5 for AV is likely lower than shown; E:C:Bl for both is likely
lower than shown. ......................................................................................................... 75

Figure 25. Sugar release on 1% Avicel after 18 h; enzyme blanks subtracted. E:C: E1 and
CBHl (1:4 ratio); E:C:B0.l: E1, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: El,
CBHl and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG (22.8 pg) (1:4:1
ratio); E:C:B2: El, CBHl and BG (45.5 pg) (1:4:2 ratio). ............................................. 77

Figure 26. Sugar release on 1% CMC after 18 h, enzyme blanks subtracted. E:C: El and
CBHl (1:4 ratio); E:C:B0.1: E1, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: E1,

xiv

CBHl and BG (11.4 pg) (1:4:0.5 ratio); E:C2B1: El, CBHl and BG (22.8 pg) (1:4:1
ratio); E:C:B2: E1, CBHl and BG (45.5 pg) (1:4:2 ratio). ............................................. 77

Figure 27. Sugar release on 1% Avicel after 18 h; enzyme blanks subtracted. E:C: El and
CBHl (1:4 ratio); E:C:BO.1: E1, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: E1,
CBHl and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG (22.8 pg) (1:4:1
ratio); E:C:B2: El, CBHl and BG (45.5 pg) (12422 ratio). Data for series “Plant TSP
alone” is same as Figure 25, shown here for comparison. .............................................. 79

Figure 28. Sugar release on 1% CMC after 18 h; enzyme blanks subtracted. E:C: El and
CBHl (1:4 ratio); E:C:BO.1: El, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: E1,
CBHl and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: El, CBHl and BG (22.8 pg) (1:4:1
ratio); E:C:B2: E1, CBHl and BG (45.5 pg) (124:2 ratio). Data for series “Plant TSP
alone” is same as Figure 26, shown here for comparison. .............................................. 80

Figure 29. Sugar release on 1% ACS afier 24 h; blanks subtracted. E:C: E1 and CBHl
(1:4 ratio); E:C:BO.1: E1, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: El, CBHl
and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: El, CBHl and BG (22.8 pg) (1:4:1 ratio);
E:C:B2: E1, CBHl and BG (45.5 pg) (1:4:2 ratio) ......................................................... 81

Figure 30. Sugar release on 1% ACS after 24 h; blanks subtracted. E:C:Novo: E1 and
CBHl (1 :4 ratio) plus Novozyme 188; SP:B0.1: Spezyme CP (SPC) and BG (2.3 pg);
SP2BO.5: SPC and BG (11.4 pg); SPzBl: SPC and BG (22.8 pg); SP2B2: SPC and BG
(45.5 pg). ...................................................................................................................... 83

Figure 31. PCR for FLC of maize plants co-transformed with either XYLl (X-) or
pMSFlS. W: Reaction containing only water; -C: nontransgenic negative control maize
DNA; P: pGreen plasmid DNA. .................................................................................... 95

Figure 32. PCR for F LC on maize plants co-transformed with BG and FLC. W: Reaction
containing only water; L: 100 bp ladder (NEB), P2 pGreen. ........................................... 96

Figure 33. Southern blots of maize co-transformed with pSMFlS (left side) or XYL (right
side) and pGreen. HiII: Non-transgenic negative control; P: pSMFlS; F: pGreen. ......... 96

Figure 34. Northern blots of maize co-transformed with pSMFlS (left side) or XYLl (X—;
right side) and pGreen. Ethidiurn-bromide stained RNA bands are shown below the blot
to show amount loaded. —C and —C(M): Non-transgenic negative control. ..................... 97

Figure 35. XYL Construct. CaMV 35S: Cauliflower mosaic virus 35S promoter; 9:
tobacco mosaic virus translational enhancer; Prla: signal peptide from tobacco
pathogenesis-related protein 1a;XYL1: xylanase gene from C. carbonum; nos: nos
terminator .................................................................................................................... 100

Figure 36. Maize plants transformed with XYL] and FLC. ........................................... 102

XV

Figure 37. PCR for XYL in maize and tobacco plants. Numbers followed by M indicate
maize plants; numbers followed by T indicate tobacco plants. W: Reaction containing
only water; -C: Untransformed maize plant; P: XYL] plasmid ...................................... 103

Figure 38. Southern blot of maize and tobacco plants transformed with XYL] and FLC.
Numbers followed by M indicated maize plants; numbers followed by T indicate tobacco
plants. P: XYL] plasmid; -C2 non-transformed maize (M) or tobacco (T). .................... 104

Figure 39. Northern blot of maize and tobacco plants transformed with X11] and FLC.
Numbers followed by M indicated maize plants; numbers followed by T indicate tobacco
plants. The probe was a 397 bp PCR-generated fragment of the XYL] gene. Upper panel:
blot probed with PCR-labeled DIG probe; bottom panel: same blot probed with random
primed DIG labeled probe. -CM2 non-transforrned maize; -CT non-transformed tobacco.
Ethidium bromide-stained bands from the agarose gel prior to transfer are shown below
the blot to show relative amounts of RNA loaded. ....................................................... 105

Figure 40. RT-PCR of maize and tobacco plants transformed with XYL] and FLC.
Numbers followed by M indicated maize plants; numbers followed by T indicate tobacco
plants. Top panel: RT reaction; bottom panel: No-RT control. -CM: non-transformed
maize; -CT non-transformed tobacco; M: 100 bp molecular weight marker (NEB). ..... 106

Figure 41. Western blot of maize plants transformed with XYL] and FLC.
Letters/numbers above the lanes represent individual plants; 1M, 2M and 4M correspond
with the plants that survived to maturity and are represented in the other figures. —C2
Untransformed maize negative control. Size markings are indicated on the right side. . 107

Figure 42. pSMF 15. rch: rice Rubisco small subunit promoter, TP: rice rch chloroplast
signal peptide; Syn-cbh] 2 synthetic CBHI ; nos: nopaline synthase 3’ non-coding region.
.................................................................................................................................... 111

Figure 43. Southern blot of maize plants transformed with Syn-CBHI and FLC. P:
pMSF 15 plasmid; -C: non-transformed maize. ............................................................ 113

Figure 44. Northern blot of maize plants transformed with Syn-CBHI and FLC. The
probe was a 801 bp PCR-generated fragment of the Syn-CBHI gene. Left panel: blot
probed random primed DIG labeled probe; right panel: same blot'probed with PCR-
labeled DIG probe. -C: non-transformed maize. Ethidium bromide-stained bands from the
agarose gel prior to transfer are shown below the blot to show relative amounts of RNA
loaded. ........................................................................................................................ 1 14

Figure 45. RT-PCR of maize plants transformed with Syn-CBHI and FLC. Top panel: RT

reaction; bottom panel: No-RT control. -C: non-transformed maize; M: 100 bp molecular
weight marker (N EB). Arrow indicates 801 bp. ........................................................... 114

xvi

Figure 46. pZD408. 358: Cauliflower Mosaic Virus (CaMV) 358 Promoter; SynCBHI:
Synthetic CBHI coding region. .................................................................................... 115

xvii

KEY TO SYMBOLS AND ABBREVIATIONS

3SS—t: cauliflower mosaic virus (CaMV) 35$ terminator
4CL2 4-courmarate CoA ligase

ACS: AF EX-treated corn stover

Actl-S’: rice actin 5’ region (promoter)

AF EX: ammonia fiber explosion

AGP: ADP-glucose pyrophosphorylase

bar: bar gene encoding Bialaphos herbicide resistance
BG: B-glucosidase from Buoirivibriofibrisolvens H17c
bglA: gene encoding B-glucosidase

C3H: 4-coumarate 3-hydroxylase

CAD: cinnamyl alcohol dehydrogenase

CAldSH: coniferaldehyde 5-hydroxylase

CaMV 35S: cauliflower mosaic virus (CaMV) 358 promoter
CBHI: cellobiohydrolase I from T richoderma reesei
CCR: cinnamoyl CoA reductase

CMC: carboxymethyl cellulose

C02: carbon dioxide

DNS: dinitrosalicylic

E12 Endoglucanase E1 from A cidothermus cellulolyticus

EICAT: catalytic domain of endoglucanase E1 from Acidothermus cellulolyticus
E2: endoglucanase E2 of T hermomonospora fusca

E3: cellobiohydrolase E3 of T hermomonospora fusca

ER: endoplasmic reticulum

FLC: FLOWERING LOCUS C, a floral repressor gene identified in Arabidopsis
Hill: maize inbred line developed for its ability to produce highly proliferating, Type II
embryogenic callus from immature embryos

H VA 1 2 barley H VA 1 gene

IUPAC: International Union of Pure and Applied Chemistry

LB: T-DNA lefi border

MU: the fluorophore 4-methylumbelliferone

MUC: 4-methylumbelliferone B-D-cellobioside

N: number of replicates

(3’) nos: polyadenylation signal from the nopaline synthase gene

nptII: gene conferring bacterial resistance to kanamycin

NREL: National Renewable Energy Laboratory

OMT: O-methyl transferase

pBY520: plasmid containing bar and barley H VA 1 genes

pDM302: plasmid containing the bar gene

pGreen: binary vector containing the bar, nptII and FLC genes

pinII: potato proteinase inhibitor 11 terminator

pMSF 15: plasmid containing the Syn-CBHI gene

xviii

pMZ766-E1CAT2 plasmid containing Elc AT from A cidothermus cellulolyticus
pNPBG: p-nitro-phenyl-B-D-glucopyranoside

pNP: p-nitrophenol

Prla: tobacco pathogenesis-related protein 1a(Pr1a)
pUC1813: plasmid containing the ngA gene

RB: T-DNA right border

S.D.2 standard deviation

S.E.: standard error

SAR: systemic acquired resistance

TSP: total soluble protein

VT: vacuole-targeting sequence

XYLl: vector containing the XYL] gene

XynA: xylanase from Clostridium thermocellum
xynA: xylanase gene from Neocallimastix patriciarum
xynB: xylnanase from Streptomyces olivaceoviridis

Q: tobacco mosaic virus (TMV) translational enhancer

xix

1. LITERATURE REVIEW

Production of Heterologous Hydrolysis Enzymes within Crop Biomass for Biofuel

Ethanol

1. Introduction

Ethanol fuel is a promising alternative to fossil fuels, which damage the
environment by contributing to net carbon dioxide increase. In addition, they will
eventually be depleted, and increase dependence on foreign oil imports. According to a
recent report from the Natural Resources Defense Council and the Institute for the
Analysis of Global Security, the dependence of the United States on foreign petroleum
both undermines its economic strength and threatens its national security (Bordetsky er
al. 2005). The use of ethanol fuel, obtained either from grain or from cellulosic materials,
can help decrease the need for petroleum fuel (Bordetsky et al. 2005). Accordingly, the
ethanol fuel industry has been growing significantly in many countries throughout the
world. In the US, ethanol production capacity reached 3.5 billion gallons in 2004, up by
303 million gallons from 2003 (Renewable Fuels Association 2007). Ethanol fuel is
clean-buming and does not contribute to net carbon dioxide increase, is renewable, and
can be produced using resources the country already possesses.

Ethanol is produced from the fermentation of sugars (usually sucrose or glucose)
by yeast. The carbon (sugar) source is called the feedstock. Most feedstocks are plant
materials. The most widely used feedstocks today are sugarcane and maize grain. The
sugar in sugarcane is easily extracted and used directly for fermentation, while the maize

grain must be milled and its starch hydrolyzed to glucose by a-amylase. In the US,

ethanol is mostly produced from the starch of maize grain with a net energy balance of
1.34; that is, for every unit of energy expended in growing corn and converting it to
ethanol, 1.34 units of energy (automotive fuel) are obtained (Biomass Program: Net
Energy Balance for Bioethanol Production and Use; Shapouri et al. 2002). The most
efficient farming and ethanol production systems in place can achieve a balance of 2.09
(Biomass Program: Net Energy Balance for Bioethanol Production and Use). Starch
fermentation is thus relatively efficient. However, there is a very rich source of glucose
that has so far been underutilized: cellulose.

Cellulose, composed of B-glucose units, is the most abundant polymer on earth. It
is a structural component of the plant cell wall. It has traditionally not been used as a
carbon source because its location inside microfibrils, which are wrapped in
hemicellulose and embedded in a matrix of lignin, makes it inaccessible to hydrolysis
enzymes unless the plant material goes through extensive pretreatment. However, recent
advancements have made using this resource a possibility. In this chapter, we explore the
problems, challenges, and solutions to ethanol production from cellulosic materials, with

a focus on utilizing plants as biofactories for hydrolysis enzyme production.

2. The Plant Cell Wall

The plant cell wall is a highly organized structural component composed of a
myriad of different polysaccharides, proteins, aromatic substances and other compounds.
It has several important functions: it provides structure to the cell, thus determining its
shape and even function; it aids in defense against invading pathogens; and it contains
signaling molecules that can alert the cell to various environmental stimuli, including

pathogenic attack (Carpita and McCann 2002). It is a dynamic structure, and its

configuration and composition can vary by plant species, age, tissue, cell types and even
within cell wall layers (Ding and Himme12006; Bothast and Schlicher 2005). The
primary cell wall is formed first from the cell plate during cell division and forms the
outside of the cell. Between primary cell walls of adjacent cells is the middle lamella.
Secondary cell wall synthesis, if present, usually begins after the primary cell wall has
stopped growing, being deposited on the interior of the primary cell wall, ofien in layers
(Carpita and McCann 2002).

Polysaccharides are the primary constituents of the cell wall and form its main
structural scaffold. They are composed of long chains of sugar molecules that are
covalently linked at various positions and may have side chains. They are made up of
various combinations of the 11 monosaccharide sugars commonly found in plant cell
walls: glucose (from which all the others are derived), rhamnose, galactose, galacturonic
acid, glucouronic acid apiose, xylose, arabinose, mannose, mannuronic acid and fucose

(Carpita and McCann 2002).

2.1 Cell wall components

2.1.1. Cellulose

Cellulose is a long, unbranched polymer of up to 15,000 molecules of anhydrous
glucose. The glucose molecules are arranged in B-l,4 linkages, which means that each
unit is orientated 180° relative to the unit it is attached to. In other words, cellulose is
composed of cellobiose units (diglucose molecules connected via B-1,4 linkages).
Cellulose is an important polysaccharide found in the primary and secondary cell walls in
the form of microfibrils. It makes up 15-30% of the dry mass of primary cell walls and up
to 40% of secondary cell walls. The cellulose chains in microfibrils are lined up parallel

3

to each other and consist of crystalline regions, where the cellulose molecules are tightly
packed, and amorphous (also called soluble) regions, where the arrangement is less
compact. The amorphous regions are staggered so that the overall structure remains
strong. A microfibril has a diameter of around 30 nm and consists of around 36 cellulose

chains, but the number varies with species (Carpita and McCann 2002).

2.1.2 Cross-linking glycans

Microfibrils are coated with other polysaccharides, cross-linking glycans (also
called hemicelluloses), which link them together. The two major types are xyloglucans,
found in dicots and around half of the monocot species, and glucuronoarabinoxylans,
which are found in commelinoid monocots, including the cereals and grasses.
Xyloglucans have a backbone of glucosyl residues in 1,4-13 linkages, with xylosyl units
attached; glucuronoarabinoxylans have a backbone of xylosyl residues in 1,4-[3 linkages
to which glucosyluronic acid and arabinosyl units are attached. The grasses also have a
third major cross-linking glycan, called “mixed linkage” (1—>3),(l——>4)B-D-glucans (B-
glucans), which are unbranched polymers with a 2:1 ratio of cellotriose to cellotetraose
units connected by (l—>3)[3-D-1inkages, resulting in a coiled shape. Various mannans are
also present in smaller amounts (Carpita and McCann 2002). Hemicellulose accounts for

20-40% of the total dry weight of plant matter.

2.1.3 Pectins and other substances

Pectins are a mixed group of various branched, hydrated polysaccharides
abundant in galacturonic acid. In dicots, they account for approximately 35% of the dry

weight (Carpita and Gibeaut 1993); in monocots they are much less abundant. They serve

many functions in the cell wall: they establish wall porosity, adjust wall pH and ion
balance through charged surfaces, control bonding between cells at the middle lamella,
and also function as recognition molecules to alert the cell to the presence of
microorganisms or insects (Ridley et al. 2001; O'Neill et al. 2004). Pectins are mostly
made up of homogalacturonan and rhamnogalacturonan I; rhamnogalacturonan II,
arabinans, galactans, and arabinogalactans are also present in smaller quantities. In
addition to pectins, structural proteins and aromatic substances can also be present

(Carpita and McCann 2002).

2.1.4 Lignin

Lignin is almost nonexistent in primary cell walls but is a chief constituent in
some secondary walls, and accounts for about 10-25% of the total dry weight. It is
composed of aromatic compounds called phenylpropanoids arranged in complex systems.
These networks are linked to the carbohydrates, including cellulose and xylose, in various
bonds, including ester, ester; phenyl, phenyl; and covalent bonds (Carpita and McCann
2002). Lignin protects the cell against pathogen invasion and will often be deposited in

response to attack, providing additional structure and strength (Mosier et al. 2005).

2.2 Two major types of primary cell wall

The basic structure of primary cell walls consists of the scaffold of cellulose and
cross-linking glycans, embedded in a second (and sometimes third) complex. There are
two types of primary cell wall that differ in the kind of cross-linking glycan, which
determines the wall type. Type I walls are found in those plants that have xyloglucans;

they have approximately equal amounts of xyloglucan and cellulose. Xyloglucans coat

the cellulose microfibrils and bind them together, and this complex is embedded in a
matrix of pectin. Type II walls are found in plants whose major cross-linking glycans are
glucuronoarabinoxylans; they lack pectin and structural proteins, instead amassing
phenylpropanoids (Carpita and McCann 2002). Type 11 cell walls are found in cereals and

grasses, and thus are of greatest interest for cellulosic ethanol research.

3. Cell wall degradation

3.1 Microorganisms

Several microorganisms (bacteria and fungi) have been studied for their ability to
break down cell walls, including anaerobes (such as those present in the rumen) and
aerobes (such as those that decompose dead plant matter). Most organisms that can
degrade cellulose produce a number of enzymes, which form a system that hydolyzes
various polysaccharides, since the enzymes first have to penetrate the hemicellulose
shield before they can attack the cellulose (Warren 1996).

Anaerobic microorganisms known for their cell—wall degrading ability include the
bacteria Butyrivibriofibrisolvens Hl7c, F ibrobacter succinogenes S85, Ruminococcus
flavefaciens 17, R. albus, Prevotella ruminicola Bl 4, Clostridium thermocellum, C.
cellulovorans, C. cellulolyticum, C. stercorarium and Caldocellulosiruptor
saccharolyticus, and the fungus Neocallimastixfiontalis. Aerobic microorganisms
include the bacteria Acidothermus cellulalyticus (Tucker er al. 1989), Pseudomonas
fluorescens subsp. cellulosa, Streptomyces lividans 66, S. reticuli, S. halstedii,
Cellulomonasfimi, C. uda and Microbispora bispora, and the fungi Thermomonospora

fusca, Trichoderma reesei and Phanerochaete chrysosporium (Warren 1996).

6

These organisms produce many different enzymes that may be grouped according
to their primary activities: endoglucanases, exoglucanases (also called
cellobiohydrolyases), B-glucosidases, cellodextrinases, xylanases, xylosidases,
lichenases, mannanases, laminarinases, arabinofuranosidases and avicelases. In order to
decrystallize and hydrolyze cell walls, they must produce systems of many different
enzymes (for each of the cell wall components) that act synergistically; this has been well
documented. The enzymes vary in their substrate specificity: some exclusively act on a
particular substrate, while others can utilize more than one; some have more activity on
one substrate over another; and some can break only certain bonds, while others can
cleave more than one bond type. In addition, different enzymes often produce different
products from the same substrate. Therefore microorganisms may produce several
different enzymes, for specific substrates or bonds or both. Some microorganisms, such
as Clostridia spp., produce cellulosomes (Demain et al. 2005), complexes of multiple
enzymes held together in a specific conformation by proteins that are very efficient at cell

wall hydrolysis (Warren 1996).

3.2 Hydrolysis

The major classes of enzymes needed for cell wall hydrolysis are cellulases,

hemicellulases and ligninases.

3.2.1 Cellulases
Three types of cellulases are needed to obtain glucose from cellulose:
endoglucanase (E1; EC. 3.2.1.4), cellobiohydrolase (also called exoglucanase) (EC.

3.2.1.91), and B-glucosidase (EC. 3.2.1.21) (Ziegler et al. 2000; Ziegelhoffer et al.

2001). Enzymatic hydrolysis of plant cell wall polysaccharides to glucose is a three-step
process. First, endoglucanase randomly cleaves the crystalline regions of cellulose,
exposing chain ends. Then, cellobiohydrolase attaches to the chain end and threads it
through its active site, processively cleaving off cellobiose units; it can also act on
amorphous regions with exposed chain ends without prior endoglucanase activity.
Exoglucanases work from either the reducing or non-reducing end of the sugar, not both;
cellulase hydrolysis is more efficient if both types are produced. Finally, B-glucosidase

breaks the bonds of cellobiose to produce single glucose units (Warren 1996).

3.2.2 Hemicellulases

For cellulases to access the cellulose, the hemicellulose surrounding it must be
removed. While cellulose consists of a single monosaccharide and type of bond,
hemicelluloses are amorphous and diverse. Since the major constituent of hemicellulose
is B-l,4-xylan, the most abundant class of hemicellulase is xylanase, which can have both

endo- and exo- activity (Warren 1996).

3.2.3 Ligninases

Lignin degradation by microorganisms is less well understood than that of
polysaccharides. The most effective lignin-degrading microbes in nature are thought to be
white rot fungi (D'Souza et al. 1999), especially Phanerochaete chrysosporium and
T rametes versicolor. The three major families of lignin-modifying enzymes produced by
fungi are laccases, manganese-dependent peroxidases, and lignin peroxidases
(Boominathan and Reddy 1992; Hatakka 1994; Kirk and Farrell 1987; Thurston 1994).

They oxidize compounds by using or creating radicals.

4. Ethanol production

4.] Maize grain ethanol production

Ethanol produced from maize grain is a mature technology. It is attractive because
it benefits farmers and local communities by providing jobs, a valuable resource, and
valuble coproducts (such as distillers grains and corn gluten). As of 2007, 124
biorefineries are in operation and 76 more are being constructed. Ethanol production
currently stands at nearly 6.5 billion gallons a year and will reach 12.9 billion gallons per
year upon the plants’ completion (RFA - The Industry - Plant Locations), which could
displace 4.7 and 9.3 billion gallons of gasoline respectively (if E85, 3 firel blend of
gasoline and up to 85% ethanol, is used). However, this only covers around 3% or 6.7%
respectively of the total gasoline consumed annually in the US (137 billion gallons in
2006; (U .S. Prime Supplier Sales Volumes of Petroleum Products).

