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Michig mate
University

 

 

 

This is to certify that the
dissertation entitled
MANIPULATION OF THE GENES AND ENZYMES OF THE
SHIKIMATE PATHWAY

presented by

Sunil S. Chandran

has been accepted towards fulfillment
of the requirements for

Ph.D. degree in CHEMISTRY

fl W/

Major professor

Date January 11, 2001

MS U is an Affirmative Action/Equal Opportunity Institution 0-12771

PLACE IN RETURN BOX to remove this checkout from your record.
TO AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

 

DATE DUE

DATE DUE

DATE DUE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

6/01 cJCIRClDaIeDuepss-pjs

 

MANIPULATION OF THE GENES AND ENZYMES OF THE
SHIKIMATB PATHWAY
By

Sunil S. Chandran

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Department of Chemistry

2000

 

The ~
\iurnins ha
disposition i
driersc ranj

manipulaiet

 

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prmiding ill
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DHQ synti
the Zn‘: 3]

On

for [he mi

ABSTRACT

MANIPULATION OF THE GENES AND ENZYMES OF THE
SHIKIMATE PATHWAY
By

Sunil S. Chandran

The shikimate pathway for the biosynthesis “of aromatic amino acids and aromatic
vitamins has been the target of intensive research over the last decade. Its central
disposition in the life-cycle of a microbial or plant cell has been well established. The
diverse range of enzymes and genes responsible for each step in this pathway can be
manipulated to suit various purposes. For example, one or more enzymes in the
shikimate pathway can be inhibited to render the cell unable to sustain growth, thereby
providing an option for antibiotic or herbicidal design. The enzyme DHQ synthase is a
prime candidate for this purpose. In the present study, inhibition of DHQ synthase has
been attempted by taking advantage of the metal co-factor in the active Site.
Protocatechuic acid, catechol, and derivatives of these aromatics, sharing the common
element of an ortho dihydroxylated benzene ring were synthesized. These molecules
allowed examination of the role that the metal co-factor plays in the binding properties of
DHQ synthase. The results indicated marked differences in binding properties between
the Zn+2 and Co+2 metalloforms of DHQ synthase. These inhibitors provided evidence
attesting to the important role that the active site metal played in substrate binding.

On the other hand, instead of inhibiting the shikimate pathway, it can be exploited

for the microbial synthesis of industrially applicable chemicals. A major portion of this

 

thesis inie~

>liiklllllk. at.

 

synthetic so:
iennentaiio:
limiting in.
alleviated.

glucose uptu-
in 27‘} yield

Anei

 

 

 

fornication dL
fermentation
most lilch

1nSUL'menlnl

thesis investigates the development of a biocatalytic process for the production of
Shikimic acid, which has recently emerged as a suitable starting material for a number of
synthetic schemes. Synthesis of Shikimic acid was successfully achieved under fed-batch
fermentation conditions, utilizing various recombinant Escherichia coli biocatalysts. The
limiting factors for obtaining high Shikimic acid titers and yields were identified and
alleviated. Accounting for low intracellular E4P concentration and loss of PEP due to
glucose uptake, proved to be key determinants. Shikimic acid titer of 71 g/L was realized
in 27% yield.

Attempts were also made at understanding the mechanism of quinic acid
formation during microbial synthesis of Shikimic acid. Uptake of shikimic acid from the
fermentation broth followed by its in vivo processing to quinic acid was shown to be the
most likely source of the contamination. Identifying this role of shikimate uptake was

instrumental in developing methods designed to eliminate production of quinic acid.

 

Copyright by
Sunil S. Chandran

2000

 

To my family

For their love and support

 

Tin
guidance. .
have learn:
aggressixe:
addition. I
llaleczka.
the preparu

1 ar
stages in n
“inch she
gratitude it
\lontcham
Saliam’ahe
Barton. M
lam fOI'IUn
Padmesh V

has
Pifiems anc
me under 2

h .- .
Lens 13 d5(

 

ACKNOWLEDGMENTS

The first person I would like to thank is Prof. John Frost for his patience,
guidance, and encouragement throughout the course of my graduate career. From him I
have learned the valuable lesson that to practice science you have to live science. His
aggressiveness towards challenging problems has been a source of inspiration for me. In
addition, I would like to thank the members of my graduate committee, Prof. Robert E.
Maleczka, Prof. Mitch Smith, and Prof. Katherine Hunt for their intellectual input during
the preparation of this thesis.

I am especially grateful to Dr. Karen Draths for her invaluable insight at various
Stages in my projects. I thank her for allowing me to pursue the Shikimic acid project,
which she and Dave Knop were instrumental in developing. I would like to extend my
gratitude towards the past and present members of the group including Dr. Jean-Luc
Montchamp, Dr. Feng Tian, Dr. John Arthur, Dr. Kai Li, Dr. Spiros Kambourakis,
Satyamaheshwar Peddibotla, Jian Yi, Jiantao Guo, Ningqing Ran, Wei Niu, Tom
Bannon, Michael Gyamerah, and Mapitso Molefe for their assistance and their friendship.
I am fortunate to have shared these years in the company of Jessica Barker, Chad Hansen,
Padmesh Venkitasubramanian, and Dave Knop, whose friendship I will always cherish.

Last but not least I would like to thank the most important people in my life, my
parents and sister for their love and support in all my endeavors. Their undying faith in
me under all circumstances Spurred me to keep trying, irrespective of the odds. This

thesis is dedicated to them.

-vi-

 

TABLE OF CONTENTS

LIST OF TABLES .......................................................................................................... x
LIST OF FIGURES ........................................................................................................ xi
LIST OF ABBREVIATIONS ........................................................................................ xv
HAPTE 1 ................................................................................................................... 1
INTRODUCTION ....................................................................................................... l
The Shikimate Pathway ........................................................................................ 2
Derailing the Shikimate Pathway: A Viable Option for
Herbicidal and Antimicrobial Agents .................................................................... 5
Inhibition Studies of DHQ synthase ...................................................................... 7
Biocatalytic Synthesis of Value-Added Chemicals .............................................. 10
Metabolic Engineering of Escherichia coli for Optimization of
Product Titer and Yield ....................................................................................... 28
HAPTER 2 ................................................................................................................. 37
INHIBITION OF DHQ SYNTHASE VIA UTILIZATION OF METAL-DIOL
INTERACTIONS ...................................................................................................... 37
Background ........................................................................................................ 37
Rationale Behind Inhibitor Design ...................................................................... 41
Synthesis of Phosphonate 4 and Homophosphonates 5-6 .................................... 45
Inhibition Studies of 4-8 with DHQ synthase ...................................................... 47
Interpretation of the Inhibition Pattern Exhibited by Aromatic Inhibitors 4-8 ...... 49
HAPTER ................................................................................................................. 52
BIOCATALYTIC SYNTHESIS OF SI-IIKIMIC ACID USING RECOMBINANT
ESCHERICHIA COLI ............................................................................................... 52
Introduction ........................................................................................................ 52
Biocatalysts and Fed-Batch Fermentation Conditions ......................................... 54
A) Shared Genomic Traits and Plasmid Elements ........................................ 54
B) Fed-Batch Fermentor Conditions ............................................................. 57
Overexpression of Transketolase and Transaldolase Under
Fed-Batch Fermentor Conditions ........................................................................ 59
Production of SA as a Function of Increased PEP availability ............................. 69
Examination of Lowered Osmotic Stress on SA Production ................................ 85
Comparison of SA Production Capabilites Between ............................................ 91

-vii-

 

 

CHAPITJQ
lNVESTI
lntrt‘“
Fedl
Séct
Sour

 

Stor
(:uh
(Sen
(Ana
(3er

E. coli B and E. coli K12 .................................................................................... 91

Discussion .......................................................................................................... 96
A) Comparison of Titers and Yields ............................................................. 96

B) Effects of Increased PEP Availability ...................................................... 98

C) E. coli B v/s E. coli K12 ........................................................................ 100
Synthesis of Phenol from Shikimic Acid ........................................................... 102
HAPTER 4 ............................................................................................................... 104
INVESTIGATIONS INTO SHIKIMIC ACID—QUINIC ACID EQUILIBRATION. 104
Introduction ...................................................................................................... 104
Fed-Batch Fermentor Conditions ...................................................................... 106
SA/QA Equilibration: Is Uptake of SA the ....................................................... 107
Source of QA Contamination? .......................................................................... 107
Experimental Evidence Supporting QA Formation Via SA Uptake ................... 115
Discussion ........................................................................................................ 125
HAPTER ............................................................................................................... 130
EXPERIMENTAL .................................................................................................. 130
General Methods .............................................................................................. 130
General Chemistry ............................................................................................ 130
Reagents and Solvents ...................................................................................... 130
Chromatography ............................................................................................... 13 l
Spectroscopic and Analytical Measurements ..................................................... 131
Enzyme Assays ................................................................................................. 132
General Information .................................................................................... 132
DAHP synthase assay ................................................................................. 133
Transketolase activity measurement ............................................................ 134
Bacterial Strains ............................................................................................... 135
Storage of Bacterial Strains and Plasmids ......................................................... 135
Culture Medium ............................................................................................... 135
General Fed-Batch Fermentor Conditions ......................................................... 137
Analysis of Fermentation broth ......................................................................... 138
Genetic Manipulations ...................................................................................... 144
General Information .................................................................................... 144
Large Scale Purification of Plasmid DNA ................................................... 145
Small Scale Purification of Plasmid DNA ................................................... 146
Restriction Enzyme Digestion of DNA ....................................................... 147
Agarose Gel Electrophoresis ....................................................................... 148
Isolation of DNA from Agarose .................................................................. 148
Treatment of DNA with Klenow fragment .................................................. 149
Treatment of Vector DNA with Calf Intestinal Alkaline Phosphatase .......... 149
Ligation of DNA ......................................................................................... 150
Preparation and Transformation of Competent Cells ................................... 150
Synthetic Procedures ........................................................................................ 152

- viii -

 

Strut

QC.

Cup

BlBLlO(

Synthesis of PCA phosphonate 4 and PCA homophosphonate 5 .................. 152

Synthesis of catechol homophosphonate 6 .................................................. 157
Enzyme Kinetics ............................................................................................... 159
Determination of Inhibition Constants (K) for Inhibitors of DHQ synthase. 159
Strain Constructions ......................................................................................... 159
Strain SP1.1 ................................................................................................ 159
Strain EB 1.1 ............................................................................................... 160
Strain SPl.lpts ........................................................................................... 161
Plasmid pKDlZ.O36A ................................................................................. 161
Plasmid pKDlZ.047A ................................................................................. 162
Plasmid pKD12.112A ................................................................................. 162
Plasmid pKD12. 138A ................................................................................. 162
Plasmid pKD15.07lB ................................................................................. 163
Plasmid pSC5.112B .................................................................................... 163
Plasmid pSC6.090B .................................................................................... 163
Plasmid pSC6.14ZB .................................................................................... 164
Plasmid pSC6. 162A .................................................................................... 164
Plasmid pSC6.301A .................................................................................... 165
Strains Constructions ........................................................................................ 165
Strain SC1.0 ................................................................................................ 165
Strain SP1.lshiA ......................................................................................... 166
Plasmid pSC5.214A .................................................................................... 166
BIBLIOGRAPHY ................................................................................................... 167

-ix-

 

 

Table 1. ln
forms of D

Title}. D
Table} D
TableJ. I

Table 5. E
SA b} SP1

Table 6. T

Tablei. C_
EBl.l andi

Tiblfi 8 C;
EBl.l andf

Table 9, 1 i

 

LIST OF TABLES

Table 1. Inhibition constants (uM) for binding to the Co+2 and Zn+2
forms of DHQ synthase. ................................................................................................ 48

Table 2. DAHP synthase activities (umol/min/mg) for PEP limited biocatalysts. .......... 66

Table 3. DAHP synthase activities (umol/min/mg) for non-PEP limited biocatalysts. 74

Table 4. Titers and yields for SA production by SP1.1 based biocatalysts. .................... 84
Table 5. Effect of varying betaine concentrations on production of

SA by SP1.1/pKDlS.071B. ........................................................................................... 90
Table 6. Titers and yields for SA production by EB1.1 based biocatalysts. ................... 94

Table 7. Comparison of DAHP synthase activities (umol/min/mg) between
EBl.1 and SP1.1 biocatalysts. ....................................................................................... 95

Table 8. Comparison of transketolase activities (umol/min/mg) between
EB1.1 and SP1.1 biocatalysts. ....................................................................................... 95

Table 9. Titers and yields of SA produced under various glucose-limited conditions. . 114

 

Figure l. I
aromatic \t

 

Enm5.Q
Figureo. i
Figure'l. (
EgureS. t
Figure 9. I
Figure 10,
F1:lure 11.

Figure 12.
Tim Phent

figure 13,

 

LIST OF FIGURES

Figure 1. The common pathway of aromatic amino acid and

aromatic vitamin biosynthesis. ........................................................................................ 4
Figure 2. Compounds proven to disrupt the common pathway. ....................................... 6
Figure 3. Proposed mechanism of DHQ synthase. .......................................................... 7
Figure 4. Comparison of chemical and biocatalytic routes to acrylamide. ..................... 11
Figure 5. Comparison of chemical and biocatalytic synthesis of L-lactic acid. .............. 13
Figure 6. Structures of L-lysine and aspartame .............................................................. 13
Figure 7. Chemicals accessible from the aromatic amino acids. .................................... 14
Figure 8. Chemical synthesis of indigo. ........................................................................ 15
Figure 9. Biocatalytic synthesis of indigo from D-glucose. ........................................... 16
Figure 10. Chemical production of vanillin from catechol. ........................................... 17
Figure 11. Biocatalytic synthesis of vanillin from D-glucose .......................................... 17

Figure 12. Comparison of the chemical and microbial synthesis of PHB

from phenol and D-glucose respectively. ....................................................................... 19
Figure 13. Comparison of chemical and microbial catalyzed synthesis of PABA. .......... 20
Figure 14. Microbial synthesis of QA. .......................................................................... 21
Figure 15. Various synthetic routes to hydroquinone. ................................................... 22
Figure 16. The various synthetic applications of DHS. ................................................. 23
Figure 17. Conversion of D-glucose to gallic acid and pyrogallol .................................. 24
Figure 18. Comparison of chemical and biocatalytic routes to catechol ......................... 26
Figure 19. Chemical and microbial synthetic routes to adipic acid. ............................... 27
Figure 20. Reactions catalyzed by transketolase (TktA) and transaldolase (TalB). ........ 31
Figure 21. The PT S-3ystem for glucose uptake. ............................................................ 32

-xi-

Figure 33.

1“
lisure -2.
b

 

Ezure 34.

5

for glueo~e
Figure 35.
Fizure 36..

5

Figure 37.