US maize growers produced 10.5 billion bushels of maize grain in 2006 (World of
Com 2007); 18.3% was used in ethanol production (World of Com 2007). This is the
equivalent of 2.2 billion bushels, or 6.2 billion gallons of ethanol, which likely displaced
4.4 billion gallons of gasoline (3.2% total consumption). Since current ethanol plant
capacity is 2.3 billion bushels, becoming 4.6 billion bushels, grain production must
increase to meet capacity, or must be diverted from other uses. Currently, 50.8% of total
production, or 6 billion bushels, is used for livestock feed (World of Com 2007). Much of
this could be successfully diverted to ethanol fuel production as the grain could be
replaced with nutritious distillers grains. To meet capacity, only 1.7% (currently) or 40%
(when the plants are completed) need be diverted from grain destined for livestock feed

9

(0.1 billion bushels and 2.4 billion bushels respectively). Therefore meeting production
capacity from maize grain is an attainable goal and likely to be realized. However, if all
the maize grain produced in the US were used for ethanol fuel production, only 29.4
billion gallons would be produced, the equivalent of 21.2 billion gallons of fuel, or 15.4%
of current usage (Houghton et al. 2005). Clearly, an alternative to maize grain ethanol is

needed.

4.2 The promise of cellulosic ethanol

According to Kim and Dale (2004), worldwide wasted crops and lignocellulosic
waste crop residue could translate into 129.7 billion gallons of ethanol and replace 93.4
billion gallons of gasoline (about 32% of current worldwide consumption) if E85 is used.
About 90% of this estimate comes from crop residue waste. This number could be much
higher if biofuel crops were grown to supplement this amount and if the technology were
in place to produce it. Worldwide availability of lignocellulosic feedstocks is estimated at
over 1.7 billion tons per year (Kim and Dale 2004), with some estimates reaching 10—50
billion tons of crop biomass annually (Greene et al. 2004). In addition to being
inexpensive and widely available, lignocellulosic biomass has the added benefit of being
renewable and thus sustainable (Kim and Dale 2004; Greene et al. 2004). It is believed
that with proper management, roughly 1.3 billion tons of crop and forest residues and
energy crops can become available annually in the US (Perlack et al. 2005), the majority
of which could be used for conversion to alcohol fuels, yielding the equivalent of
approximately 108.5 billion gallons of gasoline (Kim and Dale 2004).

A current goal for enhancing US economic security is to meet 10% of chemical

feedstock demand by 2020 with plant-derived materials, or a fivefold increase over

10

current usage levels (Singh et al. 2003). Crops that have a high amount of lignocellulosic
biomass, such as corn, rice, sugarcane and fast growing perennial grasses have been
recommended for conversion to alcohol fuels (Knauf and Moniruzzaman 2004; Sticklen
2004)

Construction of commercial cellulosic biomass ethanol facilities is currently
underway in the US. These facilities will have the capactiy to collectively produce 226.4
million gallons per year. They include: Abengoa Abengoa Bioenergy, NE; Akico, Inc.,
FL; Bluefire Ethanol, CA; Broin Companies, IA; Iogen Biorefinery Partners, ID; and
Range Fuels, GA (Bruce Dale, Michigan State University, pers. comm). In Canada,
Iogen Corporation has a demonstration biomass ethanol plant currently in operation that

can produce about 660,000 gallons of ethanol per year.

4.2.1 Cellulosic ethanol production

To produce ethanol from biomass, several events must take place: the hydrolysis
enzymes must be produced (usually in microbial fermentation tanks), the biomass must
undergo a pretreatment process to disrupt the lignin and expose the cellulose, the
enzymes must be added to the pretreated feedstock, and the resulting sugars must be

fermented and distilled.

4.2.2 Challenges to cellulosic ethanol production

Although production of fermentable sugars for alcohol fuels fiom plant biomass is
an exciting and attractive idea, and substantial efforts have been made toward improving
ethanol yield through this technology and reducing its production costs (Ingledew 1995;

Lynd et al. 2005), major roadblocks still stand in the way of widespread commercial

ll

implementation of this technology. These include prohibitive costs of pretreatment
processing of the lignocellulosic matter, with estimates of up to $0.30/gallon (Mosier et
al. 2005) and production of microbial cellulase enzymes used in the conversion of
cellulosic matter to fermentable sugars (Kabel et al. 2006).

Removal of lignin is the major roadblock to this process and an area of intense
research because of the high cost involved. Although research is ongoing in the area of
fungal ligninases (mentioned above) and reduction of lignin content (described below) in
order to decrease the necessity (and thus the cost) of pretreatment, pretreatment is
currently required. Several pretreatments have been deve10ped so far, including dilute
acid, flow-through, ammonia fiber explosion (AFEX), ammonia recycle percolation,
steam water explosion, lime, and organosolv pulping (Eggeman and Elander 2005;
Mosier et a1. 2005; Wyman et al. 2005b; Wyman et al. 2005a; Pan et al. 2005).

Currently, production of hydrolysis enzymes in microbial fermentation tanks is
expensive (Knauf and Moniruzzaman 2004; Howard et al. 2003). Although decades of
research have been devoted to reducing microbial production costs, resulting in
significant decreases since 1980 (Knauf and Moniruzzaman 2004; Wyman 1999),
enzyme production is still costly (Knauf and Moniruzzaman 2004). The latest cost-
reduction model designed by the National Renewable Energy Laboratory (NREL) and
Genencor is to produce cellulases at around $010-$020 per gallon of ethanol (Genencor
Celebrates Major Progress in the Conversion of Biomass to Ethanol - Genencor a
Danisco division). A possible solution to these problems is to use biomass crops as

biofactories to produce these enzymes on a large scale.

12

5. Production of Hydrolysis Enzymes in Biomass Crops

5 . 1 Plants as molecular biofactories

Plants are already being used successfully for molecular farming (Horn et al.
2004) of enzymes (Hong et al. 2004; Chiang et al. 2005) and other proteins (Liu et al.
2005), carbohydrates (Schulman 2002; Sahrawy et al. 2004), lipids (Qi et al. 2004),
polymers such as polyhydroxybutyrate (Bohmert et al. 2002); (Saruul et al. 2002; Zhong
et al. 2003) and pharmaceuticals (Howard and Hood 2005). Plant-based production of
enzymes has several critical advantages compared to microbial fermentation or
bioreactors. For example, plants can use the sun’s energy directly, requiring fewer energy
inputs. Furthermore, proteins produced in plants generally display correct folding,
glycosylation, activity, reduced degradation and increased stability (Horn et al. 2004). In
addition, the infrastructure and expertise are already available for plant genetic
transformation, growing, harvesting, transporting and processing plant matter (Horn et al.
2004).

The US Government has recently urged the agricultural and petrochemical
industries to discover and employ alternatives to fossil fuels to both decrease dependence
on foreign oil and promote a cleaner environment. A specific recommendation was to
develop technology that would allow production of cellulases and other hydrolysis
enzymes in plants (Ragauskas et al. 2006; Sticklen 2007b; Sticklen 2007a; Sticklen 2004;
Sticklen 2006), which has the potential to reduce enzyme production costs. Extraction of
plant total soluble protein (TSP) from leaves is quick and easy, and could be done at the
ethanol production facilities; alternatively, the enzymes could be extracted and

lyophilized for inexpensive storage and easy transport.

13

5 .2 Successful plant-produced hydrolysis enzymes

The catalytic domain of the thermostable endo-l,4-l3 glucanase (E 1) of A.
cellulolyticus (Tucker et al. 1989; Baker et al. 1994) has been successfully produced in
Arabidopsis (Ziegler et al. 2000), tobacco (Ziegelhoffer et al. 2001), rice (Oraby et al.
2007), and maize (Biswas et al. 2006; Ransom et al. 2007). The full-length peptide has
been expressed in potato (Dai et al. 2000b) and tobacco (Dai et al. 2000a; Dai et al. 2005;
Ziegelhoffer et al. 2001). Expression of the catalytic domain yielded more activity than
the full-length enzyme (Ziegelhoffer et al. 2001). The thermostable endoglucanase E2 of
Thermomonosporafirsca was expressed in tobacco, potato and alfalfa (Ziegelhoffer et al.
1999).

Rice- and maize-produced transgenic TSP containing E1 was able to convert
AF EX-treated corn stover to glucose in conversion analyses with the addition of B-
glucosidase (Novozyme 188, Sigma). The enzyme retained activity in dry tissue and after
several months’ storage in the freezer (Oraby et al. 2007; Ransom et al. 2007).

Exoglucanases have also been expressed in plants. The thermostable
cellobiohydrolase E3 of Thermomonosporafirsca was expressed in tobacco, potato and
alfalfa (Ziegelhoffer et al. 1999). T richoderma reesei cellobiohydrolase I (CBHI) was
produced in transgenic tobacco (Dai et al. 1999). Although a low amount of protein was
produced in both cases, biological activity has been low as well.

B-glucosidases have been expressed in plants, although traditionally for reasons
other than production of enzymes for hydrolysis. Human acid B-glucosidase was
successfully expressed in transgenic tobacco seeds for medical purposes (Reggi et al.

2005). Maize B-glucosidase was expressed in tobacco to study cytokinins (Kiran et al.

14

2006). Butyrivibriofibrisolvens H17c B-glucosidase was expressed in tobacco to study
whether it could effect enhanced immune response through systemic acquired resistance
(SAR) (Yao 2004). This work is the first report of heterologous microbial B—glucosidase
expressed in a plant for the purpose of obtaining biologically-active enzyme for use in the
production of ethanol fuel.

Microbial xylanases have also been produced in plants. A modified xylanase gene
(xynA) from the rumen fungus, Neocallimastix patriciarum, was successfully expressed
in barley endosperm, retaining activity afier desiccation and storage (Patel et al. 2000). A
thermostable xylanase from Clostridium thermocellum was expressed in the apoplast of
tobacco (Herbers et al. 1995) and the catalytic domain of XynA from the same organism
was expressed in both cultured tobacco cells (Kimura et al. 2003a) and rice (Kimura et
al. 2003b). The xynB gene of Streptomyces olivaceoviridis Al was expressed in potato
(Solanum tuberosum); its enzyme activity was retained over several generations (Yang et
al. 2007). The purpose in this case was to produce xylanase as an additive for animal

feed.

5.3 TSP must be extracted prior to pretreatment

It was originally proposed that hydrolysis enzymes be produced in biomass crops
such as maize and switchgrass; then the plants could be subjected to pretreatment, and no
additional enzymes would need to be added during the conversion because they would be
already present. However, it was found that one of the mildest pretreatments available,
Ammonia Fiber Explosion (AF EX), reduced the activity of E1 by about two-thirds
(Teymouri et al. 2004), and this approach was abandoned. The new idea is to grow the

enzymes in biomass crops, which have the potential to produce large amounts of protein

15

due to a great amount of plant tissue, then extract the TSP and use it in the conversion
step. Any plant matter (rice straw, switchgrass, wheat straw, etc.) can be used in
conversion, including the plants used for growing the enzymes, after undergoing a

pretreatment process.

5.4 T hermostable enzymes are desirable

Thermostable enzymes from thermophilic microbes are usually stable at high
temperature and pH (Bruins et al. 2001) and are thus favorable in industry applications
requiring high temperatures to diminish contamination by unwanted microbes. They have
the additional advantage of being less active at ambient temperatures and becoming
activated upon heating; in the case of cell-wall-degrading enzymes, controlling the
activity is important for enzyme production in plants to avoid degradation of the plant

before extraction.

5 .5 Subcellular targeting and sequestration

Proteins can be targeted to various subcellular compartments, such as the
endoplasmic reticulum (ER), chloroplast, vacuole, or mitochondria, with the use of
targeting and/or retention signal peptides. These compartments house various
environments that make them desirable for expression of different proteins. Subcellular
localization also plays a crucial role in affecting the output of foreign proteins by
controlling the interconnected processes of folding, assembly and post-translational
modification.

Experiments have compared the targeting of antibodies to the secretory pathway

rather than the cytosol and have shown it to be generally more advantageous and superior

16

for folding, assembly and high-level accumulation (Schillberg et al. 1999). In the
secretory pathway, proteins first accumulate in the ER and those without a retention
signal (H/KDEL carboxy-terminal tetrapeptide tag; Conrad and F iedler 1998); are
secreted to the apoplast. The ER supplies an oxidizing environment and a profusion of
molecular chaperones, with few proteases (Fischer et al. 2004). These qualities are
probably the most critical influences on protein folding and assembly. In particular, the
molecular chaperone BiP has recently been shown to interact specifically with antibodies
in transgenic plants that are targeted to the secretory pathway (Nuttall et al. 2002).
Proteins are less stable when they are secreted rather than retained in the lumen of the
ER; therefore higher accumulation is possible, generally 2-10 fold greater (Schillberg et
al. 2003).

It would be strategic to confine the enzymes to different cellular compartments or
grow separate enzymes in separate plants. During TSP extraction of enzymes confined to
separate cellular compartments, the enzymes would be mixed; if grown in separate plants,
the TSP of different plants could be combined before the conversion step. Physical
separation of the enzymes will also help avoid enzyme activation until required, as
multiple enzymes are needed for synergy to complete hydrolysis, as explained above.
Also, if the enzymes are sequestered in various compartments they will not have access
to their substrate. In the same vein, if the heterologous protein is toxic to the plant, for
example, as avidin produced in the cytosol of tobacco plants, targeting to an organelle (in
this case, the vacuole) can allow its accumulation without damaging the host cell (Murray
et al. 2002). Several microbial hydrolysis enzymes have already been successfirlly

expressed in plants with subcellular targeting (Table 1).

17

Table 1. Subcellular targeting of various hydrolysis enzymes.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Gene Plant Target Reference

E 1_ C A T Arabidopsis Apoplast Ziegler et al. 2000

E 1_ C AT Tobacco Apoplast Ziegelhoffer et a1. 2001

E 1_ C AT Rice Apoplast Oraby et al. 2007

E 1_ C AT Maize Apoplast Biswas et al. 2006; Ransom et al. 2007

E1 Potato Apoplast Dai et al. 2000

E1 Potato Chloroplast Dai et al. 2000

E1 Potato Vacuole Dai et al. 2000

E1 Tobacco Apoplast Ziegelhoffer et al. 2001; Dai et al.
2005

E1 Tobacco Cytosol Ziegelhoffer et al. 2001; Dai et al.
2005

E1 Tobacco Chloroplast Dai et al. 2000; Ziegelhoffer et al.
2001; Dai et al. 2005

E1 Tobacco ER Dai et al. 2005

E1 -cd Tobacco Apoplast Zfigelhoffer et al. 2001

E 1 -cd Tobacco Cytosol Ziegelhoffer et al. 2001

E1 -cd Tobacco Chloroplast Ziegelhoffer et al. 2001

E2 Tobacco Cytosol Ziegelhoffer et al. 1999

E2 Potato Cytosol Ziegelhoffer et al. 1999

E2 Alfalfa Cytosol Ziegelhoffer et al. 1999

E3 Tobacco Cytosol Ziegelhoffer et al. 1999

E3 Potato Cytosol Ziegelhoffer et al. 1999

E3 Alfalfa Cytosol Ziegelhoffer et al. 1999

CBHI Tobacco Cytosol Dai et al. 1999

Human acid 30 Tobacco Cytosol Reggi et al., 2005

Maize 30 Tobacco Chloroplast Kiran et al. 2006

B. fibrisolvens Tobacco Vacuole Yao 2004

H17c 30

B. fibrisolvens Maize Vacuole Ransom 2007

H17c BG

xynA Barley Cytosol Patel et al. 2000

xynZ Tobacco Apoplast Herbers et al. 1995

XynA-cd Tobacco Cytosol Kimura et al. 20033

X nA-cd Rice Cytosol Kimura et al. 2003b

xynB Potato Cytosol Yang et al. 2007

{ynB Potato Apoplast Yang et al. 2007

 

The apoplast is a popular and excellent compartment for targeting because it is

spacious and can thus accumulate large quantities of foreign proteins (Ziegler et al.

18

 

2000); for E1 endoglucanase and possibly other cellulases, its pH is also similar to its
native source, A. cellulolyticus, 5.5-5.6. However, for the reasons outlined above, the ER
may be an even better localization goal.

The plant could have a suite of hydrolysis enzymes selected based on optimal
synergy, each targeted to a different compartment (apoplast, chloroplast, ER,
mitochondria, vacuole, etc.). Alternatively, if the same enzyme were targeted to several
compartments in the same plant, production could be maximized (Ragauskas et al. 2006).
Recently, this approach was investigated with xylanase targeted to both chloroplasts and

peroxisome (Hyunjong et al. 2006).

6. Other Approaches

To be economically viable as a technology, plant-produced hydrolysis enzymes
must be less expensive than those produced in microbes while retaining the same activity.
It would be ideal if the plants used to produce the enzymes had less lignin or were more

amenable to pretreatment, requiring less pretreatment and/or chemicals.

6.] Microbial engineering

Researchers have been steadily increasing both the efficiency of production and
the activity of the enzymes using synthetic enzymes and engineered microbes, resulting
in a dramatic decrease in the cost of cellulase production in microbes (Knauf and
Moniruzzaman 2004; Ragauskas et al. 2006). Microbial molecular geneticists also have a
goal to produce designer microbes that secrete all the necessary hydrolysis enzymes and

are able to use all the resulting sugars as a feedstock for fermentation in an effort to

19

achieve “consolidated bioprocessing” (Lynd et al. 2005). This technology features
cellulose production, hydrolysis and fermentation in one step. Ideally, the enzymes
produced by these organisms would be modified to have optimized activity both
individually and in specific combinations for maximum synergy. Although potentially

feasible, this technology has not been realized to date.

6.2 Lignin pathway manipulation

As pretreatment is still a costly necessity of the cellulosic ethanol process,
reducing or eliminating this need is a key goal. One way to achieve it could be reduction
of lignin or modification of its structure in the feedstock biomass (Ragauskas et al. 2006).
Lignin contains few components, so regulation of its pathway genes should be relatively
straightforward. It is derived from three precursors, para-courmaryl, coniferyl and sinapyl
alcohols, which are synthesized in separate but interconnected pathways that are also
involved in other cellular functions, including defense (Ragauskas et al. 2006). Other
industries are interested in modifying plant lignin for additional purposes, such as
increasing digestibility, decreasing bleaching necessity and reducing chemical usage
(Boudet 2000; Dean 2004; Ralph et al. 2006; Ralph 2006).

Down-regulation of a major lignin pathway gene, 4-coumarate 3-hydroxylase
(C3H), in alfalfa (Medicago sativa) resulted in a dramatic shift in the lignin profile and
consequent altered lignin structure that were speculated to explain an earlier study
(Reddy et al. 2005) reporting improved digestibility of C3H-deficient alfalfa lines in
ruminants (Ralph et al. 2006). Down—regulation of cinnamyl alcohol dehydrogenase
(CAD) in alfalfa also resulted in increased in situ digestibility and modified lignin

composition although no overall reduction in lignin content (Baucher et al. 1999).

20

Suppression of O-methyl transferase (OMT) in tobacco (Nicotiana tabacum) resulted in
increased biomass production and a shift from the structural to non-structural fraction in
biomass partitioning, but again no overall decrease in lignin content (Blaschke et a1.
2004). Tobacco with reduced CCR showed a decrease in lignin and also an increase in
xylose and glucose associated with the wall (Chabannes et al. 2001).

Experiments in Populus spp. have studied alteration in lignin composition as well.
When cinnamoyl CoA reductase (CCR) was down-regulated in poplar, Clostridium
cellulolyticum was better able to digest the polysaccharides, and twice as much
fermentable sugar was released (Boudet et al. 2003). Down-regulation of 4—courmarate
CoA ligase (4CL) in P. tremuloides resulted in a 45% decrease in lignin and a
corresponding 15% increase in cellulose (Li et al. 2003; Hu et al. 1999) which was
enhanced even further (52% less lignin and 30% more cellulose) when coniferaldehyde
S-hydroxylase (CAldSH) was also present (Li et al. 2003). Down-regulation of CAD
resulted in improved lignin solubility in alkaline medium, allowing easier delignification;
this could decrease costs associated with pretreatment because fewer chemicals are
needed (Pilate et al. 2002).

While these results are encouraging, it is important to remember that lignin is
essential for structure and for defense against pathogens and insects. Alteration of lignin

structure and amount must be done without sacrificing vital needs of the plants involved.

6. 3 Up-regulation of cellulose pathway genes to increase sugar content

Several research groups are actively involved in elucidating cellulose synthesis in
plants (Kawagoe and Delmer 1997; Arioli et al. 1998; Bolwell 2000; Persson et al. 2005;

Andersson-Gunneras et al. 2006; Haigler 2006). So far, most modifications in the

21

pathway genes have been used in basic research in the study of cellulose synthesis.
However, manipulation of pathway component genes to increase polysaccharide content
for the purposes of improved feedstock production holds promise and should be an area

of greater research activity.

6.4 Delayed flowering to increase biomass

Increasing feedstock biomass is one way to increase the amount of cellulose
available for hydrolysis and fermentation. A promising strategy is to increase the duration
of the vegetative state, which can be achieved by engineering plants that have delayed
flowering. A floral repressor gene identified in Arabidopsis, FLOWERING LOC US C
(FLC), maintains a vegetative state unless the plant is exposed to vemalization; in this
event, the gene is turned off and the plant flowers (Sheldon et al. 1999; Michaels and
Amasino 1999). This gene was used to engineer tobacco, with a successful late-flowering
result, and a concomitant increase in biomass (Salehi et al. 2005). Accordingly, work is
in progress to test expression, delay in flowering and increase in biomass in the biomass

crop maize (Sticklen laboratory, unpublished).

6.5 Genetic manipulation to increase biomass

Other ways to increase plant biomass could include modification of plant growth
regulators such as gibberellins. Hybrid poplar displayed improved growth and biomass
when gibberellin biosynthesis was increased (Eriksson et al. 2000). Another strategy

could be to adjust the plant’s physiology to harness or enhance biological functions such
as rate of photosynthesis (Richards 2000); uptake of C02, nitrogen and other resources;

utilization of nutrients, oxygen and water; respiration; synchronization of circadian clock

22

and external light-dark cycle (Dodd et al. 2005); and carbon allocation (Luo et al. 1997).
These processes could be augmented through genetic manipulation of the plants, which
has the potential to boost growth and thus biomass production. In a study on
transformation of rice to increase endosperm activity of ADP-glucose pyrophosphorylase
(AGP), a key enzyme in starch biosynthesis, an unexpected 20% increase in plant

biomass was observed (Smidansky et al. 2003).

7. Conclusion

The US needs a competitive substitute for fossil fuels; ethanol biofuel is an
attractive choice in a suite of alternatives. It is more environmentally benign and because
its use can decrease dependency on foreign oil. It can be produced from starch or
enzymatic hydrolysis of cellulosic biomass. Production of ethanol involves fermentation
of sugars, which can come from plants as simple sugars, starches, or complex structural
polysaccharides of the plant cell wall.

Several microorganisms and the enzymes they produce have been studied for their
ability to degrade cell walls. The enzymes necessary for this process include cellulases
(endo- and exo—glucanases and B-glucosidases) and hemicellulases (most importantly
xylanases). Ligninases are also produced by some organisms but are less well understood.

Biomass ethanol production involves several steps: production of hydrolysis
enzymes; biomass pretreatment; enzymatic hydrolysis; fermentation of the resulting
sugars and distillation. Although biomass is abundant, it has not been traditionally used
because of the costs involved in pretreatment and enzyme production. There are few pilot

facilities at this stage, but more are planned. Roadblocks stand in the way of this

23

technology becoming mature and economically feasible, including the high costs of
enzymes and pretreatment.