Figure 28.

Figure 39.

 

Figure 3.0,
Figure 3.1 _

. a
HEW :2.

Figure 33.
Figure 3.1

Figure 35.

Figure ,

Figure 37.

Figure 38

Figure 39

HEW40

Fig

F

F

figure 44

"J
CIN

sure 41
”Elite .13

lngIe 43

Figure 22. Phosphorylation of D—glucose during its uptake. .......................................... 33
Figure 23. Reaction catalyzed by PEP synthase. ........................................................... 34
Figure 24. Comparison of the PTS and facilitated diffusion systems

for glucose uptake. ........................................................................................................ 35
Figure 25. Active site of DHQ synthase ........................................................................ 40
Figure 26. Interactions postulated to occur at the active Site of DHQ synthase. ............. 41
Figure 27. Inhibitors functioning via complexation of Zn”. .......................................... 42
Figure 28. Unsaturated inhibitors of DHQ synthase. ..................................................... 43
Figure 29. Aromatic compounds designed for inhibition of DHQ synthase. .................. 44
Figure 30. Synthesis of PCA phosphonate 4 and PCA homophosphonate 5. ................. 46
Figure 31. Synthesis of catechol homophosphonate 6. .................................................. 47
Figure 32. Dixon plot for PCA phosphonate 4. ............................................................. 48
Figure 33. Neuraminidase inhibitor GS4104 ................................................................. 52
Figure 34. Strain SP1.1 and Plasmid pKD12.112A ....................................................... 56
Figure 35. Preparation of Plasmid pKD12.036A. .......................................................... 60
Figure 36. Preparation of Plasmid pKDlZ.047A. .......................................................... 61
Figure 37. Preparation of Plasmid pKD12.112A. .......................................................... 62
Figure 38. SP1.1/pKD12.112A fed-batch fermentation time course. ............................. 63
Figure 39 . Preparation of Plasmid pKD12.138A. .......................................................... 64
Figure 40. SP1.1/pKDlZ.138A fed-batch fermentation time course. ............................. 65
Figure 41. Preparation of Plasmid pSC4.295A .............................................................. 67
Figure 42. SP1.1/pSC4.295A fed-batch fermentation time course. ................................ 68
Figure 43. Preparation of Plasmid pKD15.07lB. .......................................................... 70
Figure 44. SP1.1/pKD15.07lB fed-batch fermentation time course. ............................. 71

-xii-

 

Fisure 45.

Figure 46.

Figure 48.
Figure 49.
Figure 50.
Figure 51.
Figure 52.
Figure 53.
Figure 54.
Fltitre 55.

figure 56.
:02 SA [0

Dmfil

Flé‘ure 58.

10 mil bet,
l

FlEJUre 59.
mmMN

 

Figure 60,
30 m“ b6!

EWML

5m“;

FiEUre 63_

Figure 64.

F1titre 65_

“Wm

Figure 45. Preparation of Plasmid pSC5.112B. ............................................................. 72
Figure 46. SP1.1/pSC5.112B fed-batch fermentation time course. ................................ 73
Figure 47. Preparation of Plasmid pSC6.090B. ............................................................. 75
Figure 48. SP1.1pts/pSC6.090B fed-batch fermentation time course. ........................... 77
Figure 49. SP1.1/pSC6.090B fed-batch fermentation time course. ................................ 77
Figure 50. Preparation of Plasmid pSC6.142B. ............................................................. 79
Figure 51. Preparation of Plasmid pSC6.162A .............................................................. 80
Figure 52. SP1.1/pSC6.162A fed-batch fermentation time course. ................................ 81
Figure 53. Preparation of Plasmid pSC6.301A .............................................................. 82
Figure 54. SP1.1pts/pSC6.301A fed-batch fermentation time course. ........................... 83
Figure 55. SP1.1/pSC6.301A fed-batch fermentation time course. ................................ 84
Figure 56. Time course for fed-batch fermentation experiment involving addition of

20 g SA to SP1.1/pKD15.071B culture .......................................................................... 86
Figure 57. Common osmoprotectants used in microbial fermentations .......................... 87

Figure 58. SP1.1/pKD15.071B fed-batch fermentation time course with

10 mM betaine. ............................................................................................................. 88
Figure 59. SP1.1/pKD15.071B fed-batch fermentation time course with

100 mM betaine. ........................................................................................................... 89
Figure 60. SP1.1/pKD15.071B fed-batch fermentation time course with

30 mM betaine. ............................................................................................................. 89
Figure 61. EBl.l/pKD12.112A fed-batch fermentation time course. ............................ 92
Figure 62. EBl.l/pKD12.l38A fed-batch fermentation time course. ............................ 93
Figure 63. EB 1.1/pKD15.071B fed-batch fermentation time course .............................. 94
Figure 64. Comparison of phenol synthesis from benzene and D-glucose .................... 103

Figure 65. SP1.1/pKDlZ.112A fed-batch fermentation time course.
Glucose limited conditions, K. = 0.1. ........................................................................... 104

- xiii -

Fzgure 66.

Figure 67.
Glucose l:'

Figure 63'.

Figure 69.
Glucose ll”

Figure 70.
Glucose li:

EHETT
Glucose lii

 

Figure 724
Figure ?3_
Figure 74.
Figure 75

Figure 76
Glucose l

Flgtlre 77
Glucose l
IDESA:

FlSure 7i
1098A

FlElite 7

Figure 66.

The two possible pathways leading to QA production ................................ 108

Figure 67. SP1.1/pKD12.112A fed-batch fermentation time course.

Glucose limited conditions, K. = 0.8 ............................................................................ 110
Figure 68. Structure of methyl-Ot-D-glucopyranoside (MGP). ..................................... 111
Figure 69. SP1.1/pKD12.112A fed-batch fermentation time course.

Glucose limited conditions, Kc = 0.1, lmM MGP. ....................................................... 112
Figure 70. SP1.1/pKD12.138A fed-batch fermentation time course.

Glucose limited conditions, Kc = O. 1, 0 mM MGP. ...................................................... 113
Figure 71. SP1.1/pKD12.138A fed-batch fermentation time course.

Glucose limited conditions, K. = 0.1, 1 mM MGP. ...................................................... 114
Figure 72. Construction and selection of SC1.0. ......................................................... 116
Figure 73. The modified shikimate pathway in SC1.0 ................................................. 116
Figure 74. Effect of inactivating DAHP synthase in SP1.1 .......................................... 119
Figure 75. Preparation of Plasmid pSC5.214A ............................................................ 120
Figure 76. SP1.1/pSC5.214A fed-batch fermentation time course.

Glucose limited conditions, no extra aromatic amino acids added ................................ 121
Figure 77. SP1.1/pSC5.214A fed-batch fermentation time course.

Glucose limited conditions; aromatic amino acids added at 0 h, 18 h and 30 h;

10 g SA added at 12 h. ................................................................................................ 122
Figure 78. SP1.1shiA/pSC5.214A fed-batch fermentation time course.

Glucose limited conditions; aromatic amino acids added at 0 h, 18 h and 30 h;

10 g SA added at 12 h. ................................................................................................ 124
Figure 79. SP1.13hiA/pKDlZJ38A fed-batch fermentation time course.

Glucose limited conditions, KC = 0.1, 0 mM MGP. ...................................................... 125
Figure 80. Crude 1H NMR of fermentation broth. ....................................................... 140
Figure 81. 1H NMR of Shikimic acid. ......................................................................... 141
Figure 82. 1H NMR of 3-dehydroshikimic acid ........................................................... 142
Figure 83. 1H NMR of quinic acid. ............................................................................. 143

-xiv-

 

A:
ADP
AMP
Ap

KW

Bu

GAP
Cm
CONII
BAH
DAHP
DCL'
DEAE
DHQ

DHS

Du
El

E4P

 

Ac

AMP
Ap

ATP

Bu

CI
CIAP
Cm
COMT
DAH
DAHP
DCU

‘ DEAE
DHQ
DHS
DO
DTT
El

E4?

LIST OF ABBREVIATIONS

acetyl

adenosine diphosphate

adenosine monophosphate

ampicillin

adenosine triphosphate

base pair

butyl

chemical ionization

calf intestinal alkaline phosphatase
chloramphenicol
catechol-O-methyltransferase
3-deoxy-D-arabino-heptulosonic acid
3-deoxy-D-arabin0-heptulosonic acid 7- phosphate
digital control unit

diethylaminoethyl

3-dehydroquinate

3-dehydroshikimic acid

dissolved oxygen

dithiothreitol

electron impact

D-erythrose 4-phosphate

-xv-

 

EPSP
FAB

FBR

GA

Glu

it
LB
it
Me
MGP

Mic

Hm

MOPS

 

EPSP

FAB

FBR

GA

Glu

His

HMPA

HRMS

Kan

MGP

MIC

MOPS

5-enolpyruvylshikimate 3-phosphate
fast atom bombardment

feed back resistant

fourier transform

gallic acid

glutamate

hour

histidine

hexamethyl phosphoramide

high resolution mass spectrometry
kanamycin

inhibition constant

Michaelis constant

rate constant

kilogram

luria broth

molar

methyl

methyl a-D-glucopyranoside
minimal inhibitory concentration
milliliter

millimolar

4-morpholinepropanesulfonic acid

-xvi-

 

.\lS
min
SAD
NADH
XADP
.\'.\lR
OD
PABA
PCA
PEG

PEP

Ppm

Pps

PTS

MS

NAD
NADH
NADP
NMR
OD
PABA
PCA
PEG
PEP
Ph
PHB
PID

PCR

PPm

iPr

PTS

QA

SA

mass spectrometry

minute

nicotinamide adenine dinucleotide, oxidized form
nicotinamide adenine dinucleotide, reduced form
nicotinamide adenine dinucleotide phosphate, oxidized form
nuclear magnetic resonance

optical density

p-aminobenzoic acid

protocatechuic aid

polyethylene glycol

phosphoenolpyruvate

phenyl

p-hydroxybenzoic acid
proportional-integral-derivative

polymerase chain reaction

parts per million

PEP synthase

isopropyl

phosphotransferase system

quinic acid

room temperature

revolutions per minute

Shikimic acid

- xvii -

 

SDS

Te

DIS

TSP

l'i'

 

SDS

Tc

THF

TMS

TSP

UV

sodium dodecyl sulfate

tetracycline

tetrahydrofuran

trimethylsilyl

sodium 3-(trimethylsilyl)propionic-2,2,3,3-d4

ultraviolet

- xviii -

 

 

l
the shikiri
microorg.-.
understun
exploitatio
in the me
antimicroh
the other h

rtltt‘am Cl

 

studiing bi

The

CHAPTER 1

INTRODUCTION

The common pathway for aromatic amino acid biosynthesis, also referred to as
the shikimate pathway, is a vital metabolic cascade of events found in plants and
microorganisms. Researchers have for long studied this pathway, gaining a better
understanding of each enzyme catalyzed reaction. This understanding allowed
exploitation of the common pathway to suit a variety of purposes. Given its central role
in the metabolism of a cell, scientists have been successful at obtaining potent
antimicrobial activity by inhibition of one or more enzymes in this metabolic route. On
the other hand, the common pathway has also been manipulated to produce industrially
relevant chemicals at a low cost. The work reported in this thesis is an attempt at
studying both aspects of the shikimate pathway.

The medical profession is currently facing a formidable adversary with the
rampant emergence of antibiotic resistant bacteria. The search is underway to design new
and more potent drugs that can combat microbes impervious to present day treatment.
This can be achieved by targeting a pathway hitherto underutilized as a possible target for
drug design, the shikimate pathway. Chapter 2 of this thesis discusses inhibition of one
enzyme belonging to the common pathway, namely 3-dehydroquinate (DHQ) synthase.1
The second enzyme in this pathway, DHQ synthase belongs to a small, unique family of
enzymes. The uniqueness of the enzymes in this family is their catalytic utilization of
nicotinamide adenine dinucleotide (NAD).2 Enzymes in this class, which we will label as

NADzymes, also include myo-inositol l-phosphate (MIP) synthase,3

 

   
   
  
   

S‘udenosj
diphosphu
contenie'
ini'estigut;
be center:

the acute

resulting i
gallic aeiti

Common r

 

describes g
The focus
lQ-ll and

TDD] SlUCllt

S—adenosylhomocysteine hydrolase,4 S-ribosylhomocysteine hydrolase,5 and uridine
diphosphate galactose 4-epimerase.6 The catalytic role of N AD has been used as a very
convenient tool to achieve inhibition of DHQ synthase in the past. The current
investigation will also utilize this aspect of inhibition. However, the major emphasis will
be centered around trying to better understand the role that the metal co-factor, present in
the active Site, plays in binding to substrate and inhibitors.

The shikimate pathway has been a target for metabolic engineering in the past,
resulting in Strains capable of producing chemicals like 3-dehydroshikimic acid (DHS),
gallic acid, vanillin, etc. A major portion of this thesis deals with manipulation of the
common pathway in order to biocatalytically produce Shikimic acid (SA). Chapter 3
describes studies undertaken to improve the overall titer and yield of SA from D-glucose.
The focus of Chapter 4 is on efforts made to better understand the source of quinic acid
(QA) and DHS contamination commonly seen in SA fermentations. Results arising from

both studies will ultimately result in a more efficient SA producing biocatalyst.

The Shikimate Pathway

The shikimate pathway present in plants and microorganisms is responsible for
the biosynthesis of the aromatic amino acids and aromatic vitamins.7 It is made up of
seven enzyme-catalyzed reactions culminating into the biosynthesis of chorismic acid
(Figure 1). Chorismic acid is processed further into L-tryptophan, L-tyrosine and
L-phenylalanine by three terminal pathways. Besides providing a metabolic route to the
aromatic amino acids, the common pathway also provides a means of synthesis of the

various isoprenoid quinones, the folic acid family of co-enzymes and enterochelin, via

pathtt 33‘s
lies in el;
intolsed '
iron sequ:

It:
iPEPt u 2:
acid T-p'n
Figure 1)
feedback i

GWH ence

 

 

 

 

l502}mes l
encoded E
reaction c
Tedllt‘lltin
the (NOE
Shikimate
3‘PDOSph;

amk’il

(

(IO

5-61101er
Slinlhacet
Chommat

pathways branching out from chorismic acid.7 The primary role of the quinone family
lies in electron transport while co-enzymes derived from folic acid are known to be
involved in the biosynthetic transfer of one carbon fragments. Enterochelin acts as an
iron sequestering agent involved in iron uptake in microorganisms.

The first reaction in the pathway is the condensation of phosphoenolpyruvate
(PEP) with D-erythrose 4-phosphate (E4P) resulting in 3—deoxy-D-arabino-heptulosonic
acid 7-phosphate (DAHP), a reaction catalyzed by the enzyme DAHP synthase
(Figure 1).7 Three isozymes of DAHP synthase exist in Escherichia coli, each of them
feedback inhibited by one of the three aromatic amino acids. The genes aroF, aroG, and
aroH encode the tyrosine-sensitive, phenylalanine-sensitive, and tryptophan-sensitive
isozymes of DAHP synthase respectively. DAHP is then converted to DHQ by aroB-
encoded DHQ synthase. Elimination of a water molecule from DHQ results in DHS, a
reaction catalyzed by the aroD-encoded enzyme, DHQ dehydratase.8 Subsequent
reduction of DHS by shikimate dehydrogenase affords SA. The enzyme is encoded by
the aroE gene and it utilizes NADPH to carry out the reaction.9 Two isozymes of
shikimate kinase catalyze the ATP aided phosphorylation of SA to generate shikimate
3-phosphate. The two isozymes of shikimate kinase are encoded by the aroLlo and
aroK” genes. Condensation of shikimate 3-phosphate with PEP to produce
S-enolpyruvylshikimate 3-phosphate (EPSP) is catalyzed by aroA-encoded EPSP

2 The final Step in the common pathway is catalyzed by aroC—encoded

synthase.I
chorismate synthase and involves loss of inorganic phosphate from EPSP resulting in

chorismic acid.”

 

i“
H203PO

 

Figure 1.
“Gimme,

Genenc 10
Oral). DH,
QTOA‘ EPS

cozn

ubiquinone ‘—
folic acid“—

X- OH PHB benzoic acid

H3PO4

   

 

_PP_S, 4kcozn H3PO4 HO," COZH H3P04 HO,” COZH
tal arof3 = OH aroB 0 . OH aroD
—-> OH O are no OH 5
OH
1’“ H203POMH arOH
2 Ft: P03H2; DAHP DHQ
E48“ R=H; DAH
0402+! NADPH NADP 0:02” ATP ADP 0°?” PEP H3Po4 ‘
o 3. OH 3’05 Ho" 5 OH Sigif H203P0" = OH 3'“
OH 0 OH ‘
DHS SA sap
cozI-I H3PO4 COZH COZH
NH2
JL aroC JL ——> -—-> __>L-tryptophan
H203P0" _ o COZH = o 002 H
OH OH anthranilic acid
EPSP chorismic acid

cozii
i

cozn HO"

OH
O

2 ,3-dihydroxy

L-phenylalanine
L-tyrosine

prephenic acid

X;NH2 PABA l

enterochelin

Figure 1 The common pathway of aromatic amino acid and aromatic Vitamin

biosynthesis.

Genetic loci are as follows: aroF aroG aroH, DAHP synthase; aroB, DHQ synthase
aroD DHQ dehydratase; aroE, shikimate dehydrogenase; aroL aroK, shikimate kinase;

aroA, EPSP synthase; aroC, chorismate synthase.

 

Di~

demonstru

and until

type cells
microbes d
necessars

mammaliu

 

Dl‘lQ 5}“!
lmplicuti-or
against bag
mUmITOns
mOCTClS 38
lmIIiUne Sh
An
alallabili]
DUOrOShik
p'aminobé
agalIlSI a r
to a mint

C0u1d be r

Gills; aron

Derailing the Shikimate Pathway: A Viable Option for Herbicidal
and Antimicrobial Agents

Disruption of the shikimate pathway has been proven to be detrimental to plants
and microorganisms. Microbes with a disrupted common pathway have been
demonstrated to be characterized by severely attenuated growth in comparison to wild
type cells when introduced into a mammalian system.” This result indicated that
microbes deficient in a functional common pathway have a difficult time scavenging the
necessary aromatic amino acids and vitamins from the rich growth environment of a
mammalian system. More recently, researchers also demonstrated that aroB encoded
DHQ synthase was required for microbial pathogen virulence.” Understanding the
implications of this huge body of work has ramifications in terms of developing vaccines
against bacterial infections. Live, attenuated strains of Salmonella typhimurium carrying
mutations in aroB", aroD”""6“, aroA‘W, and aroC“"‘b have been introduced into mouse
models as potential vaccines. The effect of the attenuated strains was to stimulate the
immune system towards infection caused by the wild type parent strain.

Among all the aromatic amino acids and aromatic vitamins, it appears that
availability of p-aminobenzoic acid is most critical. The compound (6S)-6-
fluoroshikimic acid is a potent antimicrobial agent, deriving its activity from inhibition of
p-aminobenzoic acid biosynthesis.17 It possesses an MIC value of less than lug mL'l
against a range of bacteria. Mice injected with (6S)-6-fluoroSA also exhibited immunity
to a number of bacterial infections. However, in vitro inhibition of bacterial growth
could be reversed by supplementation with p-aminobenzoic acid, but not with any of the

other aromatic amino acids or aromatic vitamins.

 

l

result in l

glyphosat

 

attributed
in l998.
fll’lt‘fimp/
tft'ectite
falt‘tpar“
l$€\ ETC C
resistant
313 Cliff:
bl Either
that COLT

C0militia

EDIE-me
be“slit.

amino 2

002H

,IF
(HoiziivNVcozH
Ho" , OH
OH
(6S)-6-fluoroshikimic acid glyphosate

Figure 2. Compounds proven to disrupt the common pathway.

Perhaps the best example to prove that inhibition of the shikimate pathway can
result in potent biological activity is the herbicide glyphosate. Marketed as Roundup®,
glyphosate is a postemergence, non-selective, broad spectrum herbicide.‘8 Its activity is
attributed to competitive inhibition of EPSP synthase. An important discovery was made
in 1998, indicating the presence of a functional common pathway in the phylum
Apicompleita.19 Agents capable of blocking the common pathway should therefore be
effective against parasites belonging to this family which include, Plasmodium
falciparum (malaria), Toxoplasma gondii (toxoplasmosis), and Cryptosporidium parvum
(severe diarrhea). In fact, glyphosate proved effective against malaria strains that were
resistant to an anti-malarial medicine, pyrimethamine, which interrupts folate processing
at a different point. But mice injected with lethal doses of T. gondii could not be rescued
by either glyphosate or pyrimethamine. However, doses of glyphosate or pyrimethamine
that could not protect the mice when used alone, rescued infected mice when used in
combination even when the mice were allowed to eat diets with folate.

This emphasized the viability of concomitantly targeting either single or multiple
enzymes along the common pathway with the goal of deriving medical and commercial
benefit. Mammals lack a functional shikimate pathway and rely on their diet for aromatic

amino acids and aromatic vitamins. Hence, using herbicides and antimicrobial agents

 

based on
With th:

understu':

I‘atultzir;4
carbocycl

acids and

 

-I ‘-
02321

I
Oi-

oxldfillg

lllfr . .
J‘gdm

based on disruption of the shikimate pathway will have minimal side effects on humans.
With this aim in mind, it would be of considerable importance to gain a better

understanding of the enzymes in this pathway.

Inhibition Studies of DHQ synthase

3-dehydroquinate synthase is the second enzyme of the Shikimate pathway
catalyzing the conversion of DAHP to DHQ. This step sets into place the six membered
carbocyclic unit which will ultimately form the benzenoid portion of the aromatic amino

acids and the secondary metabolites derived from the common pathway.

OH HO HO
_ 1 4 OH _ 0 OH _ OH
02 H 6 7 0P3.2 020 H 020 "sq
DAHP 0 er 0 O

A 00 {$9 a one

Figure 3. Proposed mechanism of DHQ synthase.

The mechanism20 for DHQ synthase begins with oxidation of the 05 alcohol of
DAHP utilizing enzyme-bound NAD, resulting in intermediate A and NADH. This
oxidation results in acidification of the C-6 proton facilitating easy elimination of

inorganic phosphate and formation of the enol ether intermediate B. Reduction of the

 

ietone 2
NAD. It 3
unmaski

con-dens :

stoicnio .
tumble

releases

to oxidat
bound to
is redox
80 nM.
metalloe
absorptit '

mDDOme

isotope e
during it
pOSTTIOnS
rate I 1 IT] 1
enlime;

“hasten

 

"_
—
—

_-_

ketone at C-5 by enzyme-bound NADH results in intermediate C and regeneration of
NAD, thereby completing the redox cycle. Ring opening of intermediate C results in
unmasking of the C-2 carbonyl in intermediate D, setting it up for an intramolecular aldol
condensation to afford DHQ.

Enzymes utilizing NAD as a co-factor in their catalytic cycle normally use it
stoichiometrically in the form of a co-substrate. The enzyme binds to the co-factor,
catalyzes the oxidation of substrate with concomitant reduction of NAD, and finally
releases the product and NAD. DHQ synthase on the other hand utilizes NAD
catalytically implying that the initial reduction of enzyme-bound NAD is always coupled
to oxidation of the NADH back to NAD during a single turnover. The NAD always stays
bound to the enzyme during its lifetime and the overall enzyme catalyzed transformation
is redox neutral. DHQ synthase binds to 1 equivalent of NAD with an apparent Km of
80 nM, the low value reflecting the efficiency of binding.” DHQ synthase is a

2°“ Atomic

metalloenzyme requiring either Zn*2 or Co+2 for its catalytic activity.
absorption analysis demonstrated that the enzyme requires one divalent metal cation per
monomer for activity.20a

Evidence for the mechanism (Figure 3) came from several observations : (1)
isotope exchange experiments20b proved that the C-6 proton was lost to the medium
during turnover; (2) kinetic isotope effectszob'c were observed at the C-5 and C-6
positions, indicating that NAD oxidation of C-5 and elimination of phosphate at C-6 were
rate limiting; (3) NAD was indispensable for maintaining stability and activity of the

l.20b,d

enzyme; (4) a trapping experiment20d with tritiated sodium borohydride gave results

consistent with formation of a keto group at the C-5 position.

synthase
phospha:
condens;
0“nehn
indicated.

 

 

 

Dblfllned .

DHstr
Lite 3.31“.
The role
absenct,
did in d,

bill d1 n

(Ir)

Earlier mechanistic studies led to some suspicion to fall on the role that DHQ
synthase played in catalyzing some of the steps in the conversion, especially the

phosphate eliminationziam

and the ring opening followed by intramolecular aldol
condensation.21d Several results suggested that the phosphate monoester catalyzed its
own elimination with the enzyme being a mere spectator. Certain experiments also
indicated that DHQ synthase only served as a template to control the stereochemistry of
the last two steps, but not to catalyze them. It was therefore considered more appropriate

b However, results were also

to regard DHQ synthase as a simple oxidoreductase.21
obtained consistent with theories contradictory to those stated above.22

All controversies were put to rest with the elucidation of the crystal structure of
DHQ synthase from Aspergillus nidulans.23 A close analysis of the interactions between
the active Site residues and the substrate analogue carbaphosphonate, served to resolve
the role that DHQ synthase played in every step of the catalysis. For example, the
absence of basic residues in the vicinity of the C-6 confirmed that the phosphate group
did indeed catalyze its own elimination. However, the enzyme provided a phosphate
binding pocket to orient the phosphate group in a position suitable for abstraction of the
C-6 proton. Interactions between the carboxylate group and active side residues locked
the C-2 conformation so as to prevent epimerization during the final two steps in the

mechanism. Hence, the labeling of DHQ synthase as an oxidoreductase would be quite

erroneous given its active involvement in every step of the transformation.

—...-n- v

suzu. C t.

muteriub

 

o tuined
Besides :
renett ab}
feedstock

Abundant

 

 

current. a‘
Chemical
ill high If
Constituti
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friendly,
lllClUClin:
Cheap an
of Orgar
Slifiams
CC‘DTEnd

D0 Vt" be

Biocatalytic Synthesis of Value-Added Chemicals

The search for cleaner, safer, and cheaper chemical processes is a never ending
saga. Current chemical production protocols rely on abiotic catalysts and on starting
materials derived mostly from petroleum. For instance aromatics are predominantly
obtained from the benzene, toluene, xylene (BTX) fraction of petroleum refining.
Besides the environmental issue, the cost of isolating these components from a non-
renewable feedstock is important to consider. The non-renewability of petroleum
feedstock may not be a concern to the current generation or the many more to follow.
Abundant stocks of petroleum exist to justify our being indifferent to its use at the
current, alarmingly high rate. But we owe it to the future to start probing for alternatives.
Chemical processes typically involve use of toxic, corrosive, and carcinogenic materials
at high temperatures and/ or pressures. By-products typically find no other use and
constitute a waste hazard, the disposal of which has enormous monetary and
environmental ramifications.

The use of biocatalysis has the potential to usher in a new age of environmentally
friendly, industrial scale processes, capable of producing a wide spectrum of chemicals,
including aromatics at a reasonable cost. Biocatalysis relies on the use of renewable,
cheap and non-toxic carbohydrate resources. Reactions are typically run in water instead
of organic solvents at near-ambient temperatures and atmospheric pressures. Waste
streams are generally benign and easily degraded. Biocatalytic processes initially had to
contend with problems of low product yieldS/ titers, paucity in terms of accessible
molecules and inconvenient reaction times and volumes. Most of these problems can

now be addressed with the advent of improved molecular biology and fermentation

-10-

hes the;

Acrylan

-

starting :
Clll'l’EIlll}
flocculur

actilamit

 

Problems
ttgenern
hl'drol}s
SFPfiratit

”311516”

Figure

techniques as well as a better understanding of cellular mechanisms. The remainder of

this section will discuss biocatalytic routes to several industrially relevant chemicals and

how they compare to their chemical counterparts.

Acrylamide

Acrylamide is one the most important commodity chemicals widely used as
starting material in the polymerization industry. Worldwide production of acrylamide
currently stands at 200 x 106 kg.24 Polymers derived from acrylamide find applications as
flocculants, stock additives, and in petroleum recovery. Conventional synthesis of
acrylamide involves hydration of acrylonitrile using copper salts as catalyst (Figure 4).24
Problems associated with this process include difficulty in preparing, recycling, and
regenerating the catalyst. There is also the danger of polymerization or complete
hydrolysis occurring during the reaction because of the heating, which complicates the

separation and purification of acrylamide. Industry has now switched to a biocatalytic

transformation of acrylonitrile to acrylamide.25

2
[05". CU+ , H20, A ’ WNHZ

 

 

O
_ Pseudomonas NH 2
//—C :N + N
chlomraphis 823 O
acrylonitrile acrylamide

Figure 4. Comparison of chemical and biocatalytic routes to acrylamide.

The strain Pseudomonas chlororaphis B23 was isolated from soil as an

isobutyronitrile-utilizing bacterium. Under optimized fermentor conditions at 10 °C

-11-

quantitu:

COI‘Iilil‘IC-T

lactic at

 

and app:
ltutorin;
recent us

of the ne

 

acid. Pg
it could
Lactic 3
am L-is
acid lite

0T aceta

blocata
blo’x‘ate

5)?“

depem

using methacrylate as an inducer, immobilized P. chlororaphis B23 converts acrylonitrile
quantitatively to acrylamide (Figure 4). There is no purification step, the reaction

conditions are very mild and the immobilized cells can be used repeatedly.