Pretreatment is necessary because the cellulose is locked away inside microfibrils,
which are wrapped in hemicellulose and lignin, making it inaccessible to hydrolysis
enzymes. Current pretreatments include dilute acid, flow-through, ammonia fiber
explosion (AFEX), ammonia recycle percolation, steam water explosion, lime, and
organosolv pulping.

Cellulases break down cell walls in a three-step process: endoglucanase randomly
cleaves crystalline regions; exoglucanase (also called cellobiohydrolase) breaks these
chains down further into cellobiose (two glucose molecules) units; and B-glucosidase
completes the reaction by releasing the single glucose units from cellobiose.

Cellulase enzymes are produced in microbial fermentation tanks, but could be
produced in crops to reduce costs. An ideal candidate is the biomass crop maize: it
produces a large amount of biomass, is already widely grown and used as a biofactory for
enzymes and other industrial products, and systems for production and distribution are
already in place. In addition, enzymes could be targeted to different subcellular
compartments to produce a suite of cellulase enzymes in the same plant, optimize
expression or avoid hydrolysis until required. In addition to using crops as enzyme
biofactories, other solutions to the problems mentioned above include reducing or
altering the plants' lignin content to make it more amenable to pretreatment, and
microbial engineering. Genetic approaches for increasing biomass include up-regulation
of cellulose pathway enzymes, delay in flowering, and manipulation of plant growth

regulators. Several of these research avenues will need to be combined before the

24

technology can be perfected. However, the major landmarks reached to date indicate that
the transition from fossil fuels to biofuels is achievable.
The objectives of this work were to:
1) Produce the cellulases endoglucanase El from Acidothermus cellulolyticus and
B-glucosidase from Butyrivibriofibrisolvens H17c in transgenic maize.
2) Perform molecular analyses to show that these enzymes were produced in the
plants.
3) Test the enzymes’ activities and test their ability to convert cellulose and
APEX-pretreated biomass to fermentable sugars.
4) Combine plant-produced cellulases to test ability to convert cellulose and
AF EX-pretreated biomass to fermentable sugars.
These experiments will confirm the feasibility of producing biologically-active
hydrolysis enzymes in the crop plant maize and demonstrate their usefulness in

conversion of lignocellulosic biomass to fermentable sugars for reducing costs associated

with ethanol fuel.

25

11. MATERIALS AND METHODS

1. Transformation Vectors

1.] Genes of Interest

pMZ766-EICAT. Vector pMZ766-E1C AT (Ziegler et al. 2000) encodes the

catalytic domain of endo-l,4—B-glucanase E1 from A. cellulolyticus, targeted to the
apoplast with the signal peptide from tobacco pathogenesis-related protein la(Pr1a),
under regulation of the CaMV 35S promoter, the tobacco mosaic virus translational

enhancer (Q), and the polyadenylation signal from the nopaline synthase gene (3’ nos)

(Figure l).

Sphl BamHI
HindIII Sbfl

Xbal NCO] EcoRV SaII Sacl Pstl SalI SacI Xbal

 

 

 

 

l l I l l L
‘i CaMV 353 Q m. EI-cat lnos WC"

 

 

 

 

 

 

SP
0.8 kb 0.2 kb 0.09 kb 1 kb 0.3 kb
4___./

V

2.36 kb

Figure 1. pMZ766-ElCAT. CaMV 3SS: Cauliflower Mosaic Virus (CaMV) 35S
promoter; Q: tobacco mosaic virus (TMV) translational enhancer; PrlaSP: tobacco
pathogenesis-related protein (apoplast targeting signal); E I-cat: coding sequence of the
catalytic domain of endo-1,4-B-gluccanase E] from A. cellulolyticus; nos:
polyadenylation signal from the nopaline synthase gene.

pUC1813. Vector pUC1813 (Yao 2004) contains the Cauliflower Mosaic Virus

(CaMV) 358 promoter, endoplasmic-reticulum leading sequence (ER); ngA gene

26

encoding Butyrivibriofibrisolvens H17c B-glucosidase, a vacuole-targeting sequence

(VT) and the CaMV 35$ terminator (Figure 2).

30915le M Sal EmRISacI

 

 

 

 

I I} Cauvass ERIbglAVT 3584 1' I 9001813

 

 

 

Figure 2. pUC1813. CaMV 358: Cauliflower Mosaic Virus (CaMV) 35S promoter; ER:
endoplasmic-reticulum leading sequence; bglA: gene encoding B-glucosidase; VT:
vacuole-targeting sequence; 3SS-t: CaMV 35S terminator.

1.2 Selectable Markers

pBY520. Vector pBY520 (Xu et al. 1996) contains the barley H VAI coding
sequence regulated by the rice actin 1 (Actl) promoter and potato proteinase inhibitor 11
(pinIl) terminator, as well as the bar coding sequences regulated by the CaMV3SS

promoter and nos terminator (Figure 3).

 

 

 

 

l l
4—l|m|m|ph ]-|-|353|bir|ml->W
Mb 1.05 0.95 0.85 056150.35

Figure 3. pBY520 (Xu et al. 1996). Act1-5’2 rice actin 5’ region (promoter); HVAI:
barley H VAI gene; pinll: potato proteinase inhibitor 11 terminator; 358 5’2 Cauliflower
Mosaic Virus 35S promoter; bar: bar gene encoding Bialaphos herbicide resistance; nos-
3’: nos terminator.

pDM302. Vector pDM302 (Cao et a1. 1992) contains the bar coding sequence

under the control of the Act] promoter and nos terminator (Figure 4).

27

 

 

SaII BamI-II SaII EooRI
HindIII, SmaI, BamHI
EooRI, 20ml, BamHI,
SmaI Smal, 8511

Figure 4. pDM302. Actl-S’: rice actin (Actl-S’) 5’ region (promoter); bar: bar gene
conferring Bialaphos herbicide resistance; nos: nos terminator.

—|-L Actl-S’ * bar
Ech I | l

20:01,

Hindu]

pGreen. Vector pGreen (Hellens et al. 2000) is a binary vector; it contains the bar
gene regulated by the 35S promoter and nos terminator and the FLOWERING LOCUS C
(FLC) gene regulated by the 358 promoter and nos terminator. In addition it has T-DNA

left and right borders and carries the nptII gene for bacterial resistance to kanamycin

(Figure 5).

T-DNA
III/ |
3581 bar/ lHnos H358

0.8 i056 0.3 0.8

 

 

 

 

Figure 5. pGreen. LB: T-DNA left border; 35S: Cauliflower Mosaic Virus (CaMV) 358
promoter; bar: bar gene conferring resistance to Bialaphos; nos: nos terminator; FLC:
FLOWERING LOCUS C (FLC) gene; RB: T-DNA right border; nptII: gene conferring
bacterial resistance to kanamycin.

28

2. Maize Transformation, Acclimation and Care

Highly proliferating, immature-embryo-derived Type II embryogenic callus
(Armstrong et al. 1991) was used in transformation experiments. Two to four hours prior
to bombardment, callus was transferred to 1.5-cm circles in the center of a Petri dish,
containing an osmotic (Vain et al. 1993; conditioning) medium. Conditioned callus was
bombarded with ethanol-washed tungsten particles combined with a total of 10 pg of 1:1
mixture of two plasmids (one containing the gene of interest, the other containing the
selectable marker gene), according to the manufacturer’s protocol (BioRad PDS
1000/He® Biolistic gun) at a pressure of 1100 PSI.

The bombarded callus was kept on the same conditioning medium for 24 hours,
transferred to callus proliferation medium (Armstrong and Green 1985; Armstrong et al
1995; Chu et al. 1975) for five days, and then placed on selection medium containing 2
mg/L Bialaphos, where they were maintained for six to eight weeks with biweekly
subcultures onto fresh medium. All cultures were maintained in the dark. The detected

Bialaphos-resistant surviving callus clones were placed in regeneration medium (Biswas

et al. 2006) and exposed to continuous light (60 pmol quanta m-2 - s-1 from cool-white

40 W Econ-o-watt fluorescent lamps; Philips Westinghouse, USA) for four to six weeks.
Plantlets were transferred further to rooting medium containing 2 mg/L Bialaphos
selectable herbicide (Biswas et al. 2006), and maintained for two to four weeks under the
above light conditions.

Rooted plantlets 8-10 cm in height were transferred to pots containing soil. Pots
were kept covered with plastic bags to maintain humidity, and acclimated in the growth

chamber. When plants showed new growth, they were transplanted to 2- or 5-gallon pots

29

and either kept in the growth chamber until maturity or transferred to the greenhouse

under similar conditions.

Fertile To plants were self— or cross-pollinated. In some cases, transgenic ears

were pollinated with wild type pollen due to lack of sufficient transgenic pollen. Plants
were allowed to mature and seeds were harvested after dry-down when the abscission

layer had formed, 35-45 days after pollination.

3. DNA Analyses

3 . 1 Extraction

Genomic DNA was extracted from leaf tissue with C-TAB as described (Saghai-

Maroof et al. 1984).

3.2 PCR

E1. The oligonucleotide primers 5’-GCG GGC GGC GGC TAT TG-3’ and 5’-
GCC GAC AGG ATC GAA AAT CG-3’ were designed, synthesized and used to amplify
a 1.0 kb fragment spanning the catalytic domain of the endo-l,4-B-endoglucanase gene.

BG. The oligonucleotide primers 5’ GCA TTG ATC TAG AAT GGA GAA ATG
GGC AAG AAT 3’ (left) and 5’ AAT AAT AGT CGA CAG CGG CTT TGA GCT TAG
TCG 3’ (right) were used (Yao 2004); they amplify the entire bglA coding sequence, 2.6
kb. The conditions used in PCR were as follows: 30 cycles of 94°C for 30 sec, 68°C for
1.5 min, and 72°C for 2 min.

H VA 1 . The oligonucleotide primers 5’-TGG CCT CCA ACC AGA ACC-3’

(forward) and 5’-ACG ACT AAA GGA ACG GAA AT-3’ (reverse) (Oraby et al. 2005)

30

were used to amplify a 0.7 kb fragment of the H VAI gene. The PCR conditions were as
follows: 94°C for 3 min; 35 cycles of 94°C for 45 s, 56°C for 45 s, and 72°C for 45s; and
72°C for 10 min.

The PCR products were analyzed by electrophoresis in 0.8% agarose gels

containing ethidium bromide, and visualized under a UV light.

3.3 Southern blots

General procedure: Genomic DNA was digested with appropriate restriction
endonucleases (see specific entries below) and fractionated on a 1% agarose gel. The gel
was depurinated, denatured and neutralized, and blotted onto a Hybond-N+ nylon
membrane (Amersham-Pharmacia Biotech, Buckinghamshire, UK) according to the

manufacturer’s instructions.

E1. Five micrograms of genomic DNA and 10 pg plasmid DNA (pMZ766-

Elc AT) were digested with HindIII or Sacl. Non-radioactive labeling and detection were

carried out with a probe representing the El-C AT coding region.

BG. Thirty micrograms of genomic DNA and 1 ng plasmid DNA (pUC1813)
were digested overnight with either BglII or NcoI, which cut the construct once (Ncol
cuts at the beginning of the CaMV 35S promoter; BglII cuts at the end of the ngA coding
sequence). Radioactive labeling and detection were carried out with a probe representing
the ngA coding region, generated by PCR. Copy number was determined by counting the

resulting bands.

31

4. RNA extraction

Total RNA was extracted using TRIZOL® Reagent (Invitrogen Corporation,

Carlsbad, California 92008, Cat. # 15596-026) as specified by the manufacturer.

5. Northern blots

Twenty pg RNA were separated on 1.2% (w/v) agarose-formaldehyde denaturing
gels (Sambrook et al. 1989) and blotted onto Hybond-N+ nylon membranes (Amersham-

Pharmacia Biotech).

6. Labeling, hybridization and detection for Southern and northern blots

For non-radioactive labeling and detection, the PCR DIG Probe Synthesis Kit
(Roche Applied Science, Penzberg, Germany, Cat # 11 636 090 910) and/or DIG High
Prime Labeling and Detection Starter Kit 11 (Roche Applied Science, Penzberg,
Germany, Cat # 11585614910) was used according to the kit’s instructions to generate a
probe labeled with digoxigenin-dUTP. Probe hybridization and immunological detection
were carried out using the DIG High Prime DNA Labeling and Detection Starter Kit 11
(Roche Applied Science, Penzberg, Germany, Cat # 11585614910) with the instructions

therein.

Radioactive probe labeling was achieved with a-[32P]-dCTP (GE Healthcare)

with the Random Primers DNA Labeling Kit (Invitrogen, Carlsbad, CA, Cat. # 18187-
013) according to the kit’s instructions. For hybridization, PerfectHbeM Plus

Hybridization Buffer (Sigma-Aldrich, St. Louis, MO 63178; Cat. # H7033) was used at

32

62°C according to the instructions. Detection was done according to standard procedures
(Sambrook et al. 1989). Blots were exposed to X-ray film and developed in a Kodak RP

X-OMAT Processor.

7. Extraction of TSP

E1. TSP was extracted from leaf tissues as described (Ziegelhoffer et al. 2001).
Briefly, 100 mg fresh leaf tissue was ground in the sodium acetate grinding buffer and
precipitated with saturated ammonium sulfate. Extracts were quantified using the
Bradford method (Bradford 1976) using a standard curve generated from bovine serum
albumin (BSA). For the large-scale TSP extraction (to check the activity on biomass), an
automatic solvent extractor (Dionex) was used. To a total of 9 gm pulverized transgenic
maize residue, 60 ml grinding buffer was added and used by the machine to extract TSP.
The extracted TSP were precipitated by adding an equal volume of saturated ammonium
sulfate and allowing to stand overnight at 4°C. The precipitated TSP was collected by
centrifirgation and concentrated by re-suspending in 5 ml grinding buffer. This TSP
concentrate was measured for activity (described below) and used without any firrther
dilution.

BG. TSP was extracted from leaf tissues as described (Carrao-Panizzi and
Bordingnon 2000) and quantified as above. Briefly, extraction buffer (0.05M citrate
buffer, pH 4.8, 10% glycerol, protease inhibitor cocktail (Sigma-Aldrich, St Louis, MO,
Cat. # P9599; or Complete, Mini, EDTA-free Protease Inhibitor Cocktail Tablets, Roche

Applied Science, Indianapolis, IN, Cat. # 11836170001) was added to pulverized (via

liquid N2) leaf tissue in approximately a 2:1 ratio to achieve a viscous slurry. Samples

33

were incubated at room temperature for 1 hr, followed by centrifugation at 15,000xg for 5

min; the supernatant was collected and used in subsequent analyses.

8. Activity Assays

8.1 MUCase Activity Assay for E1
E1 activity was assessed as described (Ziegelhoffer et al. 2001). Briefly, a series

of soluble protein dilutions ranging from 10'1 to 10‘3 were developed, representing

concentrations of 0. 1-10 ng/pl. In a 96-well plate, 10 pl samples (representing 1-100 ng
TSP) was mixed with 100 pl reaction buffer containing 4-methylumbelliferone B-D-
cellobioside (MUC). The fluorophore 4-methy1umbelliferone (MU), as the product of E1
hydrolization of the substrate MUC, was measured as follows. Plates were covered with
adhesive lids and incubated at 65 °C for 30 minutes. The reaction was stopped with the
addition of the stop buffer, and the fluorescence was read at 465 nm using SPECTRAmax
M2 device (Molecular Devices Inc., Sunnyvale, CA) at an excitation wavelength of 360
nm. After subtracting background fluorescence contributed by the control, activity of
each sample was calculated using a standard curve representing 4 to 160 pmol MU and

compared to the activity of pure E1 reported in Ziegelhoffer et al. (2001).

8.2 IUPA CAssay for 80

BG activity was measured as described (Ghose 1987). TSP from plants was added
to assay buffer (0.5M citrate buffer, pH 4.8; 0.015M cellobiose) and incubated at 50°C
with gentle agitation on a rotary shaker (90 rpm). Samples were taken at different times

depending on the experiment (see BG RESULTS).

34

8.3 p-Nitrophenol (pNP) Assay for B G

BG activity was determined by measuring the hydrolysis of p-nitro-phenyl-B-D-
glucopyranoside (pNPBG), slightly modified as follows from the procedure described by
Cai et al. (1999). The incubation mixture was made of 2 mM pNPBG, 50 mM sodium

phosphate buffer (pH 6.5) and 30 pl TSP in a total volume of 100 pl. The reaction was

carried out at 40°C for 15 min and terminated by the addition of 300 pl 1.0 M NazCO3.

The amount of p-nitrophenol (pNP) released was determined spectrophotometrically by
measuring the absorbance of the solution at 415 nm. Standards between 0-100 nmol pNP
were also included. One unit of enzyme activity was defined as the amount of TSP that

produced 1 nmol of product per min under the conditions of the assay.

8. 4 DNS Assay for Reducing Sugars

DNS is a colorimetric reagent that detects reducing sugars, and has been used in
standard assays (Miller 1959; Decker et al. 2003). A protocol for using DNS to test
hydrolysis of commercial enzyme preparations in a microplate was developed in the Dale
laboratory. After a hydrolysis step with Avicel, CMC, xylan, or other substrate, 50 p1
samples were taken in a new plate, 100 pl DNS was added, the color developed at 100°C
for 30 min, and a reading taken with a 100-pl sample using a UV spectrophotometer at
540 nm. The readings were compared to glucose standards and the glucose released as
well as percent conversion calculated. The assay calculations were standardized for a 4%
conversion (IUPAC method), so the amount of enzyme had to be diluted in order to

achieve this.

35

8.5 Glucose analyzer

Samples were put in 1.5 ml microfuge tubes and placed in the glucose analyzer
turntable; the machine took the readings directly. At least 500 pl salmple is needed; if
there was not enough sample, it was diluted by half and the numbers adjusted

accordingly.

9. Western Analysis

9.1 General Procedure
For Western blotting, the Invitrogen NuPAGE® Bis-Tris Discontinuous Buffer
System with a 10% NuPAGE ® Novex Bis-Tris Pre-Cast Gel was used (Invitrogen,

Carlsbad, California). One microgram TSP was run on the gel and blotted onto a

nitrocellulose membrane (Amersham HybondTM ECLTM; Amersham-Pharmacia

Biotech; Buckinghamshire, UK) according to the manufacturer’s instructions. The
membrane was blocked with 1x PBS, 5% non-fat dry milk, 0.1% Tween—20 and
incubated with primary antibody and secondary enzyme conjugate. The Pierce
SuperSignal® West Pico Chemiluminescent Substrate was used for detection following
the manufacturer’s protocol (Pierce Biotechnology, Rockford, IL). The blot was exposed

to X-ray film for one minute and developed in a Kodak RP X-OMAT Processor.

9.2 Specific conditions

E1. The primary antibody was monoclonal mouse anti-E1, 1 pg/ml, courtesy of

National Renewable Energy Laboratory (NERL). The secondary enzyme conjugate was

36

anti-mouse IngHRPO (BD Transduction LaboratoriesTM, BD Biosciences, San Jose,

CA; 1:2000).

10. Pretreatment of Biomass

Milled corn stover (about 1 cm in length) was pretreated using the AF EX
technology (Teymouri et al. 2004). In more detail, the crop biomass was transferred to a
high-pressure reactor (PARR Instrument Col, IL) with 60% moisture (kg water/kg dry
biomass) and liquid ammonia ratio 1.0 (kg of ammonia/kg of dry biomass) was added.
The temperature was slowly raised and the pressure in the vessel increased. The
temperature was maintained at 90 °C for five minutes before explosively releasing the
pressure. The instantaneous drop of pressure in the vessel caused the ammonia to
vaporize, causing an explosive decompression and considerable fiber disruption. The
pretreated material was kept under a hood to remove residual ammonia and stored in a

freezer until further use.

11. Conversion Analyses

11.1E1

El biomass conversion ability was assessed by measuring the reaction of TSP
extracted from El-expressing corn leaves with amorphous cellulose (CMC), crystalline
cellulose (Avicel) and material containing both amorphous and crystalline cellulose, i.e.

APEX-pretreated corn stover (Teymouri et al. 2004).

37

The enzyme hydrolysis was performed in a sealed scintillation vial. A reaction
medium, composed of 7.5 m1 of 0.1M, pH 4.8 sodium citrate buffer, was added to each
vial. In addition, 60 pl (600 pg) tetracycline and 45 pl (450 pg) cycloheximide were
added to prevent the growth of microorganisms during the hydrolysis reaction. The
substrate was hydrolyzed at a glucan loading of 1% (wzv). The TSP from the plant
producing the E1 was concentrated to 1.8% and 250 p1 were added to the substrate. The
reaction was supplemented with 64 pNPGU/g glucan (Novozyme 188 Cellobiase from
Aspergillus niger, Sigma-Aldrich, St. Loius, MO, Cat. #C6105) to convert the cellobiose
to glucose. Distilled water was then added to bring the total volume in each vial to 15 ml.
All reactions were performed in duplicate to test reproducibility. The hydrolysis reaction
was carried out at 50 °C with a shaker speed of 90 rpm. About 1 ml of sample was
collected at 72 hr of hydrolysis, filtered using a 0.2 mm syringe filter and kept frozen.
The amount of glucose produced in the enzyme blank and substrate blank were
subtracted from the respective hydrolyzed glucose levels. Hydrolyzate was quantified
using Waters HPLC by running the sample in Aminex HPX-87P (Biorad) column,

against sugar standards.

I 1.2 Microplate Hydrolysis

Hydrolysis reactions were performed in 96-well microplates. Each well contained
a steel bead, 418 p1 substrate (0.5 or 1% CMC, Avicel or APEX-treated corn stover
(AC8)), 25 pl 1 M citrate buffer and 38 pl enzyme dilution (or plant TSP) plus water for
a total volume of 500 pl. Plates were covered with foil tape and incubated at 50 °C, 350

RPM for the time specified.

38

12. Progeny Analyses
T1 seeds were germinated in vitro on 2 mg/L Bialaphos selection medium (Biswas

et al. 2006) to determine segregation ratios of the offspring. Then, PCR analyses using

the same primers and conditions as in the T0 generation were conducted to examine the

presence of transgenes in the progeny.

39

III. RESULTS FOR El

One of the challenges to the use of lignocellulosic biomass as a feedstock for
ethanol fuel production is the prohibitive cost of the enzymes needed for its
saccharification. One proposed solution has been to produce them in plants instead of
microbes. To this end, cellulases from various organisms have been produced in several
plants. Specifically, the thermostable E1 (endoglucanase) transgene from Acidothermus
cellulolyticus (Baker et al. 1994; Tucker et a1. 1989) has successfully been expressed in
several plants, including Arabidopsis (Ziegler et al. 2000), potato (Dai et al. 2000b), and
tobacco (Ziegler et al. 2000; Dai et al. 2000a); however, none of these are sizeable
enough plants to enable large-scale commercial production of enzymes. Maize was

therefore chosen in an attempt to remedy this problem.

1. Transformation
Transformation and regeneration of plants with pMZ766-E1C AT (E1) and either

pBY520 or pDM302 were performed by another researcher (Biswas et al. 2006), who

obtained a total of 9 lines, each with 4-15 plants that survived to maturity.

2. Molecular and enzymatic analyses

In preliminary work (Biswas et al. 2006), integration of the E1 coding sequence

was confirmed via PCR, which showed that 31 plants carried the E1 transgene. Southern

40

blotting further verified the integration of the E1 transgene in these plants (Biswas et al.
2006). Northern blots did not show any hybridization with the probe.

Forty plants of seven lines were tested for activity. Among the 31 PCR positive
plants, 16 showed biological activity compared to control untransformed plants, as
evidenced by percent E1 in plant leaf extract TSP (Table 2). Percentages of E1 in TSP
ranged from 0.01% to 1.16%. The assay was able to detect enzyme activity levels as low
as 0.01% E1. Western blotting confirmed the translation of El, also showing differences
in the production levels (Figure 6). In general, the signal strength observed in the Western
blot corresponded with the percentage E1 observed in activity assays (Figure 6), although
one plant, 7-6, does not show a band. The nine plants (from four different lines) that

showed the highest levels of activity were chosen for further study.

Table 2. Mean enzymatic activity and percentage E1 of total plant soluble proteins

produced by transgenic maize plants (To). N=number of replicates; S.D.=standard
deviation.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Plant (line-plant 2-8 2-5 7-6 1-1 1 1-1 2-3 1-13 1-12 1- 10
number)

% E1 1.16 0.35 0.51 0.27 0.26 0.18 0.05 0.02 0.03
N 5 9 7 6 1 l 3 2 1
SD. 0.147 0.308 0.349 0.156 N/A N/A 0.042 0.006 N/A
Activity 0.464 0.1408 0.202 0.109 0.104 0.072 0.02 0.008 0.012
(nmol/pg/min)

 

 

 

41

 

Figure 6. Western blot of 1 pg TSP from transgenic maize plants expressing E1 (To).
Lanes: +: positive tobacco control; -C: negative maize control (untransformed); 1-9:
transgenic maize plants. Invitrogen Magic MarkTM Western Standard used for size
markings. Percentages El as determined by enzyme activity assay are displayed above
hands (Table 2).

3. Conversion analyses

The hydrolytic conversion of cellulose using the plant-produced E1 was
confirmed by adding transgenic corn TSP to three types of substrates: CMC, Avicel and
APEX-pretreated corn stover. The conversion of cellulose to glucose ranged from 0.18 to
0.47 g/L when transgenic plant TSP concentrate was used on these substrates (Figure 7).
The highest sugar release, with a mean of 0.47 g/L (after 72 hrs), was observed when the

transgenic plant TSP was added to CMC (Figure 7).

42

 

 

0.50 - 0.47

 

 

 

 

 

Avicel CMC ACS

 

 

 

Figure 7. Average conversion of cellulose to glucan using E1 produced from transgenic
maize. The substrates used in the experiment were Avicel, carboxymethyl cellulose
(CMC) and AF EX-treated corn stover (ACS). The enzymatic hydrolysis was done for a
period of 72 h, at 50°C at 90 rpm. T=TSP from transgenic plants; NT=TSP fiom non—
transgenic control plants. Error bars represent standard deviation fi'om the mean.

4. Second generation
To obtain second-generation (T1) transgenic seeds, the plants were self- and/or

cross-pollinated in the greenhouse. The crosses that produced the most seeds included 2-5
x1-10,1-12 x1-10,1-7 x1-13,1-10 x1-13,1-13 x negative control. None ofthe self-
pollinations resulted in seed. These crosses resulted in 15 progeny that survived to
maturity. In addition, another researcher made two successful but not well-documented

crosses; one with an unknown plant of line 2, the other with plant 7-6. It is unknown if

43

these were the females or the males, and which were the other parents, or if they were
selfed. These crosses resulted in 22 progeny (plants).
PCR analysis confirmed the transmission of the E1 gene to the progeny (Figure

8). None of the progeny retained the H VA 1 gene.

W-C123456789101112131415+C

 

Figure 8. PCR analysis of T1 maize plants for E1. W: water; -C: non-transgenic maize
control DNA; 1-15: T1 plants from known crosses; +C: plasmid DNA (pMZ766-E1CAT).

Northern and Western blots of the 22 plants of dubious ancestry showed no
hybridization; they were not performed on the other 15 plants. Activity assays were

performed on the 15 progeny of known crosses (Table 3).

Table 3. Percentage of E1 in second generation (T 1) plants as determined by activity
assay. The first four columns show the parents and % E1, while the last two columns
show the same information for the second generation. 1-15: Second generation E1 plants;
- 2 Activity showed less than negative (nontransgenic) control; HNPC: Nontransgenic
variety ‘Honey ‘1) Pearl’ negative control; N/A: Not applicable; * PCR negative for

pMZ766-E1c AT

lst Gen. S? % E1 (9) lst Gen. 8 % E1 (6‘) 2nd Gen. Plant % E1
' ' lant n=3
1 0.0006
2-5 0.35 1-10 0.03 9* 0.0040
14 0.0131

1-12 0.02 1-10 0.03 8* -

1-7 1-13 0.05

0.0389

 

45

IV. RESULTS FOR BG

The microfibrils in plant cell walls are composed of long chains of cellulose; most
of this is crystalline but there are some amorphous regions as well. An endoglucanase is
needed to randomly cleave the crystalline cellulose to expose the chain ends that the
exoglucanase can work on. Once the exoglucanase has reduced the cellulose to
cellobiose, B-glucosidase (BG) can catalyze the final step for glucose release. BG is a
class of enzyme that breaks [3 1—>4 linkages between glucose molecules. Because of its
vital role in completing the hydrolysis reaction, it is the subject of this set of experiments.
The first goal was to transform maize with a gene encoding BG, recover enzymatically
active protein from the plants, and show that it is able to convert cellobiose to glucose.

A secondary goal was to try to increase the biomass for greater protein production
while simultaneously improving genetic confinement. To this end, the gene
FLOWERING LOC US C (FLC) from Arabidopsis was co-transformed with BG. The FLC
gene delays flowering and prolongs the vegetative state; it has been shown to delay
flowering and increase biomass production in tobacco (Salehi et al. 2005). In addition,
the FLC plasmid, pGreen, contains the linked bar gene for Bialaphos (herbicide)
resistance and thus in addition provides a selectable marker. The focus of this work is

mainly on BG. Supplementary data on FLC experiments are located in Appendix A.

1. Maize plants are transformed with 36 and FLC

A total of 150 plates of immature embryo-derived maize callus were bombarded

with pUC18l3 (containing BG); 140 of these were co-transformed with pGreen

46

(containing bar and FLC) and the remaining 10 were co-transformed with pDM302
(containing bar only). Sixty-three clones resistant to Bialaphos survived the selection

medium. Of these, 34 lines were regenerated into 196 plants (Table 4).

Table 4. Transformation events and regeneration of To maize plants co-transformed with
a combination of pUC1813 (containing 86) and pDM302 (event 11 only; containing bar)
or pGreen (all other events; containing FLC and bar).

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Event Plates Resistant N lines Line N plants per line*
bombarded clones regenerated designations

1 10 0 -- -- --

2 10 4 2 1-2 2, 14

3 9 0 -- -- --

4 6 0 -- -- --

5 5 0 -- -- --

6 5 7 4 3-6 1, 1, 2, 3

7 5 3 2 7-8 9, 9

8 10 0 -- -- --

9 18 24 17 9-25 24, 35, 16, 7, 5, 5, 4,
1, 2, 6, 5, 3,1, 6, 4, 6,
1

10 18 8 5 26-30 1, 5, 5, 3, 1

11 10 0 -- -- --

12 17 3 -- -- --

13 17 14 4 31-34 1,1,1, 5

Total 150 63 34 196

 

 

 

 

 

 

"‘ The number of plants per line refers to the line designations, e.g., from lines designated 3-6, there is one
plant from line 3, one plant from line 4, two plants from line 5 and three plants from line 6.

2. Maize plants are transgenic and express BG

PCR for BG (Figures 9 and 10) was performed on plants from eight lines (1, 2, 3,
4, 8, 9, 10, and 11). Bands were detected in five of these lines (1, 2, Figure 9; 9, 10, 11,
Figure 10) but only three had bands of the correct size, 2.6 kb (1, 2, 10). This shows that
the gene was properly integrated in plants of lines 1, 2 and 10, but it is unclear whether

the plants of lines 9 and 11 have the gene. Plants representing lines 3, 4 and 8 are unlikely

47

 

to be transgenic. However, PCR can be unreliable, so further tests had to be performed to

verify these results.

 

Figure 9. Results of PCR for BG. Individual plants indicated above the lanes are
represented by line-plant number. W: reaction containing only water; 1 kb: 1 kb marker
(NEB); P: pUC1813.

S 9 at" v" e'" .39 tr" 95" c355 9"

 

Figure 10. Results of PCR of maize transformed with BG for the 30 gene. Individual
plants indicated above the lanes are represented by line-plant number. W: reaction
containing only water; -C: Non-transformed negative control; 1 kb: 1 kb marker (NEB);
P: pUC1813.

48

Northern blots for BG (Figure 11) were performed on 12 lines (1, 2, 3, 5, 6, 7, 8,
9, 10, 11, 12, and 26). Ethidium-bromide-stained bands are shown below the blot to show
relative amounts of RNA loaded on the gel prior to transfer; this indicates that there was
reasonably even loading and thus differences in intensity of bands signify differences in
expression. Hybridization with the BG probe was detected in lines 1, 2, 3, 9 and 10,
indicating that these plants are producing 36 RNA (i.e., expressing the gene). Plants 3-1
and 104 did not show amplification by PCR (Figure 10), but 3-1 shows a very weak
signal and 10-4 a strong signal on the northern blot (Figure 11); 3-1 displays a weak
signal on the Southern blot (Figure 12; see next section). In addition, most plants (12 out
of 19; 63%) of line 9 were expressing BG. PCR of plants from line 9 showed smaller than
expected bands, and for two of the plants there were two bands (9-2, 9-4; Figure 10).
Plant 9-2 is expressing 30 RNA; however, plant 9-4 showed the same pattern after PCR
and yet was not expressing BG. As mentioned above, PCR is not always reliable and that
is why it is important to do further testing.

Lines 1, 2, 9 and 10 had representative clones that also did not hybridize with the
probe, suggesting somaclonal variation in the lines or silencing in these particular plants.
Therefore it was necessary to screen all plants and not just representatives, to avoid

missing any potentially superior lines.

49

 

Figure 11. Northern blots of maize expressing BG. Individual plants indicated above the
lanes are represented by line-plant number. Ethidium-bromide-stained RNA bands
corresponding to each of the plants from the gel before are displayed below the blots to
show relative loaded amounts. +C: Positive control (plant 10-1); —C: Untransformed
maize negative control.

50

Figure 1 1, continued.

95%.?3’53 W W5NNDk>Wb
(\u’bx‘b ~%\$N$\%999@@%N¢t.fiw¢5¢b

   

        
  

        

A A A .4. A..

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W5} Ibbbbilvlbyfi’bflo
asaeeeeeee

ON I O a

.. .’\.'\
wtwtrfhfi '0
m *9 ' If

Hflr~r ”'“ulgrusr

pscmsrén

 

51

Southern blots for 80 (Figure 12) were performed on 29 out of 34 lines (all
except 4, 14, 19, 23 and 26). Hybridization with the BG probe was detected in all lines
tested except line 8. Thirty micrograms of genomic DNA and 1 ng plasmid DNA
(pUC1813) were digested overnight with either BglII (Figure 12 A, left panel) or NcoI
(Figure 12 A, right panel). These were the two restriction enzymes to choose from that
had a single cutting site in the construct and were methylation insensitive. Cutting with
NcoI appeared to give clearer results and was used in the rest of the blots.

The blots confirm that these plants have the gene and that they are independent
transgenic lines. Apparent copy numbers of the various lines is consolidated in Table 5.
As mentioned in Materials and Methods, copy number was determined by counting the
bands. Copy numbers are only an estimate; darker bands could represent more than one
copy and the actual copy number may therefore be higher in plants with dark bands. For
some plants (those in Panel B or those that were too dark), it was impossible to quantify
copy number due to blot quality. As there were different numbers of bands and the bands
were different sizes, the gene was integrated into different locations in the genome and
the plants in fact represented independent transgenic lines. The DNA of lines 1 and 2 was
undetectable in the agarose gel prior to transfer, indicating that the DNA quantification
was inaccurate, and thus the amounts of DNA loaded were insufficient to enable

measurable detection.

52

    

Figure 12. Southern blots of maize transformed with BG. Genomic DNA (30 pg) and
pUC 1813 plasmid DNA (1 ng) were digested overnight with [A] Bglll (left panel) or
NcoI (right panel) or [B-E] Neal and fractionated on a 1% w/v agarose gel. Individual
plants indicated above the lanes are represented by line-plant number. 1 kb: 1 kb
molecular weight marker (NEB); P: pUC1813 positive control; -C: Untransformed maize
inbred line Hill. Panels D and E represent the same samples but different exposure times
(8 h and V2 h, respectively).

 

53

Figure 12, continued
:5 :x a :x a :x 2» :x to I: 5L 5‘) 5* :x :5 e
’15“) N" \n’ (5 N? 5°) '0 r3 '3’ r35 '19 ‘19 “LN “If" ’0’ '1. '13" If

  
  

      

'\

    
 

         
    

 

4+3.» .u

0;]!

r mm

54

Table 5. Plant lines (To) and apparent copy numbers based on bands on Southern blots.

 

 

 

 

 

 

 

 

 

 

 

Line Copies Line Copies Line Copies
1 2 12 ? 25 7

2 3 13 7 27 5+
3 5 15 5 28 6

5 ? 16 1 29 5+
6 ‘7 l7 8 30 5

7 ? 18 8 31 ?

8 0 20 9 32 2-3
9 5-6 21 9 33 1
10 4-5 22 8-9 34 4-5
1 1 6-7 24 7 M 4-5

 

 

 

 

 

 

 

 

3. TSP from transgenic 86 plants converts cellobiose to glucose

‘ A preliminary glucose conversion activity assay for BG (Table 6) tested the
plants’ ability to degrade cellobiose. TSP (100 pg) from plants from lines 2, 3, 8, 9, 10,
11 and 26 was added to 0.5M citrate buffer, pH 4.8, with 0.015M cellobiose substrate.
Most plants showed a decrease in glucose after the 30-min incubation at 50°C; however,
all differences were negligible and similar to negative control. A paired t-test indicated no
significant difference between readings at 0 and 30 min. The standard protocol for
determining cellobiase (B-glucosidase) activity of commercial enzymes (Ghose 1987)

was followed.

55

Table 6. Glucose (g/L) released from 0.015M cellobiose after 30 minutes using 100 pg
maize TSP from plants transformed with BG. Hill: Untransformed maize negative
control. The difference between 0 and 30 min was not significant at or=0.05 (t=-2.917,

df=12).

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Line-Plant 0 min 30 min Difference
2-1 0.1 1 1 0.109 -0.007
2-2 0.113 0.117 -0.014
2-4 0.085 0.088 —0.018
3-1 0.085 0.078 -0.014
8-1 0.100 0.086 -0.01
9-4 0.081 0.063 -0.005
9—6 0.084 0.070 +0.002
10-1 0.117 0.107 -0.012
102 0.007 0.065 0015
10-3 0.073 0.075 +0.004
11-1 0.101 0.089 -0.002
26-1 0.081 0.066 +0.004
HiII 0.072 0.076 +0.003
Mean 0.090 0.084 0006
St. Error 0.005 0.005 +0.002

 

A second conversion experiment was carried out with a S-ml reaction using 570
pl TSP, using the same protocol as above, scaled to 5 ml. In this experiment, the TSP was
obtained from 1 g leaf tissue fi'om RNA-positive plants and the resultant amount (mg) is
indicated in Table 7. Plants showing high expression of BG in northern blots (Figure 11)
were chosen for this study.

Samples were collected at several time points: 30 min, 1 h, 7.25 h and 26.5 h
(Table 7, Figures 13 and 14). The data were adjusted by subtracting the substrate blank.
After 30 min, the glucose released in most of the samples had decreased relative to 0 min
(data not shown), similar to the above experiment. After 1 h, an increase in glucose was
observed in most of the samples, but the differences were mostly negligible. After 7.25 h,
differences in glucose release compared to 0 min were becoming substantial. These

results show that 30 min is not enough time to see a conversion response; at least an hour

56

is needed, and 6 or more hours will yield quantifiable results. The longer the reaction
time, the higher the measured glucose release.

The increase in glucose release observed in plants ranged from 3.4 (plant 9-18) to
2.04 (plant 10-13) times higher than that of untransformed control. Glucose release above
0.4 g/L (4% conversion) was observed in 9—18, 9-2, 10-17, 10-24 and 9-9 after 26.5 h.
The standard assay for unit calculation is standardized for a 4% conversion; thus the
reaction (amount of TSP and time) must be optimized. This is an indirect method for
measuring BG activity. The lower panel of Figure 14 also shows relative performance of
the various lines in terms of glucose released (g/L) per mg protein. Although 9-18
performed best, 9-9 and 9-2 actually displayed greater activity, followed closely by 10-24

and 10-13.

Table 7. Glucose (g/L) released from 1% cellobiose at 0 min, 1 h, 7.25 h and 26.5 h,
using 570 pl maize TSP from plants expressing BG. Data have been adjusted by
subtracting the substrate blank. Hill: Untransformed maize negative control. The
differences between 0 min and 1 11, 0 min and 7.25 h, and 0 min and 26.5 h were all
significant at or=0.05 (t=2.667, 6.733 and 8.338 respectively, df=6).

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Total Times

Line- Protein Higher
Plant (mg) 0 min 1 h 7.25 h 26.5 h than Hill
9-18 0.951 0.16 0.20 0.35 0.57 3.41
9-2 0.630 0.24 0.29 0.37 0.51 3.01
10-17 0.968 0.05 0.16 0.28 0.49 2.92
1024 0.610 0.11 0.13 0.26 0.45 2.65
9-9 0.502 0.14 0.17 0.27 0.42 2.47
10-13 0.497 0.14 0.15 0.22 0.34 2.04
Hill 0.570 0.08 0.08 0.11 0.17 1.00
Mean 0.675 0.13 0.17 0.27 0.42

St. Error 0.076 0.024 0.022 0.022 0.03

 

 

57

 

 

 

 

 

 

 

 

 

 

 

 

 

   

9-18 . 9-2 0-17 10-24

 

 

 

 

..P.°
$838
Tl"
1
1l
1
l

'3
l
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l

8
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l
l

Gluceee releeeed (alums protein
0 o p o o o

8

l

L.
O
l

 

 

p
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9-18 9-2 10-17 10-24 9-9 10-13 Hlll
Plente

Figure 13. Top panel: Glucose (g/L) released from 1% cellobiose at 0 min, 1 h, 7.25 h
and 26.5 h, using 570 pl maize TSP from plants expressing BG. Data have been adjusted
by subtracting the substrate blank. Bottom panel: Glucose (g/L) released per mg protein
in TSP. Numbers below the bars represent maize plants expressing BG. Hill:
Untransformed maize negative control.

58

 

0.50 /)
--s1a
--sz
/- --1o17
-1cz4

—9—9

 

s

-o-10-13

0min 1h 7.25h 26.5h
11me point

 

Glucose (gIL)
O O
B 8
\

 

 

 

 

 

0.6 , ----- a

 

Glucose (gIL)
.0 .0 .0
u 3'- 0|

.0
N

 

 

 

 

 

 

 

 

Figure 14. Glucose (g/L) released from 1% cellobiose at 0 min, 1 h, 7.25 h and 26.5 h,
using 570 p1 maize TSP from plants expressing BG. Data have been adjusted by
subtracting the substrate blank. Top panel: Lines represent different maize plants
expressing BG; numbers below the lines represent sample collection time points; Hill:
Untransformed maize negative control. Bottom panel: T: transgenic plants; NT: non-
transgenic plants. Dashed lines represent 95% confidence intervals; solid lines represent
polynomial regression lines; dots represent individual data points. Images in this
dissertation are presented in color.

59

BG enzyme activity was measured with the p-nitrophenol assay and percentage
BG in TSP roughly estimated by comparing with Novozyme 188 (80% BG). The results
are shown in Table 8, and show activities between 0.268 and 5.475 pNPU and 0.15-

3.11% BG in TSP.

Table 8. Mean activity in units pNP (pNPU) and estimated %BG in transgenic plant TSP.

n=3.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Plant pNPU %BG St. Dev.
3-1 5.475 3.11 0.172
9-18 3.840 2.18 0.154
9-2 3 .242 1.84 0.163
2-1 2.768 1.57 0.109
10-24 1.604 0.91 0.073
9-14 1.586 0.90 0.053
10-17 1.563 0.89 0.100
9-9 1.554 0.88 0.037
10-12 1.547 0.88 0.065
10-13 1.193 0.68 0.080
10-30 1.182 0.67 0.066
9-8 0.832 0.47 0.026
9—15 0.805 0.46 0.027
10- 10 0.746 0.42 0.177
10-5 0.747 0.42 0.044
10-14 0.738 0.42 0.151
10-7 0.497 0.28 0.034
9-17 0.445 0.25 0.071
10-6 0.389 0.22 0.031
9-12 0.302 0.17 0.027
10-15 0.290 0.16 0.059
9-23 0.268 0.15 0.009

 

 

60

V. RESULTS OF EXPERIMENTS RELATED TO OPTIMIZING RATIOS OF PLANT-

PRODUCED HYDROLYSIS ENZYMES FOR CONVERSION

1. Introduction.

Enzymes used in the hydrolytic conversion of cellulosic biomass to fermentable
sugars act synergistically (Hoshino et al. 1997), and certain ratios of endoglucanase,
exoglucanase and fi-glucosidase (among other enzymes) are considered ideal (Zhang and
Lynd 2006; Boisset et al. 2001). Commercial cellulose mixtures are often produced from
fungal or bacterial culture media and thus have differing enzyme ratios depending on the
mix that particular species produces (Kabel et al. 2006). Companies such as Genencor are
working on optimizing enzymatic hydrolysis by determining the best available enzyme of
each class produced out of all species (and at times enhancing this activity even further
by creating synthetic genes), and optimizing the ratios of these enhanced enzymes for
maximum sugar production.

Although this is useful for bacterial or fungal enzyme production, plant-produced
enzymes may not behave as expected. Enzyme production is likely to be lower as a
percentage of total soluble protein (TSP) in plants and activity may not be as high as in
commercial enzyme preparations; this could be due to many factors, including low
expression, lack of enzyme purity, glycosylation, truncation or incorrect folding.
Therefore, it is important to test plant-produced enzyme activity and optimize ratios of
TSP necessary for maximum conversion.

Endoglucanase (E1) and exoglucanase (CBHI) were produced in maize and

tobacco by Dr. Chuansheng Mei. Several lines of each have been tested and the activities

61

measured by MUCase assay and conversion experiments (described in Materials and
Methods). Four were considered the best (i.e., had the most activity); three E1 plants and
one CBHI plant. The TSP of these plants was extracted and used in the following
experiments. B-glucosidase (BG) was produced in maize and its activity confirmed
indirectly with the IUPAC method and conversion assays, and directly with the pNPBG
assay (described in Materials and Methods and presented in BG Results).

In this set of experiments, the plant TSP containing the hydrolysis enzymes was
tested alone and in combinations to determine the best mixture to use for biomass
conversion. It was found that the ratio ofE12CBH12BG 1:421 worked best for converting
1% AF EX—treated corn stover to fermentable sugars, and this worked as well as using

commercial Spezyme CP plus plant TSP containing BG.