Lactic acid

Global lactic acid production is estimated to be more than 100 x 108 kg per year,
and approximately 75% of the lactic acid produced is used in the food industry as a
flavoring acid or baking agent and its glycerol ester is used as an emulsifier.26a More
recent uses for lactic acid have been driven by ecological interests and include production
of the non-chlorinated solvent ethyl lactate as well as the biodegradable plastic, polylactic
acid. Polylactic acid is a polymer whose properties are similar to those of polyolefins and
it could replace a significant portion of the polyethylene terephthalate-based polymers.26
Lactic acid can be synthesized chemically but such synthesis results in a mixture of D-
and L-isomers with only the L-form being commercially useful. The synthesis entails
acid hydrolysis of lactonitrile at 100 °C. Lactonitrile in turn is produced by the treatment
of acetaldehyde with hydrogen cyanide under basic conditions (Figure 5).27

Chemical synthesis of lactic acid has now been replaced in industry by
biocatalytic routes. For example, Rhizopus oryzae is a fungus widely used for
biocatalytic conversion of carbohydrates to L—lactic acid under aerobic conditions (Figure
5).28 The fermentations are very efficient and can result in yields higher that 90%

depending on the starting carbohydrate used.

-12-

fiwmi

global s.
feedstoci
all? print:

being ex.

 

St‘nuhesi

iH HO OH 100 . OH
——* /l~ ——>
CN st04 COzH
acetaldehyde lactonitrile D,L-lactic acid

I'0 H Rhizopus H 0,, H
’ /<C o H
OH oryzae 2

 

Ho OH
D-glucose L-lactic acid

Figure 5. Comparison of chemical and biocatalytic synthesis of L-lactic acid.

Among amino acids, L-lysine commands a huge share of the market with annual
global sales of well over 200 x 108 kg.29 Its primary use is as an additive in animal
feedstock. Aspartame is the highest selling food additive in the world.30a Both molecules
are prime examples of compounds previously produced by chemical processes, but now

being exclusively manufactured via biocatalytic fermentations (Figure 6).29'3°"

L-lysine aspartame

Figure 6. Structures of L-lysine and aspartame.

The sequence of events leading to the crossover from chemical to biocatalytic

synthesis for either of the aforementioned compounds is very engaging in its own right.

But in the context of the subject of this thesis, it would now be pertinent to discuss how

-13-

the shikimate pathway can be manipulated to culminate in the production of industrially
relevant value—added chemicals.

The shikimate pathway has proven to be quite prolific in terms of its applicability
for biosynthesis of aromatic compounds. Low-cost fermentations have been developed
for production of phenylalanine and tryptophan.31 Chemical and enzymatic routes exist

for conversion of the aromatic amino acids to various other value added chemicals

(Figure 7).
002' CH302C o
+
NH3+ NI)J\ENH3
H -
-——)> CC»
—->
L-phenylalanine aspartame
002'
+ o o
NH3

0
—>

 

 

OH 0 O
. . _ n
L-tyrosine melanIn
coz’
H3+ O

H
\ -—>_. O t
N 't O
A H
L-tryptophan indigo
Figure 7. Chemicals accessible from the aromatic amino acids.
0b

Phenylalanine can be chemically or enzymatically converted to aspartame,3

tyrosine can be converted to the mammalian pigment melanin,32 which is widely used in

-14-

hair d):

indigo." '

Indigo

shiiimat
indigo. 1
annual o.
product;
IFi-gure *
leedstou

presents

 

Plgllre 8

hair dyes and in sun screens, while tryptophan can be utilized for the production of

indigo.33

Indigo

The biocatalytic synthesis of indigo provides an interesting example of how the
Shikimate pathway can be interfaced with enzymes from other pathways/organisms.
Indigo, the dye that imparts the unique blue color to jeans, is a widely used vat dye with
annual worldwide production of approximately 1.3 x 107 kg.“ Current routes to indigo
production involve the use of either aniline or anthranilic acid as starting materials
(Figure 8).34 Not only are these compounds derived from non-renewable petroleum
feedstocks, but toxic waste (ammonia, cyanide compounds, formaldehyde, etc.) also
presents serious environmental concerns.

CIVC02H

(1 °' 03»
’

NH2 i) HCHO, HCN w“: OH

aniline ii) Base W

co H /
t: 2 W - “I
r ! A .
NH2 ”AcozH Indoxyl

H

 

 

anthranilic acid

OH
air oxidation
2m
't'
H

indoxyl indigo
Figure 8. Chemical synthesis of indigo.
A biocatalytic route from a renewable feedstock has now been developed for

indigo by overexpressing the tryptophanase and naphthalene dioxygenase enzymes in a

-15-

tryptophan-synthesizing microbe (Figure 9).33 Whether this process will replace current
methods of indigo synthesis remains to be seen, but it highlights the attractive alternatives

that biocatalysis can provide to chemical synthesis.

 

 

002'
\ tryptophanase> ’<I\>
- 'i‘ ”l
HO OH H H
D-glucose L-tryptophan
O
........ __.__. O *’
ne
©j§:OH airoxidation N \ N
dioxygenase |_I| O
O
cis-indole-2,3- . .
dihydrodiol '“d'9°

Figure 9. Biocatalytic synthesis of indigo from D-glucose.

Vanillin

As a food flavor, vanillin is second only to aspartame in terms of annual sales.“
About 12 x 106 kg of vanillin are produced annually throughout the world.35 Synthetic
vanillin can either be obtained from waste sulfite lye in wood pulping operations or from
catechol (Figure 10).}6 Both methods are fraught with huge amounts of toxic waste

streams and use of toxic, carcinogenic materials.

-16-

ligur

 

O

HJHrOH
CH 0 so 0
( 3 )2 2’

 

 

OH OH
catechol guaiacol
HO COZH o C02H o H
—“—>
CH3O —*CH30 CH30
OH OH
mandelic acid 4- -hydrox --3 -methoxy vanillin

phenyl g yoxylic acid

Figure 10. Chemical production of vanillin from catechol.

 

 

COZH 002"!
Mali
0 5 OH OH
OH OH
D-glucose DHS PCA
O H
002H
aryI-aldehyde
COMT dehydrogenase
OCH3 OCH3
SAM AMP, PP; 0H
v:nillic NADPH NAD” vanillin
acid

Figure 11. Biocatalytic synthesis of vanillin from D-glucose.

Recently a synthesis of vanillin from glucose has been elaborated by using a

recombinant E. coli strain (Figure 11).37 The conversion was accomplished by synthesis

-17-

b\ ani-
though

tooards

 

These 1"
also use
reactior
{Figure
Problen
The used

PEUOTe

 

 

of DHS from glucose via the common pathway followed by conversion of DHS to
vanillic acid by the action of aroZ encoded DHS dehydratase and catechol-O-methyl
transferase (COMT). The final step involved in vitro reduction of vanillic acid to vanillin
by aryl-aldehyde dehydrogenase which was isolated from Neurospora crassa. Even
though this process afforded less than 5 g/L of vanillin, it is definitely a giant step

towards large-scale environmentally-benign manufacture of vanillin.

p-hydroxybenzoic acid (PHB).

The use of PHB in liquid crystal polymers such as Zydar is well documented?“
These materials find applications in high performance instruments. Esters of PHB are
also used as food preservatives.38b Currently, PHB is synthesized via the Kolbe-Schmidt
reaction involving treatment of potassium phenoxide with 20 atm dry CO2 at 180-250 °C
(Figure 12). Neutralization of the potassium salt with a mineral acid affords PHB.
Problems associated with this process range from the high temperature and pressures, to
the use of phenol which is a toxic, corrosive chemical obtained from non-renewable
Petroleum feedstock.

An E. coli strain has been developed incorporating the ubicC gene encoding for
Chorismate-pyruvate lyase (Figure 12). This enzyme catalyzes the conversion of
Chorismate directly to PHB.39 The biocatalyst consisted of plasmid-localized aroA, aroL,
aroB, and aroC along with genome-localized tktA, aroFfl’R, and ubicC The carbon flow
into the common pathway was thus optimized leading to increased chorismic acid
production which was subsequently processed by chorismate-pyruvate lyase. Genes

encoding other chorismate-utilizing enzymes were mutationally inactivated. This

-18-

hrusot

Cl

l|
Oh

I
potass
phenc

Flgure

and D.g

 

P'amim

problem
outofi
muhg

TllIrO gr

resulted in production of 12 g/L of PHB from glucose.40 Shikimic acid can also be
converted to PHB by acid catalyzed dehydration.4o This chemical conversion can be very

profitable provided a cheap, reliable source of Shikimic acid can be found, which is the

focus of Chapter 4 in this thesis.

C 02K c 02H
20 atm C02
180- 250 °C
OK OH UbiC COZH
potassium potassium
phenoxide p- hydroxybenzoate o’lL
c02H

l:

COZH fl chorHismic acid

 

Fig. 1
.l
; Ho' 5 OH
HO OH OH
D-glucose SA

Figure 12. Comparison of the chemical and microbial synthesis of PHB from phenol

and D-glucose respectively.

p-aminobenzoic acid (PABA)

The synthesis of PABA is a process that has to deal with health, safety, and waste
problems in every step (Figure 13).“l Nitration of toluene gives a mixture of compounds
out of which p-nitrotoluene is separated out. Chromic or nitric acid oxidation of the

methyl group to a carboxylate functionality followed by iron catalyzed reduction of the

nitro group results in PABA.

-19-

 

DEC-X} ch

synthase
base is
and it \s
iltilgne

a”testis

QUTnic

Sbikini.

inanu

CH3 CH3 COzH

HNOa, H2804) HNanr
30-45 °c H2Cr04
+

 

  
 

N02 N02 F\e’ H C02H
toluene p-nitrotoluene p—nitrobenzoic
acid
CO H
OH 2 PabA COZH Pabc NH2
.I Fig. 1 6&0 JLCOZ PabB (5‘ JL PABA
: OH _ o C02H
HO OH OH IiIH2
D-glucose chorismic acid ADC

Figure 13. Comparison of chemical and microbial catalyzed synthesis of PABA.

Chorismic acid can be converted to (PABA) via initial formation of 4—amino 4-
deoxychorisrnic acid (ADC) followed by elimination of pyruvic acid (Figure 13).42 ADC
synthase is made up of two subunits encoded by the pabA and pabB loci, while ADC
lyase is encoded by pabC. All three genes have been cloned and sequenced from E. coli
and it will be only a matter of time when a PABA producing biocatalyst is successfully
designed. PABA is an ingredient in UV-blocking formulations and its ester is the local

anaesthetic benzocaine.4|

Quinic acid (QA), Hydroquinone, and Benzoquinone

A by-product of the shikimate pathway arising out of reduction of DHQ by
shikimate dehydrogenase is QA,43a which is a very useful molecule and finds applications
in a number of synthetic schemes.“3'*"’2”"22c It is currently isolated from the bark of the

cinchona plant. The Frost group has successfully designed an E. coli biocatalyst capable

-20-

tumult:

Figure

 

benzoqt
product
a phott‘
Sintbe'
bcnzen.
dent at:
results
ElPlosi
maout”.

of bent

else”:

SUpern

Ieml‘er

of producing up to 60 g/L QA (Figure 14).“4 This was achieved by overexpression of

shikimate dehydrogenase and feedback insensitive DAHP synthase in an E. coli strain

 

lacking DHQ dehydratase.
HO," COZH
- Ho" : ‘OH
HO OH OH OH
D-glucose DHQ QA

Figure 14. Microbial synthesis of QA.

The largest potential application of QA is in the production of hydroquinone and
benzoquinone, both of which are currently synthesized from benzene.45 Annual global
production scales for hydroquinone currently stand at 4.0 x 107 kg.46 It is widely used as
a photographic developer. Benzoquinone is used as an organic building block.45c

45” involves zeolite catalyzed Friedel-Craft alkylation of

Synthesis of hydroquinone
benzene to p-diisopropyl benzene (Figure 15). Air oxidation of this dialkylated benzene
derivative under strong caustic conditions at 90-100 °C along with cobalt or copper salts
results in hydroquinone. The air oxidation proceeds via peroxide chemistry, the
explosive nature of which cannot be understated. On the other hand, benzoquinone is
manufactured by oxidation of aniline with MnO2 under acidic conditions.‘5d'° Reduction
of benzoquinone over iron also affords hydroquinone.

QA produced in fermentations can be easily converted to hydroquinone in
excellent yields by the action of industrial bleach (Figure 15).“7 The fermentation culture

supernatant is treated to remove protein and cations, acidified and stirred at room

temperature for 3 h with bleach. The bleach was neutralized with isopropanol and the

-21-

resulting solution refluxed for approximately 8 h giving clean hydroquinone. This
reaction sequence represents a process which can be easily manipulated to meet industrial

requirements in terms of yield, purity and waste.

NH2
Mn02 O
—->
st04 o
aniline benzoquinone

Fe H+

\

HO C02H
bleach
H+

0|"
0""

thdroquinone
GA
02
A NaOH
benzene p-diisopropyl
benzene

Figure 15. Various synthetic routes to hydroquinone.

Dehydroshikimic acid (DHS)

The shikimate pathway intermediate that has found the most number of
applications to date is DHS.48 The antioxidant properties of DHS are well known. In
fact, DHS is as good as if not a better antioxidant than many commercially used
antioxidants49 with the added advantage that DHS can be produced at an industrially
economical scale from glucose. The Frost group has successfully developed an E. coli
strain capable of producing up to 88 g/L DHS in a fed-batch fermentor (Figure 16).50

This strain was constructed by inactivation of genomic aroE encoding for shikimate

-22-

 

Figure

dehydrogenase and overexpression of aroFcBR; tktA, and pps. Perhaps more important
than the properties of DHS itself is its conversion, chemically and biocatalytically, to

industrially relevant chemicals (Figure 16).

H2N N H2
l ‘ COZH
, N
<—— —->
HO OH HO OH
CH30 OCH3 0H 0H
OCH3 gallicacid Pyrogallol

trimethoprim T l
H CO H
002 2 0
rd —» .

o . OH OH *0
H OH

 

 

OIII

 

 

 

bendiocarb
o—gluoose D H S P C A
N H3+ l l
C05 CHO
«— —->
H O H 0 00 H3
0 H O H O H
L-dopa catechol vanIllin
H
H020 W002
adipic acid

0 l H
Wow/Wl.

nylon 6.6

Figure 16. The various synthetic applications of DHS.