Materials and Methods for this chapter are listed in the main Materials and

Methods section (Pages 35 and 38).

2. Results and Discussion
2.1. Preliminary Results

2.1.1. Commercial enzyme test plates

First, commercial enzyme preparations of Spezyme CP (a commercial cellulase
mixture containing mostly endo- and exo-glucanases), multifactorial xylanase and
Novozyme 188 (a commercial cellulase mixture containing roughly 80% B-glucosidase)

were tested on a 0.5% CMC plate to test the method (Figure 15).

62

 

1.200

1.000

.0
on
o
0

Sugar (gIL)
§

0.400

0.200

0.000

 

Figure 15. Sugar determination via DNS on a 0. 5% CMC plate. SPC: Spezyme CP;

Sugar Determination on 0.5% CMC Plate

+

if

i

m}—

 

 

711771: i7

 

112001’1 1/1

1.300 1.2 1.12 1.1400113 1.3 1.2500

Dilution of Stock Enzyme

MFX:Multifactorialxylanase;BGzNovozyme188.

The Spezyme dilutions (1:200, 1:300, 1:400 and 1:500) were below 0.2 g/L, and

there was a steady decrease. Multifactorial xylanase also showed a decreasing trend, and

at the 1/3 dilution, was getting close to 0.2 g/L. Undiluted Novozyme 188 showed under

0.2 g/L, 1/2 dilution was over 1 g/L and 1/3 was over 0.7 g/L. The results were not as

expected (Figure 15). The low glucose release of the undiluted enzyme could be either

due to inhibition due to excess enzyme or viscosity that may have caused inaccurate
pipetting.

To test the suitability of the microplate DNS assay for B—glucosidase on

cellobiose, its native substrate, a second assay was performed using Novozyme 188, a

63

 

commercial cellulase mixture that is mostly (around 80%) B-glucosidase (Figures 16 and

 

 

 

 

 

   

 

17).
,3 3 .33 3- i 33 .333 , . 333
l Sugar Determination on 1% Cellobiose Plate
l 0500 -fi~ . - 3333 ~~7 . . 33333. ..
1
1/1 1/2 1/10 1/50 0100 111000
A -0.500 -
.J
l E 1000 l
I h ' . 7
S il
l = i
‘ "’ 4.500- ‘
l
‘ -2.000 .
-2.500 ~ 3-3
Dilution of Novozyme 188
Sugar Determination on 1% Cellobiose Plate
0060 3% 33fi3733 ~ *3 33 333 a:
o 040 - l
1,. 3
3 0020 T‘ l
a l l
.. , g
s . 1
Us, 0000 :1: . . . . . __
1/2 1/10 1/50 1/100 1/1000
-o.020 « I
1 E

 

 

0040 ._333333 ~- .3 ..33_ 33333333
Dilution of Novozyme 188 1

Figure 16. Sugar determination via DNS on a 1% cellobiose plate. Bottom panel:
Undiluted Novozyme 188 is not included to show the other dilutions in more detail.

 

Difference between Novozyme 188 and substrate blank (DNS)

3.500

 

3.000 -

8 1.500 « '“°“°z"“° ’88

 

 

0.000 <
1 2 3 4 5 6

Novozyme 188 dilution

 

 

 

Figure 17. Difference between glucose released in commercial enzyme (Novozyme 188)
vs. substrate background (blank).

The readings were very low after subtracting the blank (Figure 16). It was found
that DNS cannot be used for cellobiose, which is itself a reducing sugar, because the
background is too high for detection of increased sugars (Figure 17): almost 3.5 g/L
glucose, which is nearly 10 times the amount required (4%) for proper determination via
this assay.

To determine if it would be possible to use microplate hydrolysis for Novozyme
188 before DNS color development, samples from the plates after the hydrolysis step
(which had been kept at 4°C) were analyzed for glucose. Figure 18 shows that there is no
trend and the numbers fluctuate around the same amount (8-9.5 g/L) in dilutions 1/1 to

1/ 100. At 1/1000, however, the numbers make a dramatic drop, down to 3 g/L, indicating

65

that dilutions of Novozyme 188 should start at this point and go down from here to

achieve the dilution necessary for 0.4 g/L.

Glucose released by Novozyme 188 dilutions

Glucose (gIL)

1*

1/1 1/2 1/10 1/50 1/100 1/1000
Novozyme188 dilutions i

 

 

 

 

 

 

 

 

 

 

1‘
Figure 18. Sugar determination via glucose analyzer on a 1% cellobiose plate after
hydrolysis.

2.1.2. Plant-produced E1 and CBHl

Dr. Chuansheng Mei had previously determined that four plants out of the ones he
tested showed the highest enzyme activity. Total soluble protein (TSP) of these plants
expressing E1 or CBHI was tested on a 1% CMC plate and, to determine if the plant-
produced enzymes had any activity after 1 hr hydrolysis, samples from the plates were
analyzed for glucose. The readings were very low, the highest being not quite 0.07 g/L,
much lower than the needed 0.4 g/L (Figure 19). Half the readings were higher and half

lower than the enzyme blanks.

66

 

Glucose released (1 h)

 

 

0.08
0.041
e

0-05 ‘ blank
'3
E
' 0.04 - Iadjusted
g CMC (CMC
— blank
0 subtracted)

0.02 4

o .

 

 

 

1.3 E18a 1.4 E1 Ba 1.4 E13b cbh1-7
Plant

 

 

 

Figure 19. Glucose released from 1% CMC after 1 h hydrolysis using TSP from plants
expressing E1 or CBHI.

Since the results of the 1-h hydrolysis were inconclusive, a longer reaction was
attempted. This time, plant TSP was incubated in 1% CMC or Avicel for 24 h at 50°C
and 90 rpm; a 0 time point was also taken. The results (Figure 20), determined via
glucose analyzer, showed that two of the plants’ TSP definitely had activity on CMC (1.3

E1 8a and 1.4 E1 3a); 1.3 El 8a was used in subsequent studies.

67

 

CMC

 

0.16
0.14 -
0.12 —
a 0.1 1
"' 0.08 -
0.06 -
0.04 -
0.02 -

Glueoee

 

 

 

 

0.02 1.3E18_a 1.451 3_a 1.4 E1 30 elm-7

 

Plant

 

Avicel

 

EIO
I24

  

 

 

1.3 E1 0a 1.4 E1 3a 1.4 E13b cbh1-7
Plant

 

 

 

 

Figure 20. Glucose released from 1% CMC or Avicel after 24 h hydrolysis using TSP
from plants expressing E1 or CBHI. Numbers above 24 h bars indicate times higher than
0 h data.

68

The other two had increases after 24 h that were negligible. Regardless, the
conversion did not reach even half of the needed 0.4 g/L. On Avicel, the results were not
encouraging either; while the release was nearly the same as for CMC, 1.3 E1 8a was
considerably less after 24 h than 0 min. The TSP was quite concentrated as well: a total
of273 pg 1.3 E1 8a, 463.6 pg 1.4 E1 3a, 501.6 pg 1.4 E1 3b and 486.4 pg CBH1-7 TSP
were used. In comparison, five out of six of the plants expressing BG had exceeded 0.4
g/L after 24 h on cellobiose with 500-950 pg TSP.

The assay is an initial rate assay and designed to allow calculation of enzyme
activity after only 30 min hydrolysis. Since the plant TSP needs 6-24 h hydrolysis, and
the longer the better, the assay cannot be used for direct calculation of enzyme activity.
However, it can be used for measuring sugars released during hydrolysis. In this case, it
is not necessary to reach 0.4 g/L; any amount is acceptable, since the assay will not be

used to calculate activity, only to measure sugar released.

2. 2. Optimal concentrations of transgenic plant TSP

Another researcher determined that the plant-produced E1 and CBHI described
above work synergistically together best in a ratio of 1:4. The next step was to determine
the relative amount of plant-produced BG to add to this fixed ratio that would give the
best conversion on both pure cellulose and AF EX-treated corn stover. To this end, BG
plant TSP was tested for ability to convert Avicel and CMC, and, keeping the ratio of El
and CBHl 1:4, varying amounts of BG (in terms of percentage of the total of El plus

CBHI) were added to determine the optimal balance.

69

First, enzymes E1 and CBHl were concentrated using a centrifugal concentrator
to a concentration of 4.34 and 5.71 pg/pl respectively. Then, a 24-hour hydrolysis
reaction was done on 1% CMC and 1% Avicel using a 1:4 ratio of E1 to CBHl and
varying amounts of BG relative to E12CBH1: 0.1, .5 and 1. Hydrolysis was performed at
50°C, 90 RPM in a total reaction volume of 10 ml. The amounts used were limited by the
amount of plant TSP available. In this experiment, 30 pg El, 120 pg CBHl, and 15, 75
and 150 pg BG were used. The blanks included were also substrate blank, E1 alone (on
substrate), CBHl alone (on substrate), BGO.1 (15 pg) alone (on Avicel), and BGO.5 (75
pg) alone (on CMC). Three replicates of each sample were included. More TSP blanks
were not possible due to lack of TSP. Sugar release was measured after 24 hours with a

DNS assay (Figure 21) and with the glucose analyzer (Figure 22).

70

 

Sugar release (DNS) 24 h

 

 

 

 

i? w

a BMW
E ICMC
ii.

8

 

 

 

 

Enzymes

 

 

 

Figure 21. Sugar release from 1% Avicel (AV) or CMC after 24 h by application of
various combinations of plant TSP containing hydrolysis enzymes, determined by DNS
assay. E: E1 (30 pg); C: CBHl (120 pg); 801: BG (15 pg); B0.5: BG (75 pg); E:C: E1
and CBHl (1:4 ratio); E:C:BO.1: E1, CBHl and BG (15 pg) (1:4:0.1 ratio); E:C:BO.5:
E1, CBHl and BG (75 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG (150 pg) (124:1
ratio); AV: 1% Avicel; CMC: 1% CMC.

71

 

Glucose released after 24 h (glucose analyzer)

 

0.100
0.080 -

MED J

 

0.040 r

0.020 1

Glucoee (glL)

0.000 -

4.020 ‘

 

 

 

-0.040

 

 

 

 

Figure 22. Sugar release from 1% Avicel (AV) or CMC after 24 h by application of
various combinations of plant TSP containing hydrolysis enzymes, determined by
glucose analyzer. E2 E1 (30 pg); C2 CBHl (120 pg); B0.12 BG (15 pg); 805: BG (75
pg); E:C: E1 and CBHl (1:4 ratio); E:C:BO.1: E1, CBHl and BG (15 pg) (1:4:0.1 ratio);
E:C:BO.5: E1, CBHl and BG (75 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG (150
pg) (1:421 ratio); AV: 1% Avicel; CMC: 1% CMC.

Some general trends can be seen. For the most part, E1 and CBHl alone and
together do not have much activity, releasing below 0.01 g/L sugar after 24 hours (Figure
21), most of which does not appear to be glucose (Figure 22). BGO.1 alone and in
combination with E1 and CBHl is about the same. BGO.5 released much more sugar
alone on CMC than BGO.1 on Avicel; this could indicate endoglucanase activity.
However, this amount was more than when combined with E1 and CBH] , so it could be
an anomaly, or it could be due to inhibition when combined with E1 and CBHI. Also, we

see that as the amount of BG increases, so does the sugar. This could be due to free

72

sugars present in he TSP, but since TSP blanks without substrates had not been included,
it could not be verified. The sugar increases after 24 h on both substrates were more
pronounced when determined via DNS (Figure 21) than with the glucose analyzer (Figure
22), so these free sugars are likely not glucose.

To get a better idea of how much increase was seen, the blanks were subtracted
from the 24-h data (Figures 23 and 24). Please note that because BG blanks included only
BGO.1 on Avicel and BGO.5 on CMC (due to lack of TSP), the BGO.1 blank was used for
E:C:BGO.5 and E:C:BGl on Avicel, and the BGO.5 blank was used for E:C:BGO.1 and
E:C:BGl on CMC. So in the case of Avicel, the E:C:BGO.5 amount is likely lower than
shown; in the case of CMC, the E2C:BG0.1 amount is likely higher than shown; and in

the case of both substrates, the E:C:BGl amount is likely lower than shown.

73

 

Sugar release after 24 hours minus enzyme blanks

 

p
8

EAV
ICMC

3'10" (0M

$3

    

E:C E:C:BOJ E:C:BO.5 E:C:B1

a
g

 

 

 

-o.100

 

 

Enzyme cemblnetlons

 

Figure 23. Average sugar release after subtracting enzyme blanks determined by DNS
assay. E:C: El and CBHl (1:4 ratio); E:C:BO.1: E1, CBHl and BG (15 pg) (1:4:0.1
ratio); E:C:BO.5: El, CBHl and BG (75 pg) (1:4:0.5 ratio); E:C:Bl: El, CBHl and BG
(150 pg) (1:4:1 ratio); AV: 1% Avicel; CMC: 1% CMC. E:C:B0.l for CMC is likely
higher than shown; E:C:BO.5 for AV is likely lower than shown; E:C:Bl for both is likely
lower than shown.

74

 

Glucose released minus enzyme blanks (glucose analyzer}

0.020

 

0.000 -

E2C E:C:Bl

,b
O
8

 

 

DAV I

 

Glucose (91L)
6
E

,b
i

-0.080 -

 

 

 

 

0.100

 

 

 

Figure 24. Glucose release after subtracting enzyme blanks determined by glucose
analyzer. E:C: E1 and CBHl (1:4 ratio); E:C:BO.1: E1, CBHl and BG (15 pg) (1:4:0.1
ratio); E:C:BO.5: E1, CBHI and BG (75 pg) (1:4:0.5 ratio); E:C:BI: E1, CBHl and BG
(150 pg) (1:4:1 ratio); AV: 1% Avicel; CMC: 1% CMC. E:C2B0.1 for CMC is likely

higher than shown; E:C:BO.5 for AV is likely lower than shown; E:C:Bl for both is likely
lower than shown.

After subtracting the blanks, it is clear that glucose release is very low, even with
the highest relative concentration of BG. Total sugars are also low (around 0.15 g/L).
Many of the readings were also negative.

Due to the many problems with this experiment, a second reaction was done. The
BG and E1 TSP had been used in the previous experiment, and the (maize) plants had
since dried down, so TSP was extracted from leaf tissue from entire plants. Although E1

from dried material had been shown to be active in a previous study, this E1 was the

75

result of a different transformation event and subcellular localization (ER vs. apoplast),
and it was unknown whether it or BG would maintain activity after dry-down.

In this experiment, El, CBHl and BG were concentrated to 4.9 pg/pl, 6.6 pg/pl
and 5 pg/pl respectively. A 1:4 ratio of E1 to CBHl was maintained, and 1:0.1, 120.5,
121, and 1:2 ratios of E12CBH1:BG were tested on both 1% Avicel and CMC. As before,
TSP availability determined amount used; 4.6 pg E1, 18.2 pg CBHl, and 2.3, 11.4, 22.8
and 45.5 pg BG were used in a total volume of 500 pl. Three replicates were prepared for
every sample, and data were adjusted by subtracting the substrate blanks. Hydrolysis was
performed at 50°C, 350 RPM. All analyses were performed via DNS assays and a single
timepoint of 18 h was used. Blanks included substrate, enzyme alone in water and
enzyme alone in substrate.

Sugar released after subtracting the blanks is shown in Figure 25 for 1% Avicel
and Figure 26 for 1% CMC. On Avicel, the plant TSP is not contributing at all to the
conversion to sugar, as the values are all negative. The amount of free sugars attributable
to TSP decreases as the amount of BG increases, however, as the numbers become less
negative. This could indicate some BG activity. (On Avicel, only one replicate of E:C:Bl
and two replicates of E:C:BZ were readable by the plate reader.) On CMC, the results are
similar in that they are all negative except for BG2. lBG2 shows a release of around 0.35
g/L, nearly 4%. Perhaps a higher ratio of BG to E12CBH1 is needed to realize maximum
conversion. It appears that the plant-produced E1 and CBHl have very little, if any,

activity.

76

 

 

 

 

 

 

 

 

 

Sugar release on 1% Avicel after 18 hours
0100 ..—. 3__3_ ———-— .. _--*—~_ 33
,1 j T i i : l ' l ‘
-0.100 . E:C' E:C:BO.1 E:C:BO.5 E:C:B1 E:C:BZ
T i i . ' i
. i 2 g
A 0300 « LE2 9
.r ; . _
e i ; J
5 0500 i ' 1
O)
3
m 0.700 .
0.900
-1.100 —- — ~41
Enzyme combinations ”i

Figure 25. Sugar release on 1% Avicel after 18 h; enzyme blanks subtracted. E: C: E1 and
CBHl (1:4 ratio); E. C. BO 1 E1,CBH1 and BG (2. 3 pg) (124201 ratio); E:C:BO.5: E1,
CBHl and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG (22.8 pg) (1:421
ratio); E:C:B2: E1, CBHl and BG (45.5 pg) (1:4:2 ratio).

 

 

 

 

 

 

 

Sugar release on 1% CMC l
0.400 — 1
F17
0.200 ~ l l
A .
g, 0.000 —.— ‘i—T‘T . _._ ,. :3: .. L——1 l
: E20 E:C:BO.1 E:C:BO.5 E:C:B1 E:C:B2
1' , II
3’ 0200 . :
to i g
l l:
-0400 4 r
-0.600 ——-—-- 3-3:
Enzyme combinations ‘

Figure 26. sugar release on 1% CMC after 18 h, enzyme blanks subtracted. E: C: E1 and
CBHl (1 :4 ratio); E: C: B0.21 E1,CBH1 and BG (2. 3 pg) (124'01 ratio); E: C. B0. 5. E1,
CBHI and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: El, CBHl and BG (22.8 pg) (1:421
ratio); E:C:B2: E1, CBHl and BG (45.5 pg) (1:4:2 ratio).

77

The plant TSP was also tested for its ability to substitute for commercial enzymes.
Novozyme 188 was added to plant TSP containing El and CBHl at the same rate as used
in normal conversion experiments using commercial enzymes, scaled to 500 pl. In
addition, the various amounts of plant TSP containing BG were added to the normal
amount of Spezyme CP, scaled to 500 pl. The results are shown in Figures 27 and 28. On
Avicel, results were unreadable for E12CBH12Novozyme 188 and Spezyme CP:BO.1, and
only two replicates were readable for Spezyme CP2BO.5. For this and subsequent
analyses, the contribution of a particular enzyme in a combination was determined by
subtracting the sugar released by the other enzyme(s) from the combined total sugar
released. For example, in a combination of E12CBH1 (1:4), to determine the amount
attributable to E1, the amount of sugar released by CBHl alone would be subtracted from
the amount of sugar released by the combination of E12CBH1. Obviously, this is only an
estimate, and the amounts attributable to individual enzymes will not add up to the
amount released by the combinations due to interactions (either synergy or inhibition or

more complex interactions) or other factors, such as sample variation.

78

 

Sugar Mono on 1% Avicel after 18 hours (commercial

 

 

 

 

 

 

enzymes)

1.000

0.000 4

0.000-

0400 -
5 0200‘ BMTSP+conmuuy.
I Immm
3 0900 - INT?”

0200-
OMW

0M1

 

 

 

-0.aoo

 

 

 

 

Figure 27. Sugar release on 1% Avicel after 18 h; enzyme blanks subtracted. E:C: E1 and
CBHl (1:4 ratio); E:C:BO.1: E1, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: El,
CBHl and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG (22.8 pg) (1:421
ratio); E:C:B2: E1, CBHl and BG (45.5 pg) (1:4:2 ratio). Data for series “Plant TSP
alone” is same as Figure 25, shown here for comparison.

79

 

Sugar released on 1% care after 18 hours
(commercial enzymes)

 

 

"‘ 0.7oo~
IletTSP+oomn.enz.

IPlaraTSPoonlrbm'on
IMTSPalone

0500 ,
3 0300 4
0.100 -
-0.1oo -

 

-0.300-

 

 

 

0.51!)

 

HURTS?

 

 

Figure 28. Sugar release on 1% CMC afier 18 h; enzyme blanks subtracted. E:C: El and
CBH] (1:4 ratio); E:C:BO.1: E1, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: E1,
CBHl and BG (11.4 pg) (1:4:0.5 ratio); E:C:Bl: E1, CBHl and BG (22.8 pg) (1:4:1
ratio); E:C:B2: E1, CBHl and BG (45.5 pg) (1:4:2 ratio). Data for series “Plant TSP
alone” is same as Figure 26, shown here for comparison.

On Avicel, very little of the sugar is attributable to the plant TSP; however, this
amount is more than the combined plant TSP alone on Avicel (same data as in Figure
25). The amount of plant TSP contribution seems to decrease with the amount of BG
added, which is opposite the results of the experiment with combined plant TSP. On
CMC (Figure 28), a comparatively larger fraction of the total sugar release is attributable
to plant TSP. These results are also for the most part inconsistent with the combined plant
TSP results (same data as Figure 26), as they are mostly negative; the only exception

again is BG2. However, the amount of plant TSP contribution seems to increase with the

80

amount of BG added, consistent with the experiment with combined plant TSP. This
could again indicate some endoglucanase activity.

The ultimate goal of producing hydrolysis enzymes in plants is to use them in
actual biomass conversion. Therefore, combinations of plant-produced E1, CBHl and
BG were applied to AF EX-treated corn stover representing 1% glucose in a 24-hour
hydrolysis reaction. The ACS had been ground to a fine powder prior to AFEX treatment
to allow it to be mixed and treated as a slurry. As before, enzyme and substrate blanks
were included and all reactions were done in triplicate. Hydrolysis was performed at

50°C, 350 RPM in a total volume of 750 pl. Sugar release was determined via DNS

 

 

 

 

 

 

 

assay.
Plant TSP comblnatlone 1% A08 24 II
1.2
1 .
0.8 -
5 0.8 « ITotd auger
Ia I E:C oonlrbutlon
g 0.4 « IBGoontrlbullon
0.2 -
0 « ' ,
.0 2 E:C:BO.1 E:C:BO.5 E:C:B1 E20232

 

 

 

Plant TSP comblnatlon

 

 

 

Figure 29. Sugar release on 1% ACS afier 24 h; blanks subtracted. E:C: El and CBHl
(1:4 ratio); E:C:BO.1: El, CBHl and BG (2.3 pg) (1:4:0.1 ratio); E:C:BO.5: E1, CBHl
and BG (11.4 pg) (12420.5 ratio); E2C2B12 E1, CBHl and BG (22.8 pg) (12421 ratio);
E2C2B22 El, CBHl and BG (45.5 pg) (1:422 ratio).

81

Sugar release from combined plant TSP after subtracting blanks is shown in
Figure 29. The results show a much higher sugar release than for either CMC or Avicel.
The best combination tested appears to be a 1:421 combination of E12CBH1:BG, with
release of nearly 1 g/L. Most of this amount appears to be due to El and CBHl, but for
this ratio it also has the highest BG contribution. In fact, for all of the ratios tested, E1
and CBHl account for most of the sugar release seen, and BG contributes relatively little
or none at all.

The same pattern is present when plant TSP is compared with commercial
enzyme (Figure 30). When Novozyme 188 is added to E12CBH1 (1 :4), its contribution to
the sugar release is less than the amount attributable to plant TSP. However, plant BG

TSP does not contribute much, if anything, to the sugar release.