-23-

pyrog;

aroma:

 

rdaxu:
flwfii'
“r?” as
encode,
deh§drl
hydm;
anqn;

(”OZ dn

 

acklfro

Fig" re

Gallic acid

Gallic acid is a trihydroxy aromatic compound isolated from gall nuts and tara
powder.’l Thermal decarboxylation of gallic acid in copper autoclaves yields
pyrogallol.51 Both compounds are used as building blocks to provide the trihydroxylated
aromatic ring of biologically active molecules like the antibiotic trimethoprim, the muscle
relaxant gallamine triethiodide and the insecticide bendiocarb (Figure l6).5l Pyrogallol
also finds applications in photographic developing solutions?2 DHS can be chemically as
well as biochemically converted to gallic acid (Figure 17).”53 DHS dehydratase,
encoded by the aroZ locus and isolated from Klebsiella pneumoniae, catalyzes the
dehydration of DHS to protocatechuic acid (PCA). A mutant isozyme of p-
hydroxybenzoate hydroxylase, encoded by pabA *, has been obtained from Pseudomonas
aeruginosa which is capable of hydroxylating PCA to form gallic acid?4 Expression of
aroZ and pobA* in a DHS-synthesizing microbe resulted in production of 20 g/L gallic
acid from glucose?4 DHS can also be chemically transformed to gallic acid via copper

catalyzed oxidation in the presence of zinc (Figure 17).55

 

HO OH OH OH

D-glucose DHS . PCA
CU”. ZEN ;/PobA*
cozH
AroY
HO OH HO OH
OH' OH
gallic acid pyrogallol

Figure 17. Conversion of D-glucose to gallic acid and pyrogallol.

-24-

min
be c
fem
01" P:
acid
[on

with

Cate

 

Gallic acid can be decarboxylated to pyrogallol by aroY-encoded PCA
decarboxylase from K. pneumoniae.56 However, the decarboxylation of gallic acid is
slower than that of PCA. Hence aroY cannot be overexpressed in a gallic acid producing
microbe because PCA occurring as an intermediate between DHS and gallic acid would
be converted to catechol.”56 The problem was resolved by adding gallic acid to a
fermentor containing an E. coli strain overexpressing only aroY, resulting in a 97% yield
of pyrogallol from gallic acid.53 Thus, even though the biocatalytic production of gallic
acid is still not a stage where it can be industrially viable, the outlook is promising.
Conversion of gallic acid to pyrogallol though, is already at a stage where it can compete

with chemical decarboxylation.

Catechol

Catechol is an example of a simple, dihydroxylated, aromatic molecule which
finds applications in a number of industrial chemical processes. The variety of chemical
products derived from catechol is staggering and includes pharmaceuticals (L-dopa,
adrenaline, papavarine), flavors (vanillin, eugenol, isoeugenol), agrochemicals
(carbofuran, propoxur), and polymerization inhibitors and antioxidants (4-tert-
butylcatechol, veratrol).“3’57 Currently, the primary source of catechol is via synthesis
from benzene (Figure 18).”57 Friedel-Crafts alkylation of benzene affords cumene which
upon oxidation leads to formation of phenol and acetone. Subsequent treatment of the
phenol with 70% hydrogen peroxide in the presence of transition metal catalysts leads to
a mixture of catechol and hydroquinone. Catechol is purified away from hydroquinone

by distillation.

-25-

dex'el.‘

(Figur;

 

 

lflipn

(Mu

ingUr

Adip

dEm;

inxg'

As stated before, the enzyme PCA decarboxylase catalyzes the formation of
catechol from PCA at a rapid rate. This implies that in theory catechol can be
synthesized from glucose via DHS. Akin to gallic acid synthesis, an E. coli strain was
developed overexpressing aroZ and aroY to afford catechol via a biosynthetic route

(Figure 18).56

  
 

acetone
[2 _a, ‘2, O hydroquinone
HO <
c
benzene cumene phenol
HO
OH COZH COzH / OH
,.OH Fig.1 catechol
___) AroZ AroY
; OH O 5 OH OH
HO OH OH OH
D-glucose DHS PCA

(a) propylene, solid H3PO4 catalyst, ZOO-260 °C, 400-600 psi. (b) 02, 80-130 °C; 802‘
60-100 °C. (c) 70% H202, EDTA, Fe‘2 or C0”, 70-80 °C.

Figure 18. Comparison of chemical and biocatalytic routes to catechol.

Adipic acid

Adipic acid makes up one of the 6-carbon units in nylon—6,6. Its annual global
demand is about 1.9 x 109 kg.58" Adipic acid is currently synthesized via a route
involving nitric acid oxidation of a cyclohexanol/ cyclohexanone mixture (Figure 19).58b
The by-product of this step is nitrous oxide which has been identified as one of the main
culprits in depletion of the ozone layer and global warming.59 This synthetic route in fact

accounts for 10% of the annual increase in atmospheric nitrous oxide levels. An

-26-

might

catcch

 

Cali-Cf
Anna
lli‘} P.

and l

benze

 

environmentally benign biocatalytic synthesis of adipic acid is really called for under
such a situation.

Adipic acid may at first glance have nothing in common to catechol, and hence it
might be difficult to fathom a biocatalytic route leading from one to the other. But
catechol can be biocatalytically transformed to cis,cis-muconic acid by the action of
catA-encoded catechol 1,2-dioxygenase (Figure 19).56a The catA gene was isolated from
Acinetobacter calcoacet'icus.60 Catalytic hydrogenation of cis,cis-muconic acid using
10% Pt on carbon under 50 psi hydrogen pressure and room temperature results in adipic

acid. This would therefore classify as a synthesis of adipic acid from glucose via DHS.

OH O

0A0$o+

x
benzene cyclohexane CYC'O' CYCIO-

hexanoi hexanone
H020 /\/\/C02H

H ’Pt adipic acid
OH 2/
,.OH F“ .1
_'_g) CatA FE/COZH
- OH HO “020
HO OH OH
D-glucose catechol cis,cis-muconic acid

  
 

(a) Ni-A1,o,, H2, 370-800 psi, 150-250 °C. (b) Co, 0,, 120-140 psi, 150-160 °C. (c) Cu,
NH4VO3, 60% HNO3, 60-80 °C.

Figure 19. Chemical and microbial synthetic routes to adipic acid.

These examples prove the versatility of the shikimate pathway in providing safe,
eco-friendly, alternative routes for chemicals currently made via hazardous processes.

The successful application of genes and enzymes from other pathways and organisms in

-27-

 

induur

Mocaai

 

scde.l
amouni
lmprov
DAHP
Catlin:
Which

genes.
and u»)
achie\

Subfitra

jlilo 1h:

[ranger

conjunction with the E. coli shikimate pathway opens the gateway to an entire gamut of
compounds. As science heads into the 21St century, and with the chemical industry
coming to terms with its role in protecting the environment, the potential for biocatalysis

to make an impact is, dare we say, without any boundaries.

Metabolic Engineering of Escherichia coli for Optimization of

Product Titer and Yield

For any process, be it chemical or biocatalytic, to be successfully applied in
industry, it has to fulfill certain criteria in terms of yield and titer. In order for a
biocatalyst to produce SA in quantities high enough to justify its use on an industrial
scale, it is necessary to improve carbon flow into the common pathway.61 A considerable
amount of research has been expended towards this end. The primary target for
improving carbon flow into any pathway is the first enzyme of the pathway, in this case,
DAHP synthase. The rate of aromatic amino acid biosynthesis is modulated by the
catalytic activity of DAHP synthase. DAHP synthase exists as three isozymes each of
which is sensitive to feedback inhibition by one of the three aromatic amino acids. The
genes aroF, aroG and aroH encode for the tyrosine-sensitive, phenylalanine-sensitive,
and tryptophan-sensitive isozymes respectively. The regulation of the three isozymes is
achieved via feedback inhibition, genetic expression levels and availability of their
substrates E4P and PEP.

Overexpression of DAHP synthase can be realized by introduction of a mutation
into the locus encoding the aporepressor (TyrR) for aroF transcription. In lieu of TyrR,

transcription of aroF is derepressed and the number of molecules of AroF increases.62

-23-

Repla.
gite :5
polyni
Amplf
amF 5
canno:
Fills ls.
DAHP
syntha
technit

lmprm

amtplifi
S}'nthe\
increas;
Several
Chef at
carbgh}.
(PTS) 5

hm‘fl'er

 

Replacing the native promoter (Pomp) of aroF with a strong promoter like a Pm can also
give the same result. The TyrR repressor protein does not influence binding of RNA
polymerase to the Pm promoter resulting in a higher degree of transcription.
Amplification of DAHP synthase activity can also be accomplished by localization of the
aer gene on a multi-copy plasmid. However, the in vivo activity of DAHP synthase
cannot be increased beyond a certain point irrespective of the level of gene amplification.
This is because of the powerful role played by feedback inhibition in the regulation of
DAHP synthase activity.63 Feedback insensitive mutants of all three isozymes of DAHP
synthase have been isolated.°2'“'°5 Application of any of the gene amplification
techniques to a feedback insensitive version of DAHP synthase can now result in
improved in vivo catalytic activity.

Ultimately, DAHP synthase catalytic activity attains a point where further
amplification. of feedback resistant DAHP synthase has no incremental effect on the
synthesis of aromatic amino acids or their precursors. Attempts were then made to
increase availability of the substrates required by DAHP synthase namely, PEP and E4P.
Several cellular processes and enzymes compete with DAHP synthase for available PEP.
Chief among them are pyruvate kinase, PEP carboxylase and the PEP-dependent native
Carbohydrate uptake system, the phosphoenolpyruvate-carbohydrate phosphotransferase
(PTS) system. Mutational inactivation of pyruvate kinase66 and PEP carboxylase6‘c'67
hOwever did not give any significant improvement in aromatic amino biosynthesis.

The focus was then placed on in vivo concentrations of E4P. In 1990, Frost and
CO-workers reported that availability of E4P was an important factor limiting DAHP

Synthase activity when glucose was used as the carbon source.6”'68 Levels of E4P were

-29-

reponc'
for thi
Transl
Condn
high Ct
mainta
hmm
rcquir:
intolv
enzym
Cami};
glycer
to be

sedohr,
catal} ,
incred
With 1
heptui
amplir

meCU

 

SUbSIf

 

reported to be low even in the absence of amplified DAHP synthase activity.69 A reason
for this could be its rapid dimerization, trimerization, and polymerization in solution.70
Transformation of these polymeric forms back to monomeric E4P is quite slow.
Conditions disfavoring the monomeric E4P form include low temperature, basic pH, and
high concentration. By matching the rate of E4P production with E4P utilization, the cell
maintains a low concentration of E4, ensuring its stability as a monomer. This strategy
however meant that steady state concentrations of E4P were insufficient to satisfy
requirements of an amplified DAHP synthase activity level. Analysis of processes
involving E4P as a substrate or a product led to the pentose phosphate pathway and the
enzyme transketolase encoded by the tktA gene (Figure 20). The transketolase enzyme
catalyzed formation of E4P from the coupling of D-fructose 6-phosphate with either D-
glyceraldehyde 3-phosphate or D-ribose S-phosphate. Formation of E4P was also found
to be catalyzed by a second enzyme, transaldolase. This enzyme however utilized D-
sedoheptulose 7-phosphate, which was generated as a by-product during the transketolase
catalyzed formation of E4P. Amplification of transketolase expression levels resulted in
increased channeling of E4P into the common pathway.68 Coupling tktA overexpression
with that of aroGFBR in an E. coli aroB strain led to increased 3-deoxy D-arabino
heptulosonic acid (DAH) accumulation relative to that observed when only aroGFBR was
amplified.68 Accumulation of DAH can be considered as a measure of carbon flow into

the common pathway. Formation of DAH results from dephosphorylation of DAHP, the

substrate of DHQ synthase which is the enzyme mutationally inactivated in E. coli aroB.

-30-

 

 

 

 

 

D—lmr:

“Orkc‘

 

H203PO OH 0 Hzoapo o

M...
. + _ H *—
OH OH OH OH
D-tructose 6-phosphate D-glyceraldedyde
-phosphate

H203PO OH 0 H203PO OH 0
.. K/KJLH __..__-”<'A

OH OH OH OH OH
D-tructose 6-phosphate D-ribose 5-phosphate

H203PO OH OH OH Hzogeo 0
° TaIB

+ H ‘_—_.=

OH OH 0 OH
D-sedoheptulose D-glyceraldedyde
7-phosphate -phosphate

 

 

H203PO OH

H203PO OH

H203PO OH

0H0

 

 

D-erythrose
4-phosphate

H203PO OH OH

OH 0
D-xylulose
5-phosphate

HZO3PO OH OH OH

OH OH 0
D—sedoheptulose
7-phosphate

H203PO OH 0

OH OH OH
D-fructose 6-phosphate

Figure 20. Reactions catalyzed by transketolase (TktA) and transaldolase (TalB).

With the problem of E4P availability no longer being a factor, Liao and co-
workers re-examined the role of PEP availability in increasing aromatic biosynthesis.
The shikimate pathway and glucose uptake compete for the same intracellular levels of
PEP. Glucose uptake in E. coli is mediated by the PTS transport system (Figure 21),
which is responsible for the transport and phosphorylation of a large number of

carbohydrates, movement of cells towards these carbon sources (chemotaxis), and in the

regulation of a number of metabolic pathways.71

-31-

61d.c

 

‘“

syntax;

 

lntetm
erased.
of all
encod
26 ll(

Pig.

glucose 6- -phosphlate

®—a ®-—HPr ®-EB
PEPXB XHP rX®—l IA“XEBo g'ucose
pyruvate llAdc

cytoplasm lperiplasm
Figure 21. The PTS-system for glucose uptake.
Genetic loci are as follows: ptsI, Enzyme I (EI); ptsH, Histidine protein (HPr); crr,
glucose specific Enzyme IIA (IIAg'°); ptsG, Enzyme B and C (EB and C).

The phosphate group is transferred to the glucose via obligatory phospho-
interrnediates of proteins EI, HPr, IIAg'c, EB and C. The ptsI-encoded EI and ptsH-
encoded HPr proteins are soluble cytoplasmic proteins that participate in phosphorylation
of all PTS carbohydrates. Protein IIAglc is a soluble enzyme specific for glucose and is
encoded by the crr locus. The two membrane bound proteins EB and C, also designated
as IICB‘“, comprise the ptsG-encoded II81c domain which is specific for glucose. The
PTS-system relies on phosphoryl group transfer from PEP to initiate the cascade of
events leading to glucose uptake (Figure 21).71 The pyruvic acid by product is then used
for sustaining cell growth. As a result three carbon atoms are lost for every six carbons
imported by the cell (Figure 22). One strategy to nullify the competition between the
shikimate pathway and the PT S-system for glucose, is to overexpress PEP synthase. The
enzyme PEP synthase, encoded by the pps locus, catalyzes the phosphorylation of
pyruvate in the presence of ATP, resulting in PEP, AMP, and inorganic phosphate

(Figure 23).