82

 

Plant TSP and Commercial Enzyme 1% ACS 24 h

1.75

 

.0 re
5‘ 91

E

Sugar (0le

«0.25<

 

 

 

41.75

 

Enzyme comblnatlon

 

 

 

Figure 30. Sugar release on 1% ACS after 24 h; blanks subtracted. E:C:Novo: El and
CBHl (1:4 ratio) plus Novozyme 188; SP:B0.12 Spezyme CP (SPC) and BG (2.3 pg);

SP:BO.52 SPC and BG (11.4 pg); SP:Bl: SPC and BG (22.8 pg); SP:BZ: SPC and BG
(45.5 pg).

83

VI. DISCUSSION

1. Plant-produced E1 and BG,

Production of the hydrolysis enzymes E1 and BG has been achieved in several
plants: The catalytic domain of the thermostable El endo-l,4-B glucanase of A.
cellulolyticus (Tucker et al. 1989; Baker et al. 1994) has been successfully produced in
Arabidopsis (Ziegler et al. 2000), tobacco (Ziegelhoffer et al. 2001), rice (Oraby et al.
2007), and maize (Biswas et al. 2006; Ransom et al. 2007) and the full-length peptide in
potato (Dai et al. 2000b; Dai et a1. 2005) and tobacco (Ziegelhoffer et al. 2001). Human,
maize and B. fibrisolvens H17c B-glucosidases have been successfully expressed in
tobacco (Reggi et al. 2005; Kiran et al. 2006; Yao 2004) and the B. fibrisolvens H17c B-
glucosidase in maize (See Chapter IV. BG RESULTS).

Expression of the E1 catalytic domain yielded more activity than the full-length
enzyme (Ziegelhoffer et al. 2001); therefore it was chosen for the work in maize (Biswas
et al. 2006; Ransom et al. 2007). The expression cassette contained the strong
constitutive cauliflower mosaic virus (CaMV) 35S promoter, the tobacco mosaic virus
(TMV) Q translational enhancer, the apoplast-targeting tobacco Prla signal peptide, and
the polyadenylation signal of nopaline synthase. With the same cassette in Arabidopsis,
the plants produced El at a range of 0.01 to 25.7% of TSP (Ziegler et al. 2000). In the
work with maize (Ransom et al. 2007), the estimated El protein accumulation was up to
2.1% of TSP. Although this is several fold lower than reported in Arabidopsis (Ziegler et
al. 2000), it is in the range reported in transgenic tobacco (Dai et al. 2005) and potato

(Dai et al. 2000b). Due to the random nature of transformation, expression levels can

84

vary due to position effects. Screening a larger number of tranforrnants could yield maize
lines with higher activity.

The expression cassette for BG included the CaMV 35S promoter, an ER—leading
sequence at the 5’ end of the ngA gene, a vacuole-targeting sequence at the 3’ end, and
the CaMV 358 terminator. In tobacco, this cassette was used to study increased SAR
from B—glucosidase-induced conversion of SA 2-O-B-D-glucoside (GSA) stored in the
vacuole to free salicylic acid (SA) (Yao 2004). In the work in maize (Chapter IV), it was
useful to test the vacuole’s capacity as a storage location for heterologous hydrolysis
enzymes.

The maize-produced transgenic TSP containing E1 successfully hydrolyzed
AF FIX-treated corn stover and yielded glucose in conversion analyses afier addition of
commercial B-glucosidase. This was true even when extracted from tissue that was dry or
stored several months in the freezer (Ransom et al. 2007). TSP containing BG was also
able to hydrolyze cellobiose to glucose. Although the enzymes were biologically active,
they did not hydrolyze cell walls in planta. There are several possible explanations for
this. First, the El enzyme is thermostable and is most active at higher temperatures; its
activity assay was performed at 65°C. This is much higher than ambient temperatures or
even those experienced in summer field conditions. So, the enzyme may have had
activity that was too low to damage the cell wall before heating. Testing this hypothesis
with developing plants, however, may lead to detrimental consequences, such as heat
damage to the plant and/or plant death. A second explanation is that the enzyme may be
barred from its substrate because of its location in the ap0p1ast or vacuole and because

the cellulose itself is located inside hemicellulose and lignin. Third, while endoglucanase

85

alone could cleave random internal bonds in the crystalline cellulose, it would require
cellobiohydrolase and B-glucosidase to complete the hydrolysis. B-glucosidase alone
could hydrolyze cellobiose units, but these are likely scarce without the prior activity of
endo- and exo-glucanases. For these reasons it is safe to assume that growing individual
hydrolysis enzymes in plants, targeted to cellular compartments, and especially if they are
thermostable, poses no danger to cell wall integrity or development of the plant, as long
as transformation does not cause mutations that adversely affect cell wall integrity or
plant development, nor the transgene itself mutate to affect the same.

In the experiments for optimization of plant-produced cellulase combinations, the
highest sugar release (a little over 1.5 g/L) was observed on 1% ACS after 24 h with SPC
and BGO.5 (Figure 30), but all of the reactions except SPC2B1 achieved around the same
sugar release. The same was true for 1% CMC after 18 h; the release was nearly 1.5 g/L;
SP:B0.5 was again the highest, but all the other combinations of SP:Bx were similar and
above around 1.25 g/L (Figure 28). The next highest was on 1% ACS with E2C2Bl,
nearly 1 g/L (Figure 29). Nearly as high was SP:Bx on 1% Avicel after 18 h (Figure 27),
with amounts ranging from above 0.7 g/L (SP:BO.5) to around 0.9 g/L (SP:Bl and
SP:B2).

On 1% ACS, combinations of plant produced enzymes appear to be nearly as
effective as combinations that include SPC, although not as effective as the combination
of SPC and Novozyme 188 on 1% ACS, which routinely generates 7-10 g/L sugar (data
not shown). Plant-produced TSP on pure cellulose yielded disappointing results. It is
unclear why sugar release should be so much greater on 1% ACS rather than pure

cellulose with the same TSP.

86

Plant El and CBHl did not appear to have much activity in preliminary assays,
whereas BG appeared to have much more activity; however, in combinations, it was the
E1 and CBHl that appeared to have the most effect on sugar release rather than the BG.
One explanation is that much more 80 was used in preliminary experiments and not
much was available for further experiments. Another issue is that in preliminary
experiments using plant TSP, nearly 6.5 times the amount used in subsequent
experiments was used, yet the sugar release was much greater for 1% ACS, although
lower for pure cellulose. This is not a direct comparison, as the larger amounts were not
available-for use on 1% ACS.

In the future, the individual enzymes should be purified from the plant TSP if
possible, to compare directly with commercial enzymes and test activity directly in
comparison with purified enzymes. Although plant TSP contains a percentage of enzyme,
it is impossible to know if this enzyme maintains its total activity or if other proteins or
substances in the TSP affect its activity. Despite the drawbacks, it can be concluded that
plant TSP can be used successfully on 1% ACS to achieve conversion, and may enhance
the activity of commercial enzymes. However, the conversion is not as high as with

commercial enzymes.

2. Limitations/problems

2.1 Problems Related to Molecular Analyses

Southern blotting of E1 presented a challenge in that most enzymes were not
suitable for use in digestion because of methylation or restriction sites that were present
in the plasmid. Attempts at using Ncol or HindIII failed: Ncol did not digest maize

87

genomic DNA as thoroughly as HindIII, and HindIII digests, although nice-looking,
experienced either non-specific hybridization or no hybridization (see below). In the end,
SacI was used because it was the only enzyme that produced detectable and scorable
bands. However, since this restriction site is present on both sides of the E1 gene in the
plasmid, it was impossible to determine copy number.

During experiments on E1, results using non-radioactive DIG labeling and
detection for transgenic maize were generally poor. Probes used in Southern blots either
failed to hybridize or displayed extensive non-specific hybridization, depending on
whether random prime or PCR labeling (respectively) was used. Northern blots
consistently failed to show any hybridization although Western blots clearly showed
signal. Thus radioactive labeling and detection were used for subsequent experiments
with BG.

Non-radioactive methods had previously been used in the same laboratory for
both Southems and northems using tobacco, rice and oat. One possible explanation is that
maize has a large genome. This would require much larger amounts of nucleic acid than
recommended by the manufacturer (maximum of 5 pg). The amount used for Southems
was only 7.5 pg DNA, due to insufficient DNA available. Some have shown single-gene
copy detection in maize using DIG (Non-radioactive detection of single copy sequences
in maize; Chemiluminescent Southern detection of maize genomic single copy
sequences); in those experiments, 15-20 pg DNA were loaded. For northems, 20 pg RNA
were loaded and ethidium bromide-stained bands showed that it was present, evenly

loaded and not degraded (data not shown). The only explanation for lack of detection is

that the DIG-labeled probe is not as sensitive as 32F labeled probe for maize. Because of

88

the problems experienced here and in the experiments described in the appendices using
DIG, radioactive labeling and detection were used in subsequent BG experiments with
much better results.

For BG plants, line 8 appears to have been an escape. None of the plants tested in
PCR, Southems or northems showed evidence of presence of a gene or its transcription.
Southern blots (Figure 12) showed that lines 17 and 18 were similar to each other, and
lines 20 and 21 were similar to each other, and that both of these groups were similar to
each other, but slightly different. They all have nearly identical banding patterns;
however, lines 17 and 18 appear to have bands slightly higher than those of lines 20 and
21. However, the bands on plant 185 are slightly higher than those of 18-1 and l7-1, so
this could be due to the way the samples ran in the gel. Looking at the top of the blot, this
is indeed possible because the top of the lanes are not even across although they appeared
even on the agarose gel prior to transfer (data not shown). Since they were all
transformed on the same date, it is possible that these are not separate lines but the same
line. Or, it is possible that 17 and 18 or 20 and 21 are the same genetically. It was not
possible to determine bands on the blot of Panel B of Figure 12. The samples were too
close together and there were spotty patches, which could have been due to crystallization
of the transfer buffer dried on the blot.

Two lines had one copy (16 and 33), three had two-three copies (1, 2 and 32), two
had four-five copies (10 and 34), seven had five-six copies (3, 9, 15, 27, 28, 29 and 30),
and the rest had six-nine copies (11, 13, 17, 18, 20, 21, 22, 24, 25 and 31) (Figure 12). Of
those that showed expression in northern blots, weak signals were observed in lines 33

(one copy), 34 (four-five copies), 3, 27, 30 (five-six copies), 24, and 31 (six-nine copies);

89

and strong signals were observed in lines 1, 32 (two-three copies), 10 (four-five copies),
9, 15 (five-six copies), 11, 13, 17, 18, 20, 21 and 25 (six-nine copies). Signal was absent
from lines 16 (one copy), 28, 29 (five-six copies) and 22 (six-nine copies). Signal
strength was not correlated with copy number in these experiments.

Southern blots were performed with the aim of determining whether expression
differences were genetic or due to silencing. Where possible, samples of an expressing
and non-expressing plant from each line were tested; otherwise, two random
representatives were chosen, or in some cases, the only representative. With few
exceptions, the clones had similar banding patterns, so they were likely indeed clones,
and silencing was likely the cause of the lack of expression. Some lines had only
representatives that showed expression (3 (faint), 15, 17, 20, 21, 25, 30, 31, 32, 33); some
did not express at all (5, 6, 7, 8, 12, 26, 22, 24 28, 29). Some lines were impossible to
score due to excessive signal strength or poor blotting (5, 6, 7, 12, 27, 29, 31); all of these
did not express, except 31, which was faint, so although genetic differences could not be
detected they were not likely to play a role in expression differences. Two pairs appeared
different than their putative respective twin: 7-3 and 7-4, and 13-1 and 13-2. On the
Southern, 7-3 showed hybridization while 7-4 did not. It is possible that some escapes
exist in this line. Plant 13-1 had a different banding pattern than 13-2, which was similar

to line 15, indicating that either the line is not pure or there was a labeling mix-up.

2.2 E1 Enzyme Activity Assays

The assay used for E1 enzyme activity produced highly variable results. Also,
different plants were tested different numbers of times on different occasions. Therefore

numbers represented in tables and figures are averages and signify estimations of activity.

90

The assay was for the most part consistent with results from Western analyses.
However, in one plant, 7-6, the assay consistently placed its activity among the highest.
Yet, no band was detectable in Western blots. The only explanation for this is that the
assay is not always reliable. Perhaps this plant had high background fluorescence
interfering with the assay. Even the negative control had fluorescence that had to be
subtracted from the readings; in many cases the negative control had higher readings than
transgenic plants. In another instance, one experiment showed a plant had 9% E1 in TSP
and this was the one and only time this plant had any significant reading. Similarly,
Ziegler et a1. (2000) reported nearly 26% El accumulation in Arabidopsis; this was a
highly unusual result, even amongst the plants they studied. The inconsistencies
displayed in the highly variable results support the assertion that this assay is not reliable

and should be used with caution and in conjunction with Western analyses.

2.3 Problems Related to Materials and Recordkeeping

Another researcher responsible for transformation and care of E1 maize plants did
not keep records of individual plants so it was difficult to follow and complete his work.
The inbred line used in transformation, Hill, is generally weak and not very fertile;
however, it is the best (of very few) genotype for Type II callus production and
transformation, as maize is highly recalcitrant. In addition, growth in greenhouse
conditions is often stressful on the plants as is transformation and tissue culture.
Furthermore, these disadvantaged plants were planted in small pots so obtaining seed was
nearly impossible.

After large-scale protein extraction of several E1 plants with the highest activities,

the tubes were accidentally thrown away while being stored in a partner laboratory where

91

the glucose analysis was done. Only one plant remained, which had not been subjected to
extraction yet. Therefore, there are no replicates or comparisons available for conversion
using plant El TSP.

Second-generation analysis of El plants revealed very low activity, although the
gene appeared to have been transmitted. As the parents of these plants had, for the most
part, biologically-active El, this indicates that the gene was likely unstable in these
plants.

For experiments with BG, initial examination revealed the wrong starting
material. The lab had received a binary vector, pGLU200, from which the BG cassette
shown in Figure 2 was to be cut and put into a simple vector such as pUC19. However,
on cutting with SacI, the plasmid did not digest as it should have. It did digest with EcoRI
and give correct fragment sizes; these were cloned into pUC19 and sequenced using M13
primers. The insert had a very high match to a gene from Pseudomonas syringae; PCR
also did not result in amplification. Afier careful examination of the dissertation (Yao
2004), it was determined that during DNA manipulations, the second EcoRI site would
have been removed, so the correct plasmid should give two bands when out with Sad but
only give one band when cut with EcoRI. After alerting the people who were the source
of the material, the proper plasmids were sent. These were verified by PCR, enzyme
digestion, and sequencing.

There was one plant in the greenhouse for which the label had been lost. It
showed strong expression on northern blots (data not shown), so it was tested in the

Southems to determine which line it came from. It is indicated as plant “M” (for

92

“mystery”) on the Southern blot picture (Figure 12). However, Southern blotting failed to

elucidate its genetics because it did not seem to match any of the scorable patterns.

3. Conclusions

These experiments confirm that it is possible to grow hydrolysis enzymes in crop
plants when targeted to subcellular compartments. The enzymes retain biological activity,
are robust and storable, are able to convert pretreated feedstock biomass to fermentable
sugars, and work in synergy. This is a step forward in the quest for alternatives to current
enzyme production methods.

Much work still needs to be done. More E1 plants need to be generated and
screened for higher activity, perhaps targeting different organelles. Examination of the
next generations of the BG and new El plants will elucidate whether the transgenes are
transmitted stably and whether they retain activity. The enzymes will have to be either
stacked, i.e., the same enzyme localized to multiple compartments, or combined, i.e., a
suite of enzymes introduced into the same plant, each localized to a different organelle. It
should also be verified that the proteins are localized to the correct compartment(s).
Finally, the lines will have to be bred into more commercially acceptable cultivars. In
addition, BG plants could be tested for enhanced disease resistance or SAR response and

molecular analyses of FLC, along with delay in flowering.

93

APPENDIX A. RESULTS FOR FLC

1. Introduction

FLOWERING LOC US C (FLC) is a gene characterized in Arabidopsis that
maintains a vegetative state until it is down-regulated by vemalization, allowing
flowering to occur (Michaels and Amasino 1999; Sheldon et al. 1999). It was proposed
that it could delay flowering in other plants and this hypothesis was tested and verified in
tobacco (Salehi et al. 2005). This is an important characteristic to employ for both
increasing biomass for biofarming or feedstock for ethanol production and for transgene
containment. If plants with transgenes flower later than other plants of the same species
in the surrounding fields, it is less likely that they will cross.

It was with these goals in mind that FLC (pGreen) was used in co-transformation
with XYL] and CBHI (pMSF 15); pGreen also contains the bar gene to provide Bialaphos

herbicide resistance as a selectable marker.

2. Materials and Methods

pGreen (Figure 5) was used in co-transformation with pUC1813, XYLl and
pSMF15 as described on page 29.

For PCR, the following set of primers was used: FLC F 2 5'-CGA TAA CCT GGT
CAA GAT CC-3' (forward primer) and FLC R, 5'-CTG CTC CCA CAT GAT GAT TA-
3' (reverse primer; Salehi et al. 2005). The predicted size of the amplified DNA

fragments of the transgene was 338 bp. The PCR profile had an initial denaturation step

94

at 94°C for l min, followed by 30 cycles of 1 min at 94°C (denaturation), 2 min at 60°C
(annealing) and 3 min at 72°C (extension).

For Southern and northern analyses, a probe was generated by digesting pGreen
plasmid DNA with XhoI and SpeI to release a 0.59-kb fragment containing the FLC
coding region. Labeling and detection were done using non-radioactive methods

described on page 32.

3. Results and Discussion

Maize plants bombarded with a mixture of pGreen and XYLl, pMSF 1 5 or
pUC1813 were tested via PCR for presence of the FLC transgene (Figures 31 and 32). In
Figure 31, all of the plants tested amplified a band of the correct size with the exception
of X-l. Plants X-4 and 3 amplified a weak band. This result is not valid, however, due to
the presence of a weak amplification product in the nontransgenic negative control. In
Figure 32, all three plants amplified the correct size band; plant 2-1 however was not as
clear. The rest of the BG plants must also be tested for FLC, and analyzed using

Southems and northems as well.

W —C X-1 X-2 X-3 X4 1 2 3 P

   

'4-333 b9

 
 

Figure 31. PCR for FLC of maize plants co-transformed with either XY L1 (X-) or
pMSF15. W2 Reaction containing only water; -C2 nontransgenic negative control maize
DNA; P: pGreen plasmid DNA.

95

 

Figure 32. PCR for FLC on maize plants co-transformed with BG and F LC. W: Reaction
containing only water; L: 100 bp ladder (NEB), P: pGreen.

Southern and northern blots of maize co—transfomred with either XYLl or
pSMF 15 that had been used in hybridizations with their respective probes were stripped
and probed using the FLC probe (Figures 33 and 34). No hybridization was detected,
even with the pGreen plasmid DNA.

1235HillPF 3

           

Figure 33. Southern blots of maize co-transfonned with pSMF 15 (left side) or XYL (right
side) and pGreen. Hill: Non-transgenic negative control; P: pSMF15; F : pGreen.

96

-C(M) x-1

 

Figure 34. Northern blots of maize co-transformed with pSMF 15 (left side) or XYLl (X-;
right side) and pGreen. Ethidium-bromide stained RNA bands are shown below the blot
to show amount loaded. —C and —C(M): Non-transgenic negative control.

pGreen is a binary vector. As such, it was difficult to obtain enough DNA for
bombardment. A project was commenced to attempt to clone the FLC construct into a
simple vector. This was difficult because our plasmid map was incomplete and we did
not have the complete sequence or cutting sites. Cloning was begun but proved fi'uitless;
because the cloning site in pGreen is based on pBLUESCRlPTKS+, and this was the only
cloning vector with the appropriate restriction sites (pGreen was cut with BglIl to release
the T-DNA, and pBSKS+ cut with EcoRI and treated with Klenow for a blunt-end
ligation), it was impossible to check the sequence and very difficult to find unique sites

for cloning. The project was therefore abandoned.

97

Another project related to FLC was the attempt to obtain a polyclonal antibody
for use in western analyses. A synthetic protein sequence was generated by the Michigan
State University Research Technology Support Facility. This was used as an antigen and
sent to Cocalico Biologicals. They sent us prebleeds from five rats and a western was
performed as described on page 36. Twenty micrograms genomic DNA (Hill) and 50 ng
antigen were run in five lanes each; the blot was out after blotting and each strip
hybridized with a different prebleed diluted 1:100 in 1x PBS. Anti-ratzHRP was diluted
1210,000 leBS for the secondary antibody. There was no background detected.
Thereafter, they conjugated the antigen and injected the rats, and sent several bleeds, all
of which were tested in a similar manner and all of which showed no response to the
antigen. The antigen was then conjugated with a different carrier, thyroglobulin, and
injected as before; however the response was still the same. The project is at a standstill

as of now.

98

APPENDIX B. XYLANASE TRANSFORMATION OF MAIZE (ALONG WITH FLC),
ANALYSES OF TOBACCO AND MAIZE TRANSFORMED WITH XYL], AND

CLONING OF XYL] AND A NEW XYL

1. Introduction

The XYL] gene was cloned in Dr. Jonathan Walton’s laboratory from the fungus
Cochliobolus carbonum. Previously in the Walton lab, Apel (1996) used XYLI to
transform tobacco using two constructs (not in the same plants): one with the native
fungal signal peptide and one without. Both were not targeted to any cellular
compartment and so remained in the cytoplasm. She was not successful in finding

expression via Western blotting although she had some promising northern blots.

2. Materials and Methods

XYL] with native fungal signal peptide was sent to a company, Norclone, to put
into the apoplast-targeting vector pMZ766 in place of the E1 gene (Figure 35). This
plasmid was used for tobacco transformation (performed by another researcher) and

particle bombardment, along with pGreen (Figure 5), of maize.

99

SphI BamHI
HindIII Sbfl

Sall SacI Pstl Sail SacI Xbal

| l l l I
«L CaMV 358 9 Pg}? XYL lnosl WW9

Xbal NCOI ECORV

 

 

 

 

 

 

 

 

 

 

 

0.8 kb 0.2 kb 0.09 kb 0.7 kb 0.3 kb
J

V

2.09 kb

Figure 35. XYL Construct. CaMV 35S: Cauliflower mosaic virus 35S promoter; S22
tobacco mosaic virus translational enhancer; Prla: signal peptide from tobacco
pathogenesis-related protein la; XYLI 2 xylanase gene from C. carbonum; nos: nos
terrnrnator.

Maize callus production, bombardment, selection and regeneration were carried
out as described previously in Materials and Methods (page 29), as were DNA, RNA and
protein extraction (pages 30, 32 and 33) and Southern, northern and western blotting
(pages 32 and 36). For Southern blots, 15 pg genomic DNA was digested overnight with
HindIII and fractionated on a 1% agarose gel. Tobacco that had been transformed with
Agrobacterium by another researcher were included as well. PCR and RT-PCR were
performed using the following primers and conditions: Xyll-F: 5’ CTG CCC GTA CCA
TCA CCT AC 3’ and Xyll-R: S’GTG ATC TGG GCG TTA CCA GT 3’ (397 bp); 94°C
for 3 min; 35 cycles of 94°C for 45 s, 56°C for 45 s, 72°C for 45 s; 72°C for 10 min. The
probe used for Southern and northern hybridization and detection was the 397-hp
fragment generated by PCR described above.