-32-

 

Figurl

C0n\ex

paths;

 

benefi
Carbor.

A

OPOst j
COZH COZH -—> CO2

    

PEP pyruvate
\ g
, OH PtS ;
HO OH H203PO OH
D-glucose D-glucose 6-phosphate

periplasm cytoplasm

 

Figure 22. Phosphorylation of D-glucose during its uptake.

Overexpression of PEP synthase will hence enable PT S-generated pyruvate to be
converted back to PEP. However, no improvement in carbon flow into the shikimate
pathway was detected in E. coli aroB with amplified expression of pps and aroGFBR. The
benefits of enhanced PEP availability was realized only when pps and aroGFBR
overexpression was coupled to that of tktA. The result was a twofold improvement in
carbon flow as measured by the yield of DAH produced by E. coli aroB.6""° These
observations indicate that E4P is the first limiting metabolite in aromatic amino acid
biosynthesis. Once this limitation is removed, the full benefits of increased PEP
availability are realized. Amplified expression of transaldolase also relieves E4P
limitation in the presence of amplified PEP synthase, but no further improvements in
aromatic amino acid biosynthesis are observed relative to when transketolase is

overexpressed.72

-33-

 

Figul

 

9 9 9 <9.
Ad—Rib—O-P-O-P-O-P-O— Ad—Rib—O —P-o"‘

O_ O_ O— O
ATP AMP
+ PEP +

_ synthase
co. ———>._ _ JL _
fir oapo co2
O

pyruvate PEP
+ +
HOH O

HO-P-O—
O—

inorganic

phosphate

Figure 23. Reaction catalyzed by PEP synthase.

Alleviation of PEP deficiency by overexpression of PEP synthase can be
problematic. PEP synthase is reported to be a heavily regulated enzyme and its amplified
expression is known to be detrimental to the health of the microbe.61d A better alternative
is to channel the glucose into the cell via a mechanism which is not dependent on PEP.
An uptake system fulfilling this criteria is facilitated diffusion (Figure 24). Zymomonas
mobilis is known to transport glucose by a facilitated diffusion process which is a low-
affinity, high-velocity, non-energy dependent system.73 A large amount of interest in this
area has resulted in the cloning and sequencing of a putative glucose transporter gene
(glf).74 The deduced amino acid sequence of the glf gene product is closely related to a
large family of glucose transporters.” Most importantly, expression of the Z. mobilis glf
and glucose kinase (glk) genes was able to provide for glucose transport capability in E.

coli strains lacking a native PT S system.73W

-34-

 

4‘
pics

 

Fig

lip!

‘* e
an
Co
Ur

Ct

  

OH
.OH
carbohydrate
phosphotransferase
system 5 OH
HO OH
D-glucose
facilitated
glucose
diffusion

 

A

OP03H2
COZH

PEP

o
)LCOZH ——> co2
pyruvate

OH

  

 

PtS ; OH

g H203PO OH

i D-glucose 6-phosphate

5 ADP

E glk

ATP

5 ,.OH

——>

9" OH

perlplasm E cytoplasm

Figure 24. Comparison of the PTS and facilitated diffusion systems for glucose

uptake.

The initial results concerning DAHP synthase, transketolase and PEP synthase

were obtained under shake-flask conditions. Evaluation of the impact of aroE"BR and tktA

amplification have now been performed and confirmed under fed-batch fermentor

conditions.“76 Results reported in this thesis for SA synthesis have all been obtained

under fed-batch fermentor conditions.

This is an important consideration because

contrary to results from fed-batch fermentations, calculated yields from shake-flask

-35-

 

cultix

 

imp):
glues

COD-CC

 

 

 

cultivations do not really reflect the percent conversion of glucose to product. It is also
important to consider that under shake-flask conditions, it is not easy to maintain constant

glucose, oxygen, and pH levels which is not the case where fed-batch fermentations are

concerned.

-36-

bend.
asni
reduc

pnth

 

 

 

S}nth
tonsi
thetl
nfle

inns
lube
13Pl

but

of
di\

“i

CHAPTER 2

INHIBITION OF DHQ SYNTHASE VIA UTILIZATION
OF METAL-DIOL INTERACTIONS

Background

Inhibition studies on DHQ synthase have made significant contributions towards
better understanding the intricacies of this enzyme. It was considered inconceivable that
a single enzyme could catalyze an alcohol oxidation, phosphate elimination, carbonyl
reduction, and an intramolecular aldol condensation in a single turnover of substrate to
product. Initial results did support the mechanism outlined in Figure 3 resulting in DHQ
synthase being considered as a mechanistic marvel. However, conclusions from a
considerable body of work performed in the 1980’s and early 1990’s seemed to debunk
the theory of DHQ synthase being an extraordinarily diverse and efficient catalyst.21 The
role of DHQ synthase in catalyzing the elimination of the phosphate and the
intramolecular aldol condensation came into scrutiny, and some studies even hinted at
labeling it as an oxidoreductase. The recently solved crystal structure of the active site of
DHQ synthase finally provided evidence that the enzyme was not a mere oxidoreductant,
but did indeed play a catalytic role in the conversion of DAHP to DHQ.23

The main focus of the work discussed in this chapter deals with studying the role
of the metal co-factor. DHQ synthase has been proven to contain one tightly bound
divalent metal cation per monomer by the following observations:20a (i) upon treatment
with EDTA, the enzyme loses activity and gives rise to a stable but non-functional

apoenzyme; (ii) reconstitution of the apoenzyme with a divalent metal cation restored

-37-

acn\
came
the 7
FLAIF
has h'
repor
fornz
the a
synth
halfli

thatli

 

syster
”alum
ilier

mm

activity back to the enzyme, the level of activity restored depending on the identity of the
cation; (iii) EDTA does not cause inactivation in the presence of DAHP, indicating that
the metal is less accessible when substrate is bound to the active site; (iii) the rate of
N AD” dissociation varies with the metal ion. Whether the native metal is a Zn+2 or a Co+2
has been debated in the past. The Co+2 form of DHQ synthase from E. coli has been
reported to be significantly more stable and active than its Zn+2 counterpart.“ The Zn+2
form of DHQ synthase has in the past been prepared from the Co+2 form by first stripping
the active site of its Co+2 and then reconstituting the apoenzyme with Zn”. DHQ
synthase retained some catalytic activity upon reconstitution with Zn+2 but had a short
half life of approximately 30 min, making kinetic studies difficult. This led to the notion
that DHQ synthase existed as a C0"2 enzyme in nature.20a

This however did not seem very likely given the availability of Zn+2 in biological
systems.77 It is second in abundance only to iron among the transition metals found in
nature and has been found to be an integral component of more than 300 enzymes in
different species.78 There are several reasons why Zn+2 makes an ideal candidate for a
metal co-factor, especially for DHQ synthase.79 It has a completely filled d- shell with 10
electrons resulting in no ligand field stabilization energy when coordinated by ligands in
any geometry. Zinc is inert to oxidoreduction especially in the divalent state and does not
generate reactive radicals. The lack of redox activity allows Zn+2 to remain stable in
biological systems whose potential is in a constant flux. Properties like its flexible
coordination geometry, fast ligand exchange and intermediate hard-soft character makes
it suitable for use as a co-factor. The multiplicity of coordination numbers and

geometries as well as stereochemical adaptability denotes that Zn+2 submits readily to the

-38-

 

dem.

 

3 Vii":

hthe

beyor
lennh

ln101i

 

hrce I
group
the 3‘
enz§r

facilia

“Natl
lhe (

{hits

demands of its ligands. The ability to exchange ligands at a fast rate is essential for an
active site co-factor to be able to bind and activate substrate followed by rapid release of
product. The Lewis acidity of Zn+2 enables it to polarize carbonyl groups and promote
ionization of water molecules at physiological pH. It is known to interact very well with
a variety of amino acid residues including histidine, glutamate, aspartate and cysteine. It
is the combination of these factors that makes Zn“2 so attractive.

Elucidation of the crystal structure23 of DHQ synthase from A. nidulans proved
beyond doubt that the enzyme existed as a Zn+2 enzyme in vivo (Figure 25). The C-
tenninal domain of the enzyme contains the Zn+2 binding site and many of the residues
involved in catalysis and substrate binding. The penta-coordinate Zn+2 ion interacts with
three ligands from the protein, Glu'”, His271 and Hism as well as the C-4, C-5 hydroxyl
groups of the substrate analogue inhibitor carbaphosphonate. The positioning of Zn+2 in
the active site allows speculation about the role that it might be playing during the
enzyme catalysis (Figure 26). For example, given its polarizable nature, Zn+2 can
facilitate oxidation of C-5 hydroxy group in DAHP and reduction of the C-5 carbonyl
group in intermediate B. At the same time, it serves to immobilize the C—4, C-5 edge of
intermediate D during the intramolecular aldol condensation. This chelation will allow
rotation in intermediate E only about the C-5, C-6 carbon-carbon bond but immobilizes
the C-4, C-5 bond. The stereochemistry of the intramolecular aldol condensation may

thus be controlled.

-39-

 

   
   
 

Asn162
. "I , ”
«’0, HIS 275

O
()Wat33 \‘
Lys 356 1

K

.. His 271
Asn 268

   
     
    

Figure 25. Active site of DHQ synthase.

-40-

'0

p
.‘q
t

 

ligtl

S}'n

C00

VET

Sin

[hf

otl

\hi

Zr

bt

 

h H I?
/ \ ninz - N-Rz (\— N—Rz
— H _ H _
CONH2 CONH2 2+ CONH2 2
H - - ' ‘ 3 :2" B -;;Zn +
@- _ 7
2 H\OIH Wat?
h%"_--Asn162 H
DAHP: H \[Ol’ \ ‘ ‘ Lys 358 Hf‘A’N }
2 k/‘Afl ' a 07-. ‘s
R =30PO '=X O ’ N— Asnzea’.’ - H~ H
curfnphosfhonato: _ ,’ ; Arg 130(8) 0’
=P03 . x= CH2 Wat 33 " Lys 152 His 275
NAD: His 275
R2 = n'bose-P-P-Ad
’ ‘ D " ’znZ-o
@ Lys 250 Lys 250 Ho'” 6) HO
,,.:-‘0\ OH _, OH
A79 264‘: ;;C —__.A|’g 264‘:\ &C \L\ -02
- ‘. H O
Ly8152 ”(V Lys 152' < x H ,‘Q 0
Asn 268 H 3 3-dohydroqulnate
{N (o (‘6 ‘
H’ -H (B\-H ‘H Asn 268 Lys 152
Wat20

Figure 26. Interactions postulated to occur at the active site of DHQ synthase.

Rationale Behind Inhibitor Desi n

 

This study attempts to investigate the inhibition of DHQ synthase based on simple
cis-vicinal diol complexation to a metal co-factor. Past inhibition studies with DHQ
synthase have always been performed with the Co+2 enzyme due to practical
considerations. The Frost group has now successfully purified E. coli DHQ synthase in a
very stable Zn+2 form by simply replacing Co+2 in the purification buffers with Zn”. The
similarity of kcat and K“1 for the Co+2 and Zn+2 enzymes suggests that no major changes in
the nature of mechanism of catalysis occur when one metal is substituted with the
other.20a However, that doesnt mean that subtle differences in active site interactions for
the two metals do not exist. It has been proven that NAD+ dissociates slower from the
Zn+2 enzyme by a factor of about 10. Also, the presence of excess Zn+2 in the assay

buffer leads to decreased rates showing mixed-type inhibition kinetics. This seems to

-4]-

 

sure

the;

rok'

 

form

tony
Dth
suhst
Znei
[hand

“as 1

 

thetal
succe
active
enzyr
COR};
defig
inclu

Si‘ml

suggest that the Zn+2 may be binding to a second lower affinity inhibitory site. The same
effect is not seen with Co”.20a It would therefore be quite interesting to investigate the
role that metal co-factors played in binding of inhibitors to the enzyme and whether one
form of the enzyme was more susceptible to inhibition over the other.

While designing potential inhibitors of DHQ synthase, two factors were taken into
consideration; the crystal structure of DHQ synthase and the structures of the most potent
DHQ synthase inhibitors. The crystal structure of DHQ synthase was resolved using the
substrate analogue inhibitor carbaphosphonate. The crystal structure indicated that the
Zn+2 is tightly nestled with the C-4 and C-5 hydroxyl groups of carbaphosphonate with a
metal to oxygen distance of 2.3 A and 2.2 A respectively (Figure 25). This interaction
was utilized in the designing of new DHQ synthase inhibitors. Inhibition of Zn+2
metalloenzymes by utilization of functionalities capable of Zn+2 complexation has been
successfully achieved. For example, captopril, with its thiol group complexing to the
active site Zn+2 in a monodentate fashion is a potent inhibitor of angiotensin converting
enzyme (ACE).80 Substrate based alcohol inhibitors of ACE such as I employ bidentate
complexation of the hydroxyl and the amino group to the Zn+2 (Figure 27).81 Of course,
design of these inhibitors also takes into account other active site interactions but
inclusion of Zn+2 complexation is of equal importance. Hence inhibition of DHQ

synthase via inhibitors designed to bind to the metal co-factor is a viable option.

Ph
HS N lfl
JY ; BzNH - NJY”;
o COZH HO H o COZH
captopril 1

Figure 27. Inhibitors functioning via complexation of Zn".

-42-

 

 

ngl

 

Udnsi
The l

perh:

Inhibition of DHQ synthase has been achieved using various substrate and
transition state analogues. Cyclohexylidene phosphonate 2 and cyclohexenyl phosphate
3 are the most potent inhibitors of E. coli DHQ synthase with Ki values of 2.9 x 10'10 M

and 1.2 x 10"0 M respectively (Figure 28).22c

Hog COZH HO,’ COZH
(HO)2(O)P\¢©\OH (HO)2(O)P0\/©\OH
OH OH
2 3

Figure 28. Unsaturated inhibitors of DHQ synthase.

These inhibitors were designed to mimic an Elcb intermediate or Elcb—like
transition state during the transformation of intermediate A to intermediate B (Scheme 1).
The results obtained indicated that the unsaturation at 06 was not only tolerated but
perhaps also contributed to the overall binding.

Based on the crystal structure of DHQ synthase and the inhibitory capabilities of
carbocyclic inhibitors 2 and 3, it was decided that any new inhibitor of DHQ synthase
would have to incorporate a cis-vicinal diol at C-4 and C-5 and an sp2 center at C-6.
Protocatechuic acid (PCA) 7 was chosen as a suitable starting template leading towards
inhibitors 4 and 5 (Figure 29). The substituted PCA derivatives 4 and 5 fulfill the
requirements stated above even though they constitute a structural class radically

different from that of the enzyme substrate/transition states (Figure 29).

-43-

 

subs!
Den‘
of th
and .
has I
phos
ISOSI
isost

and

COOH COOH

OH OH
(“0’40” 0“ (Homow 0”
4 5
OOH
H H
(Homow 0 ° 0”
7 a

Figure 29. Aromatic compounds designed for inhibition of DHQ synthase.

The cis-vicinal diol complexation of the active site metal co-factor in carbocyclic
substrate analogues is now replaced by the ortho dihydroxyl complexation of a catechol.
Derivatives 4 and 5 still provide a trianionic ionization state and the relative orientation
of the phosphonomethyl and phosphonoethyl groups relative to the backbone hydroxyls
and carboxylate functionalities is maintained. Maintaining a trianionic ionization state
has been proven to be important for binding to the active site of DHQ synthase.82 The
phosphonomethyl group in 4 is isosteric and the phosphonoethyl group in 5 is non-
isosteric to the phosphate functionality of substrate DAHP. Inhibitors with a non-
isosteric phosphate monoester are often but not always better inhibitors of DHQ synthase

and hence, both 4 and 5 will be examined.2“'22"'32’83

Compound 6 was designed to
determine the contribution of the carboxylate to the overall binding. Its binding will rely
on active site interactions of the phosphonoethyl group and the metal binding of the ortho

dihydroxyl groups. PCA 7 on the other hand will derive its binding affinity via active

site interactions to the carboxylate and metal interactions with the ortho dihydroxyl

 

lun

 

 

 

 

ulth
Ran
folh

Pho

Upg
lnie

functionality. Differences in the inhibition of DHQ synthase by catechol
homophosphonate 6 and PCA 7 will provide a good measure of the importance of the
carboxylate relative to the phosphate for active site binding. Finally, catechol 8 will test
the efficacy of utilizing binding of metal to the ortho dihydroxyl group as the sole means

of achieving inhibition.

Synthesis of Phosphonate 4 and Homophosphonates 5-6

The synthesis of the PCA analogues 4 and 5 exploited commercially available
3,4-dimethoxy benzoic acid 9 (Figure 30). Fischer esterification of 9 followed by
bromination of the resulting ester 10 gave 5-bromo-3,4-dimethoxy methylbenzoate 11.84
The substitution of the bromide with a vinylic group was achieved via a Stille coupling to
give 12. Subsequent ozonolysis of the olefin 12 resulted in 3-formyl-4,5-dimethoxy
methylbenzoate 13. Reductive iodination85 of 13 followed by treatment of the iodide 14
with triisopropyl phosphite under reflux afforded the phosphonate intermediate 15.
Removal of the methoxy groups by treatment with an excess of boron tribromide
followed by acidic removal of the carboxylate and phosphonate esters furnished PCA
phosphonate 4. The reaction of 13 with the lithium anion of tetraisopropyl
methylenediphosphonate yielded the olefinic homophosphonate intermediate 16, which
upon hydrogenation under 10% Pd on carbon cleanly produced the homophosphonate
intermediate 17 . Removal of the protecting groups in 17 was performed in exactly the

same manner as for PCA phosphonate 4, to afford PCA homophosphonate 5.

-45-

 

(7-

 

Inter
redu

mbf:

COOH 002C113 COch3 OZCHS

Ja—Q» ——-9—> —9—>
OCH, Br OCH, l OCH, H OCH,
OCH,
12

OCH3 OC H3 0 001.13
9 11 13
O,CH, O,CH, O,H
__.i.__> ___g_*
e OCH, OCH, OH
cogng 1 OCH, among OCH, (HObfi OH
O
14 ° 15 4
H
OCH,
0 OCH,
13 N CO,CH, CO,CH, COOH
i , l i
I OCH, OCH, OH
, OCH _ OCH OH
(Prong 3 (uproar, 3 (Hoar,
16 O 17 o 5

Key: (a) MeOH, H2804, 98%; (b) Brz, AcONa, AcOH, 70 °C, 97%; (c)
Bu,Sn(CH=CH,), Pd(PPh3),, HMPA, 65 °C, 63%; (d) 0,, CHzClz, -78 °C, 91%; (e)
[(CH,)2HSi]zO, TMSCl, NaI, ACCN, reflux; (f) P(OiPr),, toluene, reflux, 23%; (g) (i)
BBr3, CH2C12, -78 °C to rt; (ii) conc. HCl, 62%; (h) n-BuLi, [(iPrO)2P(O)],CH2, THF, -78
°C to rt, 34%; (i) H2, 10% Pd/C, MeOH, 76%; (j) (i) BBr3, CHZCIZ, -78 °C to rt; (ii) conc.
HCl, 88%.

Figure 30. Synthesis of PCA phosphonate 4 and PCA homophosphonate 5.

Catechol homophosphonate 6 was synthesized using commercially available 2,3-
dimcthoxy benzaldehyde 18 (Figure 31). Wadsworth-Emmons reaction of 18 using
tetraisopropyl methylenediphosphonate yielded the required catechol homophosphonate
intermediate 19, which upon hydrogenation using catalytic 10% Pd on C resulted in
reduced homophosphonate intermediate 20. Subsequent deprotection with excess boron

tribromide and hydrochloric acid afforded the catechol homophosphonate 6.

-46-

K31

7:1“,

Figq

deh‘;
conx
Spec:
quanl

Co‘:

 

Uhs u
Dhfl}
COmp
comp
ShOu‘

Elen

a b C
——> —-> —>

OCH, | OCH, OCH, OH

0 (DCFI (OCH H
3 (Prong ? «Prong OC 3 (Hoar: 0”

18 0 19 0 20 0 6

Key: (a) n—BuLi, [(iPrO),P(O)],CH,, THF, -78 °C to rt, 95%; (b) H2, 10% Pd/C, MeOH,
75%; (c) (i) BBr3, CHzClz, -78 °C to rt; (ii) conc. HCl, 97%.

Figure 31. Synthesis of catechol homophosphonate 6.

Inhibition Studies of 4-8 with DH s nthase

 

Inhibition of DHQ synthase was studied using a coupled enzyme assay with DHQ
dehydratase being used as the coupling enzyme.36 DHQ dehydratase catalyzes the
conversion of DHQ to 3-dehydroshikimate (DHS), the formation of which can be
spectrophotometrically detected at 234 nM, thus providing a means for continuous
quantitation of DHQ formation.8 The Michaelis constant (Km) for DAHP binding to the
Co+2 form of DHQ synthase has been reported to depend on the assay methodzo“ and for
this work has been taken as 18 11M. The Km for the binding of DAHP to the Zn+2 form of
DHQ synthase was measured to be 16 pM. Inhibition constants (K,) were measured for
compounds 4-8 and are reported in Table 1. Compounds 4-7 exhibited classical
competitive inhibition of both forms of DHQ synthase. Surprisingly, catechol 8 also
showed some level of inhibition although its Ki value could be estimated only from an 150

determination.

-47-

"1c;

 

Ta

syl

 

 

V1215

rep].

F1.

Table 1. Inhibition constants (nM) for binding to the Co+2 and Zn+2 forms of DHQ

 

 

 

synthase.
Enzyme Inhibitor
4 5 6 7 8
DHQ synthase-00+2 21 5 200 44 880
DHQ synthase-Zn’r2 0.35 1.7 90 65 630

 

Ki values were obtained from the Dixon plot of the reciprocal initial velocities
versus inhibitor concentrations at fixed substrate concentrations (Figure 32). The Dixon

replot gave a line passing through the origin indicating competitive inhibition.87

Dixon data

 

0.7

0.54 '

0.38

1IV

0.22 ”

0.06

 

 

 

 

~50 -6.25 37.5 81 .25 125
[41 11M

Figure 32. Dixon plot for PCA phosphonate 4.

-43-

 

 

C081

1th

DA}

the}

 

 

suh’

\xas

lrer

dis

sin

tel

Lineweaver-Burke and Hanes-Woolf plots also gave results consistent with
competitive inhibition. I50 determinations were done by plotting the reciprocal initial
velocity versus inhibitor concentrations at 18 11M DAHP for the Co+2 enzyme or 16 pM
DAHP for the Zn+2 enzyme. The x-intercept gave the value for the 150, half of which was

the K,.

Interpretation of the Inhibition Pattern Exhibited by Aromatic

Inhibitors 4-8

The first piece of information that could be gleaned from these results was that the
Zn"2 form of the enzyme was more susceptible to inhibition than the Co+2 form except in
the case of PCA 7. The results obtained with inhibitors 4 and 5 also highlighted the
subtle differences between the two forms of lDHQ synthase. PCA homophosphonate 5
was a better inhibitor of the Co+2 enzyme than PCA phosphonate 4, while the reverse
trend was observed in case of the Zn+2 enzyme. The Zn"2 enzyme evidently did not
discriminate between a carboxylate and a homophosphonate group as indicated by the
similar reduction in enzyme inhibition for catechol homophosphonate 6 and PCA 7
relative to PCA homophosphonate 5. The Co+2 form of the enzyme however exhibited a
bias between binding to a carboxylate or a homophosphonate functionality based on the
different extents to which enzyme inhibition was reduced for catechol homophosphonate
6 and PCA 7 relative to PCA homophosphonate 5. These observations indicate the slight
variations that exist in the binding interactions between the two enzymes. The inhibition

displayed by catechol 8 does indicate the minimal level of inhibition that can be obtained

-49-

11h

3H

IOt

Tilt

5112

a”

3C .'

an.

 

Sir
um

fill

when all other functionalities are missing on an inhibitor and the only interactions
available are those between the C4-C5 diol and the metal co-factor.

This is the first inhibition study performed with DHQ synthase utilizing a Zn+2 co-
factor as opposed to a CO+2 co-factor. DHQ synthase has also been reported88 to be
functional with metals like Ni”, Cd”, Eu+3 and Sm+3 and this study makes it worthwhile
to consider how various metal co-factors might influence binding of substrate/ inhibitors.
The activity observed with the PCA analogues is quite important considering their novel
structural simplicity, mode of inhibition and ease of synthesis. Results discussed with the
aromatic inhibitors of DHQ synthase assume significance because of the ability to
account for the accumulation of DAH as a by-product in the microbial synthesis of gallic
acid.” This was true even when aroE-encoded DHQ synthase was overexpressed in the
gallic acid synthesizing recombinant E. coli. As was illustrated in Figure 17, PCA occurs
as an intermediate in the microbe catalyzed conversion of glucose to gallic acid.
Apparently, the PCA is inhibiting DHQ synthase resulting in accumulation of DAHP in
the cytosol. Dephosphorylation of DAHP results in its export from the cell as DAH,
which appears as the contaminant in the fermentation broth. PCA also occurs as an

15‘5 and cis,cis-muconic

intermediate in the biocatalytic synthesis of vanillin,37 catecho
acid?“ Its inhibition of DHQ synthase therefore constitutes a major impediment in the
elaboration of high yielding microbial synthesis of these compounds.

In conclusion, PCA phosphonate 4 and PCA homophosphonate 5 were
synthesized and successfully tested as inhibitors of DHQ synthase. These compounds are

unique given their structural characteristics as well as mode of action and they represent a

fundamental departure from previously reported carbocyclic inhibitors. But perhaps their

-50-

 

Dl

 

most important feature is their ability to discriminate between the Co“2 and Zn+2 forms of
DHQ synthase. These compounds can act as valuable templates on which future and

hopefully better inhibitors of DHQ synthase can be based upon.

-51-

1117;.

 

363}

The

CHAPTER 3

BIOCATALYTIC SYNTHESIS OF SHIKIMIC ACID
USING RECOMBINANT ESCHERICHIA COLI

Introduction

Shikimic acid (SA) is a seven-carbon carbocyclic intermediate occurring as an
intermediate in the common pathway for aromatic amino acid biosynthesis.89 The

stereochemical disposition of tunable functional groups in SA makes it an attractive

 

chiral synthon for use in various synthetic schemes.90 It is isolated from the fruits of the
Illicium plants at a cost of approximately $10,000/kg,91 precluding its application in
moderate to large scale synthetic processes. For example, SA has been reported to be a
very convenient starting material in the synthesis of the neuraminidase inhibitor GS4104
(Figure 33).92

00,51

:>-—O" ’ 'N H,.H,PO4

NHAc
68-41014

Figure 33. Neuraminidase inhibitor GS4104

This compound is currently being marketed by Hoffman-LaRoche as its new
influenza drug, TamifluTM. Use of SA as the starting material of choice for industrial
scale production of GS4104 will depend on identifying a Cheap and reliable source for it.

The common pathway of E. coli has been successfully manipulated for the accumulation

-52.

C37

act

by

 

pht
exp
“it
of I
am

Phi

of various intermediates and secondary metabolites. Interfacing the shikimate pathway
with enzymes alien to the pathway or to the microbe itself have also met with success. A
recombinant E. coli biocatalyst can therefore be designed, capable of producing large
amounts of SA at a low cost, making the use of SA in industrial scale synthesis
economically viable.

The key to achieving high titers and yields of SA is dependent on improving
carbon flow into the shikimate pathway. Expression levels of DAHP synthase have
proven to be critical for increasing aromatic amino acid biosynthesis. As the first enzyme
of the pathway, DAHP synthase dictates the amount of cellular carbon being directed into
the common pathway and hence towards shikimate synthesis. Regulation of the in vivo
activity of this enzyme is achieved via transcriptional repression and feedback inhibition.
Removal of these impediments have helped in achieving respectable DAHP synthase
activity levels. However, in vivo DAHP synthase activity is also known to be influenced
by intracellular concentrations of the substrates D-erythrose 4-phosphate (E4P) and
phosphoenolpyruvate (PEP). Availability of E4P has been reported to hinge on
expression levels of the enzyme transketolase. The enzyme transaldolase is also involved
with E4P biosynthesis, but its overexpression does not offer any advantage relative to that

72 On the other hand, PEP concentration can be directly improved by

of transketolase.
amplification of the enzyme PEP synthase. Also, it is known that PEP is involved in the
phosphoenolpyruvate-carbohydrate phosphotransferase (PTS) system of glucose transport

in E. coli. Hence, introduction of a non-PEP dependent glucose uptake system into the

biocatalyst can also indirectly alleviate PEP limitation.