A polyclonal antibody to XYLl was obtained from Dr. Walton’s laboratory and

used in western blots. For activity assays, the assay for reducing sugars using PABAH

(Lever 1972) was used. To a 1% oat spelt xylan substrate, 25 pl protein samples were

100

added to 0.5M acetate buffer in a total volume of 300 pl, and incubated at 37°C for 0 min,
30 min, 4 h and overnight. Samples (25 pl) were collected at the designated time points,
1.5 ml PABA solution was added, the samples were mixed, heated at 100°C for 10 min

and cooled before reading absorbance at 410 nm.

3. Results and Discussion

3.1 Transformation of maize with XYL] and F LC

More than 180 plates of maize callus were bombarded and five resistant clones
were obtained. Of these, 10 plants were regenerated but only four survived to maturity.

These plants were weak and proved to be infertile (Figure 36).

101

 

t «x

Figure 36. Maize plants transformed with XYL] and FLC.

3.2 Molecular and enzyme assays for XYL on maize and tobacco

Molecular analyses on maize and tobacco plants were performed. PCR (Figure
37) revealed that one of the maize plants and two of the tobacco plants were likely to

contain the XYL] gene sequence. All lanes, including the water, negative (untransformed

102

control) and positive control (XYLI plasmid) had primer dimers (appearing as a band of

smaller molecular weight), indicating that primer concentration was likely too high.

 

Figure 37. PCR for X11 in maize and tobacco plants. Numbers followed by M indicate
maize plants; numbers followed by T indicate tobacco plants. W2 Reaction containing
only water; -C: Untransformed maize plant; P2 XYL] plasmid.

DNA from maize and tobacco plants did not show any hybridization with the
XYL] probe in a Southern blot (Figure 38) although plants 3M, 12T and 15T showed an
amplified band after PCR. Because the method used for hybridization and detection was
non-radioactive (DIG), it may not have been sensitive enough to detect a low copy
number in transgenic plants. Similarly, northern blots for maize and tobacco plants did
not show any hybridization (Figure 38). RT-PCR (Figure 40) showed two plants had a
band (1 IT and 12T); however, the band for 1 IT was also present in the no-RT control,
indicating that the RNA was contaminated with DNA, and this is not a true amplification
from RNA. It was possible the band seen in 12T was legitimate, although it is very faint.
All samples had large primer dimers, indicating lack of template or too much primer in
the reaction. This means that it is unlikely that any of the plants were producing XYL

RNA.

103

1M 2M 3M _ P 4T 7T 11T12T 15T 20T -C(T)

  
   

A...

Figure 38. Southern blot of maize and tobacco plants transformed with XYL] and FLC.
Numbers followed by M indicated maize plants; numbers followed by T indicate tobacco
plants. P: XYL] plasmid; -C: non-transformed maize (M) or tobacco (T).

104

4M 7T 11T12T 14T 15T 20T Cu

     

          

-CM 1M 2M 3M

.4.— ‘Wi‘fit’nmnm mug

-CM M 2M 3M 4N|>TTA11T12T 14T15T 20T-C

 

Figure 39. Northern blot of maize and tobacco plants transformed with XYL] and FLC.
Numbers followed by M indicated maize plants; numbers followed by T indicate tobacco
plants. The probe was a 397 bp PCR-generated fragment of the XYL] gene. Upper panel:
blot probed with PCR-labeled DIG probe; bottom panel: same blot probed with random
primed DIG labeled probe. —CM: non—transformed maize; —CT non-transformed tobacco.
Ethidium bromide-stained bands from the agarose gel prior to transfer are shown below

the blot to show relative amounts of RNA loaded.

105

1M 2M 3M 4M 7T 11T12T 14T 15T 20T -CT M

m. ' RT Reaction

No-RT Control

 

Figure 40. RT-PCR of maize and tobacco plants transformed with XYL] and FLC.
Numbers followed by M indicated maize plants; numbers followed by T indicate tobacco
plants. Top panel: RT reaction; bottom panel: No-RT control. —CM: non-transformed
maize; -CT non-transformed tobacco; M: 100 bp molecular weight marker (NEB).

A Western blot was performed on eight of 10 maize plants (two being left out
because of lack of sufficient protein) (Figure 41). Tissue had been collected prior to plant
death. In two of the plants, 2 and 3, a single band with a molecular weight of around 40
kDa was detected and in plant 6, a band of around 20 kDa was detected. All of the
transgenic plants had a large, 100-kDa protein that was absent in the non-transformed

control.

106

CD1ME2M4M-C kDa
- so
— so
-- 50
— 40

 

-20

Figure 41. Western blot of maize plants transformed with XYL] and FLC.
Letters/numbers above the lanes represent individual plants; 1M, 2M and 4M correspond
with the plants that survived to maturity and are represented in the other figures. —C2
Untransformed maize negative control. Size markings are indicated on the right side.

The assay for reducing sugars using PABAH (Lever 1972) was used on the plants.
Absorbance readings at 410 nm for the plants after subtracting the 0 time point is shown
in for maize in Table 9 and tobacco (both total protein concentrate and extracellular fluid
wash; Herbers et al. 1995) in Table 10. An increase of 0.2 to 1 shows activity. None of

the plants showed any significant activity that was greater than negative control.

107

Table 9. Maize plants tested for XYL activity using PABAH assay for reducing sugars.
Absorbance taken at 410 nm for 0 min, 30 min and 4 h are shown, along with the

adjusted values (0 min subtracted).

Plant 0min 30 min 30 min

A 1 0.9634
B 1.4841 1.0378
0.198
0.5352 0.4622
0.3742 0.3270
0.2018 0.2131

0.7939 0.6459

0.2766 0.2637
0.3355

0.5277 0.

4
0.9630
1.0775

0.4419
0.3384
0.2642

 

4h

Table 10. Tobacco plants total protein concentrate and extracellular fluid wash tested for
XYL activity using PABAH assay for reducing sugars. Absorbance taken at 410 nm for 0
min, 30 min and overnight are shown, along with the adjusted values (0 min subtracted).

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

   

 

 

 

 

 

 

 

 

 

 

 

Plant I 0 min] 30 min I 30 min adjj Over night I O/n adj.
Total protein concentrate

-C 0.481 0.4839 0.0029 0.5454 0.0644
4 0.3341 0.3968 0.0627 0.4799 0.1458
7 0.2374 0.3268 0.0894 0.2954 0.058
11 0.2729 0.326 0.0531 0.3191 0.0462
12 0.3426 0.3818 0.0392 0.4336 0.091
14 0.3875 0.4161 0.0286 0.5045 0.117
15 0.2866 0.3193 0.0327 0.3801 0.0935
20 0.3204 0.3247 0.0043 0.3954 0.075
Extracellular fluid wash

-C 0.213 0.2597 0.0467 0.2139 0.0009
4 0.2843 0.2957 0.0114 0.2497 ' " ‘
7 0.2465 0.2875 0.041 0.218

11 0.2317 0.2587 0.027 0.1709

12 0.2619 0.291 0.0291 0.2394

14 0.254 0.2746 0.0206 0.2114

15 0.2625 0.2903 0.0278 0.2069

20 0.2603 0.2841 0.0238 0.2224

 

 

108

After all the analyses were performed, it was discovered that the company that
had made the construct, Norclone, had used available restriction sites in the original
plasmid for cloning the DNA fragment into the vector. Unfortunately, these sites cut a
fragment that contained part of the plasmid backbone (i.e., “junk”). The gene remained in
frame relative to the rest of the construct and no stop codons were present. In addition,
this DNA sequence contained the native fungal secretory signal peptide.

Several attempts were made to correct the construct by removing both the junk
and the fungal signal peptide, but failed due to mutations or incorrect orientation.
Because neither construct (with and without the fungal signal peptide) had worked
previously in tobacco (Apel, 1996), it was unlikely that it could work in maize even with
a different signal peptide (i.e., apoplast). As a result, this project was shelved, and a new

project with a new xylanase (XYL from A. cellulolyticus, gift from Edenspace Corp.) was

begun.

3.3 Cloning of new XYL

3.3.1 pMZ766 vector

Primers were ordered that added SacI sticky ends to the XYL gene, which were
then used in PCR and the gene was amplified. As before, the PCR fragment was cloned
into a T-vector, sequenced, and then cloned into the pMZ766 backbone. Several attempts
were made but all of the resulting clones had the gene in the reverse orientation. So this
project was given to another researcher. Several tobacco plants resistant to kanamycin
were generated but PCR revealed that none of them out of the l 1 that were tested had the

XYL gene.

109

3.3.2 ImpactVectors

An additional set of primers was ordered that added Neal and BglII sticky ends to
the XYL gene. This fragment was to be cloned into the five ImpactVectors. These vectors
are specifically for plant transformation and employ the rubisco small subunit (Rch l)
promoter from the Asteraceous Chrysanthemum and 1 kb of the RchI terminator
sequence. Each one utilizes a different targeting sequence: cytoplasm, secretory pathway,
endoplasmic reticulum, chloroplast and mitochondria. In addition, the vectors have a
cmyc-tag allowing identification of expressed proteins using commercially available
monoclonal antibodies and a six histidine His-tag for protein purification using a nickel
column.

After cloning was accomplished and verified with sequencing, each of the five
targeting constructs was cloned into the binary vector provided with the ImpactVectors,
pBINPLUS. As before, the sequences were verified after cloning procedures. In addition,
the vector was used to transform A grobacterium. Afier the cloning was accomplished, the
plasmid DNA in the simple vector was prepared for bombardment by maxiprep. Maize

callus transformation using the ImpactVectors with XYL is currently underway.

110

APPENDIX C. CBHI AND FLC IN MAIZE

1. Introduction

Cellobiohydrolase I from T. reesei is an exoglucanase previously shown to have
activity when expressed in tobacco (Dai et al. 1999). In this set of experiments, it was

attempted to do the same in maize.

2. Materials and Methods

A construct containing a synthetic CBHI that had been codon modified for use in
tobacco transformation was made previously in the Sticklen laboratory. It had the rice
Rubisco (rch) small subunit promoter, the rch chloroplast signal peptide also from rice,

and the Agrobacterium nopaline synthase (nos) 3’ non-coding region (Figure 42).

 

 

 

 

pSMF15
-l M HEH W“ I!”
1.110» 0.14“) 1.510:

Figure 42. pSMF15. rch: rice Rubisco small subunit promoter; TP: rice rch chloroplast
signal peptide; Syn-cth 2 synthetic CBHI; nos: nopaline synthase 3’ non-coding region.

Maize callus production, bombardment, selection and regeneration were carried
out as described previously in Materials and Methods (page 29), as were DNA, RNA and
protein extraction (pages 30, 32 and 33) and Southern, northern and western blotting

(pages 32 and 36). For Southern blots, 15 pg genomic DNA was digested overnight with

111

HindIII and fractionated on a 1% agarose gel. PCR and RT-PCR were performed using
the following primers and conditions: Syn-cbhl-F: 5’ TCT TGA TGG TGC TGC TTA
CG 3’ and Syn-cbhl-R: 5’ CCA AAC TCA GCT TCC TCA GC 3’ (801 bp); 94°C for 3
min; 35 cycles of 94°C for 45 s, 56°C for 45 s, 72°C for 45 s; 72°C for 10 min. The probe
used for Southern and northern hybridization and detection was the 801-bp fragment
generated by PCR described above. For sequencing, an additional primer was ordered,

Syn-cbhl—R—Seq: 5’ CGT AAG CAG CAC CAT CAA GA 3’.

3. Results and Discussion

3.] Transformation of maize with Syn-CBHI and FLC

Seventy-five plates of maize callus were bombarded with a combination of
pMSF l 5 (containing Syn-CBHI) and pGreen (containing FLC) and 15 resistant clones

were obtained. From these, 12 plants were regenerated but only five survived to maturity.

3.2 Molecular analyses for Syn-CBHI on maize

Molecular analyses on maize plants were performed. DNA from maize plants
showed a very faint hybridization with the Syn-CBHI probe in a Southern blot that was
not present in non-transgenic control, all around the same size as the plasmid band

(Figure 43).

112

 

Figure 43. Southern blot of maize plants transformed with Syn-CBHI and FLC. P2
pMSF 15 plasmid; -C2 non-transformed maize.

However, northern blots did not show any hybridization and RT-PCR showed that
none of the plants had a band. All samples had large primer dimers, indicating lack of
template or too much primer in the reaction. This means that it is unlikely that any of the

plants were producing Syn—CBHI RNA.

113

12345-C12345—C

ram—”E‘— "mu-ama-

 

Figure 44. Northern blot of maize plants transformed with Syn-CBHI and FLC. The
probe was a 801 bp PCR-generated fragment of the Syn-CBHI gene. Left panel: blot
probed random primed DIG labeled probe; right panel: same blot probed with PCR-
labeled DIG probe. -C: non-transformed maize. Ethidium bromide-stained bands from the
agarose gel prior to transfer are shown below the blot to show relative amounts of RNA
loaded.

 

Figure 45. RT-PCR of maize plants transformed with Syn-CBHI and FLC. Top panel: RT
reaction; bottom panel: No-RT control. -C: non-transformed maize; M: 100 bp molecular
weight marker (NEB). Arrow indicates 801 bp.

114

After all the analyses were performed, when it was discovered that some of our
constructs had mistakes, we decided to check all the constructs, so pMSF15 was
sequenced. Sequencing revealed many frameshifts caused by deletions, a large fragment
of the CaMV35S promoter, numerous stop codons, and a deletion in the start codon. It
was discovered that major mistakes had been made during the cloning.

To fix the construct, new primers were ordered to amplify the rch transit peptide
with XbaI sticky ends and insert it between the 35S promoter and Syn-CBHI gene in the

original Syn-CBHI plasmid, pZD408 (Figure 46).

FWMflflGB?
PSfl710

  

March 15. 1999

  
  
  

Xbal1512

Ewan1931

EooRl 3350 3an 2111
Sacl 2520
- EooRl
3050
.. (SaclPatl)

 

Figure 46. pZD408. 35S: Cauliflower Mosaic Virus (CaMV) 35S Promoter; SynCBHI:
Synthetic CBHI coding region.

115

This was done, and the sequence checked in the T-vector. Both pZD408 and the T-
vector were digested with XbaI and the correct fragments ligated together. PCR was
performed to verify orientation. The construct was then sequenced. A single point
deletion in the middle of the Syn-CBHI gene was discovered; this was seen in more than
one clone. So, pZD408 was sent to sequencing, and it was found that the mutation was
present in the original vector’s gene sequence. Another plasmid that had been used in the
construction of pZD408 in Dr. Dai’s laboratory contained the correct sequence
(pZD394).

However, this plasmid did not have any promoter or terminator sequences, just
the Syn-CBHI gene in a lacZ multiple cloning site. Originally, the plan was to cut out the
correct sequence and excise the incorrect sequence and re-ligate them, but there proved to
be no available restriction sites. Correcting the construct then became a complicated
matter of amplifying the various pieces adding sticky ends and ligating them together
sequentially in a simple vector. Eventually, the project was abandoned due to its

complexity and lack of a suitable vector.

116

REFERENCES

Andersson-Gunneras, S., E]. Mellerowicz, J. Love, B. Segerrnan, Y. Ohmiya, P.M.
Coutinho, P. Nilsson, B. Henrssat, T. Moritz and B. Sundberg. 2006. Biosynthesis
of cellulose-enriched tension wood in Populus: global analysis of transcripts and
metabolites identifies biochemical and developmental regulators in secondary
wall biosynthesis. The PlantJ. 45(2): 144-165.

Apel, RC. 1996. The Cloning, Comparison, and Expression of Three Family G Endo B-
1,4-xylanase Genes of the Maize Fungal Pathogen Cochliobolus Carbonum and
Analysis of Their Importance For Pathogenicity on Maize. Michigan State
University.

Arioli, T., L. Peng, A.S. Betzner, J. Burn, W. Wittke, W. Herth, C. Camilleri, H. Héfte, J.
Plazinski, R. Birch, A. Cork, J. Glover, J. Redmond, R. E. Williamson. 1998.
Molecular Analysis of Cellulose Biosynthesis in Arabidopsis. Science 279(5351):
717-720.

Armstrong, C.L., and CE. Green. 1985. Establishment and maintenance of friable,
embryogenic maize callus and the involvement of L-proline. Planta 1642207-214.

Armstrong, C.L., C.E. Green, and R.L. Phillips. 1991. Development and availability of
germplasm with high Type 11 culture formation response. Maize Genet. Coop.
Newslett 65:92-93.

Baker, J.O., W.S. Adney, R.A. Nieves, S.R Thomas, D.B. Wilson, and ME. Himmel.
1994. A new thermostable endoglucanase, Acidothermus cellulolyticus E1.
Synergism with T richoderma reesei CBH I and comparison to Thermomonospora
fizsca E5. Appl. Biochem. Biotech. 45/462245-256.

Baucher, M., M.A. Bemard-vailhe', B. Chabbert, J.M. Besle, C. Opsomer, M. Van
Montagu, and J. Botterman. 1999. Down-regulation of cinnamyl alcohol
dehydrogenase in transgenic alfalfa (Medicago sativa L.) and the effect on lignin
composition and digestibility. Plant Mol. Biol. 39(3): 437-447.

Biomass Program: Net Energy Balance for Bioethanol Production and Use.
http://wwwl .eere.energy.gov/biomass/net_energy_balance.htrnl (accessed August
1,2007)

Biswas, G.C.G., C. Ransom, and M. Sticklen. 2006. Expression of biologically active

Acidothermus cellulolyticus endoglucanase in transgenic maize plants. Plant
Science 171(5): 617-623.

117

Blaschke, L., M. Legrand, C. Mai, and A. Polle. 2004. Lignification and structural
biomass production in tobacco with suppressed caffeic/S-hydroxy ferulic acid-0-

methyl transferase activity under ambient and elevated C02 concentrations.
Physiol. Plant. 121(1): 75-83.

Bohmert, K., I. Balbo, A. Steinbuchel, G. Tischendorf, and L. Willmitzer. 2002.
Constitutive Expression of the B-Ketothiolase Gene in Transgenic Plants. A
Major Obstacle for Obtaining Polyhydroxybutyrate-Producing Plants. Plant
Physiol. 128(4): 1282-1290.

Boisset, C., C. Pétrequin, H. Chanzy, B. Henrissat, and M. Schfilein. 2001. Optimized
mixtures of recombinant Humicola insolens cellulases for the biodegradation of
crystalline cellulose. Biotechnology and bioengineering 72(3): 339-45.

Bolwell, GP. 2000. Biosynthesis of Plant Cell Wall Polysaccharides. Trends Glycosci.
Glycotech. 12(65): 143-160.

Boominathan, K., and CA. Reddy. 1992. Fungal degradation of lignin: biotechnological
applications. In Fungal Biotechnology, 763-882.

Bordetsky, A., R. Hwang, A. Korin, D. Lovaas, and L. Tonachel. 2005. SECURING
AMERICA: Solving Our Oil Dependence Through Innovation. Natural Resources
Defense Council.

Bothast, R.J., and M.A. Schlicher. 2005. Biotechnological processes for conversion of
corn into ethanol. Appl. Microbial. Biotech. 67(1): 19-25.

Boudet, A.M., S. Kajita, J. Grima-Pettenati, and D. Goffner. 2003. Lignins and
lignocellulosics: a better control of synthesis for new and improved uses. Trends
Plant Sci. 8(12): 576-581.

Boudet, Alain-M. 2000. Lignins and lignification: Selected issues. Plant Physiol.
Biochem. 3 8(1-2): 81-96.

Bradford, M. 1976. A rapid and sensitive method for quantitation of microgram
quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem.
72:248-254.

Bruins, M.E., A.E. M. Janssen, and RM. Boom. 2001. Therrnozymes and their
applications-A review of recent literature and patents. Appl. Biochem. Biotech.
902155-186.

Cai, Y.J., S.J. Chapman, J.A. Buswell, and ST. Chang. 1999. Production and
Distribution of Endoglucanase, Cellobiohydrolase, and beta -Glucosidase
Components of the Cellulolytic System of Volvariella volvacea, the Edible Straw
Mushroom. Appl. Environ. Microbiol. 65(2): 553-559.

118

Cao, J., X. Duan, D. McElroy, and R. Wu. 1992. Regeneration of herbicide resistant
transgenic rice plants following microprojectile-mediated transformation of
suspension culture cells. Plant Cell Reports 11:586-591.

Carpita, NC, and D.M. Gibeaut. 1993. Structural models of primary cell walls in
flowering plants: consistency of molecular structure with the physical properties
of the walls during growth. The PlantJ. 3(1): 1-30.

Carpita, N. and M. McCann. 2002. Chapter 2. The cell wall. In Biochemistry &
Molecular Biology of Plants, ed. B. Buchanan, W. Gruissem, and R.L. Jones, 52-
108. John Wiley & Sons.

Carrao-Panizzi, M.C., and J .R. Bordingnon. 2000. Activity of beta-glucosidase and levels
of isoflavone glucosides in soybean cultivars affected by the environment
Pesquisa Agropecua'ria Brasileira 352873-878.

Chabannes, M., A. Barakate, C. Lapierre, J .M. Marita, J. Ralph, M. Pean, S. Danoun, C.
Halpin, J. Grima-Pettenati and A.M. Boudet. 2001. Strong decrease in lignin
content without significant alteration of plant development is induced by
simultaneous down-regulation of cinnamoyl CoA reductase (CCR) and cinnamyl
alcohol dehydrogenase (CAD) in tobacco plants. The PlantJ. 28(3): 257-270.

Chiang, C.M., F .S. Yeh, L.F. Huang, T.H. Tseng, M.C. Chung, C.S. Wang, H.S. Lur, J.F.
Shaw, and SM. Yu 2005. Expression of a bi-functional and thermostable
amylopullulanase in transgenic rice seeds leads to autohydrolysis and altered
composition of starch. M01. Breeding 15:125-143.

Chu, CC, CC. Wang, C.S. Sun, C. Hus, K.C. Yin, C.Y. Chu, et al. 1975. Establishment
of an efficient medium for anther culture of rice through comparative experiments
on the nitrogen source. Scientia Sinica 182659-668.

Conrad, U. and U. F iedler. 1998. Compartment-specific accumulation of recombinant
immunoglobulins in plant cells: an essential tool for antibody production and
immunomodulation of physiological functions and pathogen activity. Plant Mol.
Biol. 38(1): 101-109.

D'Souza, T.M., C.S. Merritt, and CA. Reddy. 1999. Lignin-Modifying Enzymes of the
White Rot Basidiomycete Ganoderma lucidum. Appl. Environ. Microbiol. 65(12):
5307-5313.

Dai, 2., BS. Hooker, R.D. Quesenberry, and J. Gao. 1999. Expression of Trichoderma

reesei exo-cellobiohydrolase I in transgenic tobacco leaves and calli. Appl.
Biochem. Biotech. 77-792689-99.

119

Dai, 2., BS. Hooker, D.B. Anderson, and SR. Thomas. 2000a. Expression of
Acidothermus cellulolyticus endoglucanase E1 in transgenic tobacco: biochemical
characteristics and physiological effects. Trans. Res. 9, no. 1 (February 1): 43-54.

 

. 2000b. Improved plant-based production of El endoglucanase using potato:
expression optimization and tissue targeting. Mol. Breeding 62277-285.

Dai, 2., BS. Hooker, R.D. Quesenberry, and SR. Thomas. 2005. Optimization of
Acidothermus cellulolyticus Endoglucanase (E1) Production in Transgenic
Tobacco Plants by Transcriptional, Post-transcription and Post-translational
Modification. Trans. Res. 14(5): 627-643.

Dean, J.F.D. 2004. Synthesis of Lignin in Transgenic and Mutant Plants. In
Biotechnology of Biopolymers: From Synthesis to Patents, ed. A. Steinbuchel and
Y. Doi, 4-21. Wiley-VCH.

Decker, S.R., W.S. Adney, E. Jennings, T.B. Vinzant, and ME. Himmel. 2003.
Automated Filter Paper Assay for Determination of Cellulase Activity. Appl.
Biochem. Biotechnol. 1072689-704.

Demain, A.L., M. Newcomb, and J.H.D. Wu. 2005. Cellulase, Clostridia, and Ethanol.
Microbiol. Mol. Biol. Rev. 69(1): 124-154.

Ding, S.Y., and ME. Himmel. 2006. The Maize Primary Cell Wall Microfibril: A New
Model Derived from Direct Visualization. J. Agric. Food Chem. 54(3): 597-606.

Dodd, A.N., N. Salathia, A. Hall, E. Kevei, R Toth, F. Nagy, J.M. Hibberd, A.J. Millar,
and A.A.R. Webb. 2005. Plant Circadian Clocks Increase Photosynthesis,
Growth, Survival, and Competitive Advantage. Science 309(5734): 630-633.

Eggeman, T., and RT. Elander. 2005. Process and economic analysis of pretreatment
technologies. Biores. Tech. 96(18): 2019-2025.

Eriksson, M.E., M. Israelsson, O. Olsson, and T. Moritz. 2000. Increased gibberellin
biosynthesis in transgenic trees promotes growth, biomass production and xylem
fiber length. Nat. Biotech. 18, no. 7 (July): 784-788.

Fischer, R., E. Stoger, S. Schillberg, P. Christou, and RM. Twyman. 2004. Plant-based
production of biopharmaceuticals. Curr. Opin. Plant Biol. 7(2): 152-158.

Genencor Celebrates Major Progress in the Conversion of Biomass to Ethanol - Genencor
a Danisco division. http2//www.genencor.com/cms/connect/genencor/media
_relations/news/archive/2004/gen_2l1004_en.htm (accessed August 3, 2007).

Ghose, T. K. 1987. Measurement of cellulase activities. Pure Appl. Chem 59(2): 257-268.

120

Greene, N., F.E. Celik, B. Dale, M. Jackson, K. Jayawardhana, H. Jin, E.D. Larson, M.
Laser, L. Lynd, D. MacKenzie, J. Mark, J. McBride, S. McLaughlin and D.
Saccardi. 2004. Growing Energy: How Biofuels Can Help End America's Oil
Dependence.

Haigler, CH. 2006. Establishing the cellular and biophysical context of cellulose
synthesis. In, The Science and Lore of the Plant Cell Wall: Biosynthesis, Structure
and Function. ed. T. Hayashi, 368. Brown Walker Press (FL).

Hatakka, A. 1994. Lignin-modifying enzymes fi'om selected white-rot fungi: production
and role in lignin degradation. FEMS Microbiol. Rev. 13:125-135.

Hellens, R., P. Mullineaux, and H. Klee. 2000. Technical Focus: A guide to
Agrobacterium binary Ti vectors. Trends Plant Sci. 5(10): 446-451.

Herbers, K., I. Wilke, and U. Sonnewald. 1995. A Thermostable Xylanase from
Clostridium thermocellum Expressed at High Levels in the Apoplast of
Transgenic Tobacco Has No Detrimental Effects and Is Easily Purified. Nat.
Biotech. 13(1): 63-66.

Hong, C.Y., K.J. Cheng, T.H. Tseng, C.S. Wang, L.F. Liu, and SM. Yu. 2004.
Production of two Highly Active Bacterial Phytases with Broad pH Optima in
Germinated Transgenic Rice Seeds. Trans. Res. 13(1): 29-39.

Horn, M.E., S.L. Woodard, and J.A. Howard. 2004. Plant molecular farming: systems
and products. Plant Cell Rep. 22(10): 711-720.

Hoshino, E., M. Shiroishi, Y. Amano, and T. Kanda. 1997. Synergistic Actions of Exo-
Type Cellulases in the Hydrolysis of Cellulose with Different Crystallinities. J.
Fermentation Bioeng. 84(4): 300-306.

Houghton, J., J. Ferrel, and S. Weatherwax. 2005. From Biomass to Biofuels: A
Roadmap to the Energy Future. Proceedings of the Biomass to Biofitels
Workshop, December 7.

Howard, J.A., and E. Hood. 2005. Bioindustrial and Biopharmaceutical Products
Produced in Plants. Adv. Agron. (85)91-124, Elsevier Inc.

Howard, R.L., E. Abotsi, E.L. Jansen van Rensburg, and S. Howard. 2003.
Lignocellulose biotechnology: issues of bioconversion and enzyme production.
Afi'. J. Biotech. 2(12): 602-619.

Hu, W.J., S.A. Harding, J. Lung, J.L. Popko, J. Ralph, D.D. Stokke, C.J. Tsai, and V.L.

Chiang. 1999. Repression of lignin biosynthesis promotes cellulose accumulation
and growth in transgenic trees. Nat. Biotech. 17(8): 808-812.

121

Hyunjong, B., D.S. Lee, and I. Hwang. 2006. Dual targeting of xylanase to chloroplasts
and peroxisomes as a means to increase protein accumulation in plant cells. J.
Exp. Bot. 57(1): 161-169.

Ingledew, W.M. 1995. The Biochemistry of Ethanol Production. In The Alcohol
Textbook, ed. T.P. Lyons, D. Kelsall, and J. Murtagh, 55-79. Nottingham, UK:
Nottingham University Press.

Kabel, M.A., M.J.E.C. van der Maarel, G. Klip, A.G.J. Voragen, and HA. Schols. 2006.
Standard assays do not predict the efficiency of commercial cellulase preparations
towards plant materials. Biotech. Bioeng. 93(1): 56-63.

Kawagoe, Y., and DP. Delmer. 1997. Pathways and genes involved in cellulose
biosynthesis. Gen. Eng. 19:63-87.

Kim, S., and BE. Dale. 2004. Global potential bioethanol production from wasted crops
and crop residues. Biomass Bioen. 26(4): 361-375.

Kimura, T., T. Mizutani, K. Sakka, and K. Ohmiya. 2003a. Stable expression of a
thermostable xylanase of Clostridium thermocellum in cultured tobacco cells. J.
Biosci. Bioeng. 95(4): 397-400.

Kimura T., T. Mizutani, T. Tanaka, T. Koyama, K. Sakka, and K. Ohmiya. 2003b.
Molecular breeding of transgenic rice expressing a xylanase domain of the xynA
gene from Clostridium thermocellum. Appl. Microbial. Biotechnol. 62(4): 374-
379.

Kiran, N.S, L. Polanska, R. Fohlerova, P. Mazura, M. Vélkova, M. Smeral, J. Zouhar, J.
Malbeck, P.I. Dobrev, I. Machackova, and B. Brzobohaty. 2006. Ectopic over-
expression of the maize beta-glucosidase Zm-p60.l perturbs cytokinin
homeostasis in transgenic tobacco. J. Exp. Bot. 57(4): 985-96.

Kirk, T.K., and R.L. Farrell. 1987. Enzymatic "combustion": the microbial degradation of
lignin. Annu. Rev. Microbiol. 412465-505.

Knauf, M., and M. Moniruzzaman. 2004. Lignocellulosic biomass processing: A
perspective. Int. Sugar J. 1062 147-150.

Lever, M. 1972. A new reaction for colorimetric determination of carbohydrates. Anal.
Biochem. 47(1): 273-279.

Li, L., Y. Zhou, X. Cheng, J. Sun, J.M. Marita, J. Ralph, and V.L. Chiang. 2003.

Combinatorial modification of multiple lignin traits in trees through multigene
cotransformation. Prac. Nat. Acad. Sci. USA. 100(8): 4939-4944.

122

Liu, H.L., W.S. Li, T. Lei, J. Zheng, Z. Zhang, X.F. Yan, 22. Wang, Y.L. Wang and
LS. Si. 2005. Expression of Human Papillomavirus type 16 L1 protein in
transgenic tobacco plants. Act. Biachim. Biaphys. Sin. 372153-158.

Luo, Y., J.L. Chen, J.F. Reynolds, C.B. Field, and HA. Mooney. 1997. Disproportional
increases in photosynthesis and plant biomass in a Californian grassland exposed

to elevated C022 a simulation analysis. F unct. Ecol. 1 1(6): 696-704.

Lynd, L.R., W.H. van Zyl, J.E. McBride, and M. Laser. 2005. Consolidated
bioprocessing of cellulosic biomass: an update. Curr. Opin. Biotech.. 16:577-583.

Michaels, S., and R. Amasino. 1999. FLOWERING LOCUS C Encodes a Novel MADS
Domain Protein That Acts as a Repressor of Flowering. The Plant Cell 11:949-
956.

Miller, G.L. 1959. Use of Dinitrosalicylic Acid Reagent for Determination of Reducing
Sugar. Anal. Chem. 31(3): 426-428.

Mosier, N., C. Wyman, B. Dale, R. Elander, Y.Y. Lee, M. Holtzapple, and M. Ladisch.
2005. Features of promising technologies for pretreatment of lignocellulosic
biomass. Biores. Tech. 96(6): 673-686.

Murray C., P.W. Sutherland, M.M. Phung, M.T. Lester, R.K. Marshall and T. Christeller.
2002. Expression of Biotin-Binding Proteins, Avidin and Streptavidin, in Plant
Tissues Using Plant Vacuolar Targeting Sequences. Trans. Res. 11(2): 199-214.

Nuttall, J., N. Vine, J.L. Hadlington, P. Drake, L. Frigerio, and J.K.C. Ma. 2002. ER-
resident chaperone interactions with recombinant antibodies in transgenic plants.
Eur. J. Biochem. 269(24): 6042-6051.

O'Neill, M.A., T. Ishii, P. Albersheim, and AG. Darvill. 2004. Rhamnogalacturonan 11:
structure and function of a borate cross-linked cell wall pectic polysaccharide.
Annu. Rev. Plant Biol. 552109-39.

Oraby, H., B. Venkatesh, B. Dale, R Ahmad, C. Ransom, J. Oehmke, and M. Sticklen.
2007. Enhanced conversion of plant biomass into glucose using transgenic rice-
produced endoglucanase for cellulosic ethanol. Trans. Res. Published Online
First: 20 January 2007.

Pan, X., N. Gilkes, J. Kalda, K. Pye, S. Saka, D. Gregg, K. Ehara, D. Xie, D. Lam and J.
Saddler. 2005. Biorefining of sofiwoods using ethanol organosolv pulping:
Preliminary evaluation of process streams for manufacture of fuel-grade ethanol
and co-products. Biotech. Bioeng. 90(4): 473-481.

123

Patel M., J .S. Johnson, R.I.S. Brettell, J. Jacobsen, and GP. Xue. 2000. Transgenic barley
expressing a fungal xylanase gene in the endosperm of the developing grains.
Mal. Breed. 6(1): 113-124.

Perlack, R.D., L.L. Wright, A.F. Turhollow, R.L. Graham, B.J. Stokes, and DC. Erbach.
2005. Biomass as a Feedstock for a Bioenergy and Biapraducts Industry: The
Technical Feasibility of a Billion-Tan Annual Supply. U.S. Dept. of Energy and
US. Dept. of Agriculture.

Persson, S., H. Wei, J. Milne, G.P. Page, and CR. Somerville. 2005. Identification of
genes required for cellulose synthesis by regression analysis of public microarray
data sets. Proc. Nat. Acad. Sci. USA. 102(24): 8633-8638.

Pilate, G., E. Guiney, K. Holt, M. Petit-Conil, C. Lapierre, J.C. Leple, B. Pollet, I. Mila,
E.A. Webster, H.G. Marsttrop, D.W. Hopkins, L. Jouanin, W. Boerjan, W.
Schuch, D. Comu, and C. Halpin. 2002. Field and pulping performances of
transgenic trees with altered lignification. Nat. Biotech. 20(6): 607-612.

Qi, B., T. Fraser, S. Mugford, G. Dobson, O. Sayanova, J. Butler, J.A. Napier, A.K.
Stobart, and CM. Lazarus. 2004. Production of very long chain polyunsaturated
omega-3 and omega-6 fatty acids in plants. Nat. Biotech. 22(6): 739-745.

Ragauskas, A.J., C.K. Williams, B.H. Davison, G. Britovsek, J. Caimey, C.A. Eckert,
W.J. Frederick, J.P. Hallett, D.J. Leak, C.L. Liotta, J.R. Mielenz, R. Murphy, R
Templer, and T. Tschaplinski. 2006. The Path Forward for Biofuels and
Biomaterials. Science 311(5760): 484-489.

Ralph, J. 2006. What makes a good monoligol substitute? In The Science And Lore of the
Plant Cell Wall: Biosynthesis, Structure And Function, ed. T. Hayashi, 367.
Brown Walker Press (FL).

Ralph, J., T. Akiyama, H. Kim, F. Lu, P.F. Schatz, J.M. Marita, S.A. Ralph, M.S.S.
Reddy, F. Chen, and RA. Dixon. 2006. Effects of Coumarate 3-Hydroxylase
Down-regulation on Lignin Structure. J. Biol. Chem. 281(13): 8843-8853.

Ransom, CB. 2007. Production and Analysis of Biologically-Active Cellulases far
Ethanol Fuel in Maize Biomass. Michigan State University.

Ransom, C.B., V. Balan, G.C.G. Biswas, B.E. Dale, E. Crockett, and MB. Sticklen.
2007. Heterologous Acidothermus cellulolyticus 1,4-fi-Endoglucanase El
Produced Within the Corn Biomass Converts Corn Stover Into Glucose. Appl.
Biochem. Biotech. 36-1402207-220.

Reddy, M.S.S., F. Chen, G. Shadle, L. Jackson, H. Aljoe, and RA. Dixon. 2005.
Targeted down-regulation of cytochrome P450 enzymes for forage quality

124

improvement in alfalfa (Medicago sativa L.). Prac. Nat. Acad. Sci. USA. 102(46):
16573-16578.

Reggi S., S. Marchetti, T. Patti, F. De Amicis, R. Cariati, B. Bembi, and C. Fogher. 2005.
Recombinant human acid B-glucosidase stored in tobacco seed is stable, active
and taken up by human fibroblasts. Plant Mal. Biol. 57(1): 101-113.

Renewable Fuels Association. 2007. Building New Horizons: Ethanol Industry Outlook
2007.

RFA - The Industry - Plant Locations. http://www.ethanolrfa.org/industry/locations/
(accessed August 5, 2007).

Richards, RA. 2000. Selectable traits to increase crop photosynthesis and yield of grain
crops. J. Exp. Bot. 51, no. suppl_l: 447-458.

Ridley, B.L., M.A. O'Neill, and D. Mohnen. 2001. Pectins: structure, biosynthesis, and
oligogalacturonide-related signaling. Phytachem. 57(6): 929-967.

Saghai-Maroof, M. A., K. M. Soliman, R. A. Jorgensen, and R. W. Allard. 1984.
Ribosomal DNA Spacer-Length Polymorphisms in Barley: Mendelian
Inheritance, Chromosomal Location, and Population Dynamics. PNAS 81, no. 242
8014-8018.

Sahrawy, M., C. Avila, A. Chueca, F.M. Canovas, and J. Lopez-Gorge. 2004. Increased
sucrose level and altered nitrogen metabolism in Arabidopsis thaliana transgenic

plants expressing antisense chloroplastic fructose-1,6-bisphosphatase. J. Exp. Bot.
55(408): 2495-2503.

Salehi, H., C.B. Ransom, H.F. Oraby, Z. Seddighi, and MB. Sticklen. 2005. Delay in
flowering and increase in biomass of transgenic tobacco expressing the
Arabidopsis floral repressor gene FLOWERING LOCUS C. J. Plant Phys. 162,
no. 6: 711-717.

Sambrook, J ., E.F. Fritsch, and T. Maniatis. 1989. Molecular cloning. A laboratory
manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press.

Saruul, P., F. Srienc, D.A. Somers, and D.A. Samac. 2002. Production of a
Biodegradable Plastic Polymer, Poly-B-Hydroxybutyrate, in Transgenic Alfalfa.
Crop. Sci. 42(3): 919-927.

Schillberg, Fischer, and Emans. 2003. Molecular farming of recombinant antibodies in
plants. Cell. Mal. Life Sci. 60(3): 433-445.

125

Schillberg 8., S. Zimmermann, A. Voss, and R. Fischer. 1999. Apoplastic and cytosolic
expression of full-size antibodies and antibody fragments in Nicatiana tabacum.
Trans. Res. 8(4): 255-263.

Schulman, AH. 2002. Transgenic plants as producers of modified starch and other
carbohydrates. In Plant Biotechnology and Transgenic Plants, ed. K.M. Oksman-
Caldenetey and W.H. Barz, 255-282. New York: Basel.

Shapouri, H., J.A. Duffield, and M. Wang. 2002. The Energy Balance of Corn Ethanol:
An Update. U. S. Dept. of Agriculture.

Sheldon, C.C., J.E. Burn, P.P. Perez, J. Metzger, J.A. Edwards, W.J. Peacock, and ES.
Dennis. The FLF MADS Box Gene: A Repressor of Flowering in Arabidopsis
Regulated by Vemalization and Methylation. Plant Cell 11(3): 445-458.

Singh, S.P., E. Ekanem, T. Wakefield Jr., and S. Comer. 2003. Emerging Importance of
Bio-Based Products and Bio-Energy in the US. Economy: Information
Dissemination and Training of Students. Int. Food Agribus. Mgmt. Rev. 5(3): 1-
15.

Smidansky E.D., J.M. Martin, C.L. Hannah, A.M. Fischer, and M.J. Giroux. 2003. Seed
yield and plant biomass increases in rice are conferred by deregulation of
endosperm ADP-glucose pyrophosphorylase. Planta 216(4): 656-664.

Sticklen, MB. 2004. Production of microbial hydrolysis enzymes in biomass craps via
genetic engineering. In: Proceedings of the 2nd International Ukrainian
Conference on Biomass for Energy. Sept. 20-22, 2004. Kyiv, Ukraine. Ukraine
Natl. Acad. Sci. Press. pp. 133-136.

Sticklen, MB. 2006. Plant genetic engineering to improve biomass characteristics for
biofuels. Curr. Opin. Biotech. 17(3): 315-319.

Sticklen, M.B. 2007a. Feedstock Crop Genetic Engineering for Alcohol Fuels. Crop Sci:
in review.

 

. 2007b. Role of Transgenic Biomass Crops in Ethanol Refineries. J. Biabased
Mater. Bioenergy: xx-xx in review.

Teymouri, F., H. Alizadeh, L. Laureano-Perez, B.E. Dale, and MB. Sticklen. 2004.
Effects of Ammonia Fiber Explosion Treatment on Activity of Endoglucanase
from Acidothermus cellulolyticus in Transgenic Plant. Appl. Biochem. Biotech.
11621183-1192.

Thurston, CF. 1994. The structure and function of fungal laccases. Microbial. 140219-26.

Tucker, MP, A. Mohagheghi, K. Grohmann, and ME. Himmel. 1989. Ultra-
Thermostable Cellulases From Acidothermus cellulolyticus: Comparison of

126

Temperature Optima with Previously Reported Cellulases. Bio/Techno]. 7(8):

8 17-820.
U.S. Prime Supplier Sales Volumes of Petroleum Products.
http2//tonto.eia.doe.gov/dnav/pet/pet_cons_prim_dcu_nus_a.htrn (accessed

August 5, 2007).

Vain, P., MD. McMullen, and J.J. Finer. 1993. Osmotic treatment enhances particle
bombardment-mediated transient and stable transformation of maize. Plant Cell
Rep. 12:84-88.

Warren, RAJ. 1996. Microbial Hydrolysis of Polysaccharides. Annu. Rev. Microbial.
50(1): 183-212.

World of Com 2007. http://www.ncga.com/WorldOfCorn/main/consumptionl.asp
(accessed August 5, 2007).

World of Corn 2007. http2//www.ncga.com/WorldOfCorn/main/productionl.asp
(accessed August 5, 2007).

Wyman, CE. 1999. Biomass ethanol: Technical progress, opportunities, and commercial
challenges. Annu. Rev. Energy Environ. 24:189-226.

Wyman, C.E., B.E. Dale, RT. Elander, M. Holtzapple, M.R. Ladisch, and Y.Y. Lee.
2005a. Comparative sugar recovery data from laboratory scale application of
leading pretreatment technologies to corn stover. Biores. Tech. 96(18): 2026-
2032.

 

. 2005b. Coordinated development of leading biomass pretreatment technologies.
Biores. Tech. 96(18): 1959-1966.

Xu, D., X. Duan, B. Wang, B. Hong, T.H.D. Ho, and R. Wu. 1996. Expression of a Late
Embryogenesis Abundant Protein Gene, H VA 1 , from Barley Confers Tolerance to
Water Deficit and Salt Stress in Transgenic Rice. Plant Physiol. 110(1): 249-257.

Yang P., Y. Wang, Y. Bai, K. Meng, H. Luo, T. Yuan, Y. Fan, and B. Yao. 2007.
Expression of xylanase with high specific activity from Streptomyces
alivaceoviridis A1 in transgenic potato plants (Salanum tuberasum L.). Biotech.
Lett. 29(4): 659-667.

Yao, J O. 2004. Genetic transformation of tobacco with a beta-glucosidase gene to
induce constitutive systemic acquired resistance against tobacco mosaic virus.
Western Michigan University, Kalamazoo, MI.

Zhang, Y.H.P., and LR. Lynd. 2006. A functionally based model for hydrolysis of
cellulose by fungal cellulas. Biotechnol. Bioeng. 94(5): 888-898.

127

Zhong, H., F. Teymouri, B. Chapman, S.B. Maqbool, R. Sabzikar, Y. El-Maghraby, B.E.
Dale, and MB. Sticklen. 2003. The pea (Pisum sativum L.) rch transit peptide
directs the Alcaligenes eutraphus polyhydroxybutyrate enzymes into the maize
(Zea mays L.) chloroplasts. Plant Sci. 165(3): 455-462.

Ziegelhoffer, T., J .A. Raasch, and S. Austin-Phillips. 2001. Dramatic effects of truncation
and sub-cellular targeting on the accumulation of recombinant microbial cellulase
in tobacco. Mal. Breeding 8(2): 147-158.

Ziegelhoffer T., J. Will, and S. Austin-Phillips. 1999. Expression of bacterial cellulase
genes in transgenic alfalfa (Medicago sativa L.), potato (Salanum tuberosum L.)
and tobacco (Nicatiana tabacum L.). Mal. Breeding 5(4): 309-318.

Ziegler, M.T., S.R. Thomas, and K.J. Danna. 2000. Accumulation of a thermostable

endo-1,4-B-D-glucanase in the apoplast of Arabidopsis thaliana leaves. Mal.
Breeding 6(1): 37-46.

128

   

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