-53-

 

 

mil

 

pm

CDT

Qp‘

Ex

In this Chapter, the synthesis of SA has been investigated utilizing various E. coli
K-12 (RB791) biocatalysts under fed-batch fermentor conditions. The effects of
increased transketolase, transaldolase and PEP synthase expression levels on SA
production were studied. Synthesis of SA was also examined in an E. coli strain lacking
a native FT S glucose transport system. Transaldolase amplification gave no increment in
terms of SA yield or titer. The effect of transketolase overexpression was encouraging by
itself but was quite dramatic when coupled with that of PEP synthase. Use of a non-PEP
utilizing glucose facilitator uptake system also gave significant improvements in SA
production. Titers and yields for SA production by all the biocatalysts are reported and
compared.

Synthesis of SA was also examined in a biocatalyst derived from E. coli B as
opposed to RB791. E. coli B lacks a number of proteases, which could result in higher
DAHP synthase and transketolase activity levels, especially in the stationary phase.93
Examination of these strains under fed-batch fermentor conditions indicated enhanced
DAHP synthase and transketolase activity levels as compared to the corresponding

RB791 version. However, no improvement in SA production was observed.

Biocatalysts and Fed-Batch Fermentation Conditions

A) Shared Genomic Traits and Plasmid Elements

Three constructs, SP1.1, SP1.1pts and EBl.1, were used as the hosts for SA
synthesis. The SP1.1 host strain was derived from RB791 and lacks catalytically active
shikimate kinase resulting in SA accumulation. Construction of SP1.1 began with the

homologous recombination of the aroB gene into the serA locus of E. coli RB791

-54-

resulting in RB791 serA::aroB.48'7° RB791 serA::aroB was then subjected to two
successive P1 phage-mediated transductions to transfer the aroL478zzTn10 and
aroK::CmR loci of ALO807 onto the genome and eliminating shikimate kinase activity.”
SP1.1pts was derived from SP1.1 and lacks a native PT S system for glucose uptake. This
was achieved by P1 phage-mediated transduction of A(ptsHptsIcrr)::KanR thereby
disrupting the ptsH, pm] and crr loci involved with E. coli glucose uptake (Figure 21).
Pl phage was propagated from TP2811, which was generously provided by Professor
Sophie Levy.94 EB1.1 was constructed from E. coli B and incorporates the same
deletions as in SP1.1. All strains lack the capability for de novo aromatic amino acid and
aromatic vitamin biosynthesis, necessitating their growth in medium supplemented with
L-phenylalanine, L-tryptophan, L-tyrosine, p-hydroxybenzoic acid, p-aminobenzoic acid,
and 2, 3-dihydroxybenzoic.

DAHP synthase is known to be regulated by feedback inhibition by aromatic
amino acids. DAHP synthase catalyzes the condensation of E4P to PEP resulting in
DAHP, the first committed intermediate in the aromatic amino biosynthetic pathway.
Elevated expression levels of DAHP synthase are therefore critical for high SA yields.
All SA producing strains hence utilize a plasmid localized mutant isozyme of DAHP
synthase, designated as aroFFBR, which is feedback insensitive to the aromatic amino
acids.”76 However, increased activity levels of DAHP synthase meant that aroB encoded
native DHQ synthase levels were now inadequate to convert the higher levels of DAHP
to DHQ at a rate fast enough to avoid DAHP accumulation.” This problem was
circumvented by site-specific insertion of an additional copy of aroB into the genomic

serA locus of the E. coli strain via homologous recombination. Temperature sensitive

-55-

plasmid pKL3.82A was used for the homologous recombination.48 The procedure used
for the recombination is described in Chapter 5 of this thesis. The serA locus encodes for
3-phosphoglycerate dehydrogenase, a key enzyme required for L-serine biosynthesis.96
This strategy hence required localization of the serA gene on all plasmids, which also
provided a convenient method for plasmid maintenance during fed-batch fermentor
cultivation. Shikimate dehydrogenase encoded by the aroE gene is known to be feedback
inhibited by SA resulting in incomplete conversion of DHS to SA.95a Overexpression of
aroE is therefore necessary for improved SA yields and low DHS accumulation. This
was achieved by plasmid localization of the aroE gene under the control of the tac
promoter. The sequence of events for the construction of SP1.1 and the structure of the

basic plasmid pKD12.112A are as shown in Figure 34.

Host Strain: Plasmid:
RB791 pKD12.112
serA::aroB ”OFFER
(homologous recombination)
RB791 serA::aroB serA
fl-Iac

aroLzzTn 10
(P1 transduction)

 

RB791 serA::aroB aroLzzTn 10

 

SP1 .3

aroK::CmR
(P1 transduction)

 

 

RB791 serA::amB aroLzzTn 10 amK::CmR

||
$91.1

 

Figure 34. Strain SP1.1 and Plasmid pKD12.112A

~56-

B) Fed-Batch Fermentor Conditions

Fed-batch fermentations were performed in a 2.0 L capacity Biostat MD B-Braun
fermentor equipped with a DCU system and a Dell Optiplex Gs+ 5166M personal
computer utilizing B-Braun MFCS/win software for data acquisition and automatic
process monitoring. The fermentation vessel was modified by introduction of a stainless
steel baffle cage containing four 1/ ” x 4” baffles for all fermentations. Fermentations
were run at 33 °C, pH 7.0 and the dissolved oxygen (D.O.) level was maintained at 10%.
All three parameters were controlled by PID control loops. 'A Metler-Toledo 12 mm
sterilizable O2 sensor fitted with an Ingold A-type permeable membrane was used for
monitoring D.O. levels.

Inoculants were initially grown in 5 mL of complete M9 medium for 24 h at
37 °C and 250 rpm. This culture was then transferred to 100 mL of the same medium
and grown under identical conditions for 10 h before being transferred to the fermentor.
The initial glucose concentration was kept at 30 g/L. The entire fermentation can be
divided into three stages, each of which corresponds to a different procedure for
controlling the DO. level at 10%. In the first stage, the air flow was kept constant at
0.06 LIL/min and the stirrer was ramped up from 50 rpm to 750 rpm at a rate sufficient
enough to maintain the DC. at 10%. Once the stirrer reached its preset maximum of 750
rpm, the mass flow controller increased the air flow from 0.06 LIL/min to 1.0 LIL/min.
These two stages take anywhere from 9 h to 18 h for completion depending on the strain
under investigation. Until this stage, except for the initial 30 g/L, no additional glucose
was added. As soon as the air flow reached 1.0 LIL/min at a constant impeller speed of

750 rpm, the glucose pump was switched on and hereafter the DO. level was maintained

-57-

 

 

 

 

[(I‘

no .

 

int

dc

steady at 10% by varying the stirrer from 750 rpm to 1600 rpm. Concentration of the
glucose solution added was 60% (w/v). The rate of glucose addition was controlled so as
to maintain a glucose concentration of approximately 25 g/L in the fermentation at any
given time. Determination of glucose concentrations during the fermentation was done
using the Glucose (HK) 20 assay kit purchased from Sigma. After the third stage was
initiated, glucose concentration in the fermentation culture was measured every hour for
six hours. For the next six hours, glucose concentration was monitored every two hours
and thereafter measurements were done every six hours until the end of the run.
Fermentation runs typically entered logarithmic phase of growth 6 h after
inoculation and usually achieved stationary growth phase after 24 h. Microbial cell
density normally reached 25 g/L to 30 g/L dry cell weight. Fermentations were run for
60 h. The cultures took on a dark color as the fermentation progressed and after 60 h
were deep black in color. Rate of production of SA generally picked up after 18 h and
attained a plateau in 54 h to 60 h. DHS and quinic acid (QA) accumulation was also
observed in all the fermentations. The formation of these by-products is discussed in
detail in Chapter 4 of this thesis. In fact, the dark color associated with all SA

fermentations is due to the air oxidation of DHS to gallic acid.

-53-

Overexpression of Transketolase and Transaldolase Under Fed-
Batch Fermentor Conditions

The first biocatalyst which was tested as a shikimate producer was
SP1.1/pKD12.112A. Plasmid pKDl2.112A carries a copy of arolEFBR under the control
of its native promoter, a copy of the aroE locus under the control of the tac promoter, a
copy of the serA gene and the fl-lac gene conferring Ap resistance. This strain lacks any
capability to alleviate E4P limitations. The construction of pKD12.112A was initiated by
ligation of the 1.2-kb PmaroE fragment into pKL4.20B (Figure 35). Plasmid pKL4.20B
had been previously created in the laboratory and carries the arol'i'FBR locus on a pSU18
vector. Vector pSU18 is a 2.3-kb plasmid carrying a p15A origin of replication and has a
high copy number of approximately 12 per cell.97 It also contains a lac promoter and a
genetic marker encoding for Cm resistance. The PmaroE fragment was obtained by PCR
amplification from pIA321,98 digested with KpnI and ligated with pKL4.20B (3.6-kb)
which had been linearized by KpnI treatment. This resulted in the 4.8-kb plasmid,
pKDlZ.036A (Figure 35). The orientation of the PmaroE locus in pKDl2.036A is in the
opposite direction as that of aroFFBR. The serA locus was obtained by digestion of
pD262595b with EcoRV and DraI. Blunt end ligation of this 1.9-kb fragment into
pKD12.036, which had been linearized by SmaI digestion, afforded the 6.7-kb plasmid
pKDl2.047A (Figure 36). The orientation of the serA gene was in the same direction as

that of P aroE. The fi-lac gene was amplified by PCR from pUC18, treated with NcoI

(ac

and ligated into the NcoI site of pKD12.047A resulting in the 7.7-kb plasmid

pKDlZ.112A (Figure 37).

-59-

pIA321

lPCR

 

 

 

 

Kf’m 1.2-k6 K‘j’"
Ptac' aroE
. i) Kpnl digest
lenl digest iii) CIAP treatment

pKD12.036A

4.8-kb

 

Smal
Hinotll

Figure 35. Preparation of Plasmid pKD12.036A.

-60-

pKD12.036A

 

4.8-kb
p02625
EcoRV/
Dral digest
Dral 1 .9-kb EcoRV
serA Hinoill
i) Smal digest
lKlenow treatment in) CIAP treatment
Ligate
EcoFil
EcoRl
Kpnl
pKD1 2.047A
Ncol CmR 6.7-kb

 

Hindlll “3 ‘ ' Kp
(Smal) (Smal)

Figure 36. Preparation of Plasmid pKD12.047A.

-6l-

 

") CIAP treatment

_ i) Ncol digest
Ncol digest '

Ligate

pKD12.112A

7.7-kb

 

Figure 37. Preparation of Plasmid pKD12.112A.

~62-

SP1.1/pKDlZ.112A was examined under fed-batch fermentor conditions. After
60 h this biocatalyst had produced 38 g/L SA in 12% yield (mol of SA produced! mol of
glucose consumed) (Figure 38) (entry 1, Table 4). The maximum productivity of SA was
between 18 h and 36 h, approximately 1 g/L/h, after which it slowed down to 0.5 g/Uh,
but production never leveled off. Accumulation of DHS and QA as by-products was also
observed. The total yield of the fermentation, taking in account SA, DHS and QA, was
15%. DAHP synthase specific activity was measured every six hours, starting at 12 h
into the run, up to 48 h. DAHP synthase specific activity remained moderately high and
stable throughout the run and showed only a slight decline from 42 h to 48 h (entry 1,

Table 2).

 

4O 25
35 ‘* ‘ A A A
30 1- l A
25 -- A w l

20 -
15 -- . ‘ l i
10 -- l l l l

l l ‘
5 -I- i ‘ t. 1 ‘
. \ \ \ \
0 . ‘ L' C >_ .5. CI tl bl bl _ 0
0 12 18 24 30 36 42 48 54 60
time (h)

D
N
0

L15

I

is
dry cell wt. (g/L)

SA, DHS, QA (g/L)

 

 

Figure 38. SP1.1/pKD12.112A fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l l QA(g/L).

-63-

pKD12.112A

 

PMF5‘A 7.7-kb
l BamHl digest
BamHl 224m BamHl
tktA

ii Klenow treatment

i) Hindlll digest
iil) CIAP treatment

lKlenow treatment

Ligate

Ncol

  

(Hindlll)

(Hindlll) (Smal)

Figure 39 . Preparation of Plasmid pKD12.138A.

-64-

As stated before, DAHP synthase activity is known to be limited by E4P
availability, which is in turn controlled by transketolase and transaldolase expression.68
These enzymes are operative in the non-oxidative pentose phosphate pathway and are
responsible for the interconversion of C-4, C-5, C-6 and C-7 aldoses and ketoses (Figure
20). Therefore, the next logical avenue to pursue was to amplify either transketolase or
transaldolase expression. Plasmid pKD12.138A was created by cloning of the tktA locus,
encoding for transketolase, into pKDlZ.112A (Figure 39). Plasmid pKD12.112A was
digested with Hindlll and treated with Klenow fragment. Following digestion of
pMF51A72a with BamHI, the 2.2-kb tktA fragment was blunt ended using Klenow
fragment and ligated into linearized pKD12.112A using T4 ligase resulting in the 9.9-kb

plasmid, pKD12.138A. The tktA gene is transcribed in the same direction as the serA

 

 

 

gene.
60 30
A
,3 50 "L' ‘ ‘ ‘ "'25 A
3’ at
5 40-? -- 20 ‘2,
(,5 30 -- «15 E
I 8
o
- 20 -- «10 a
as . x U
4 ‘ ‘ .-
0. s‘ a s. 5. ti 5| fin to
O 12 18 24 30 36 42 48 54 60
time (h)

Figure 40. SP1.1/pKD12.138A fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l l QA(g/L).

-65-

Plasmid pKD12.138A was transformed into SP1.1 and this biocatalyst was
examined under fed-batch fermentation conditions. At the end of 60 h,
SP1.1/pKD12.138A had produced 52 g/L of SA in 18 % (mol/mol) yield (Figure 40)
(entry 2, Table 4). A corresponding increase in accumulation of DHS and QA was also
observed and the total yield (mol/mol) was determined to be 24% from glucose. The
DAHP synthase activity levels were however measured to be much lower than those for
SP1.1/pKDl2.112A (entry 2, Table 2). A plausible reasoning for this decline is the

metabolic burden imposed on the system due to expression of the additional tktA gene.

Table 2. DAHP synthase activities (umol/min/mg) for PEP limited biocatalysts.

 

 

DAHP synthase specific activities

 

 

Entry no. Strain [SA] (g/L)
12 h 24 h 36 h 48 h
1 SP1.1/pKD12.1 12A 38 0.37 0.35 0.37 0.31
2 SP1.1/pKD12.138A 52 0.068 0.19 0.13 0.096
3 SP1 .1/pSC4.295A 32* 1 .36 0.47 0.46 0.61

 

 

* Amount accumulated after 48 h as opposed to 60 h.

Alleviation of E4P limitation was also attempted by localization of the MIR locus,
encoding for transaldolase, in plasmid pSC4.295A (Figure 41). The 1.3-kb talB fragment
was isolated by digestion of pMF52A72‘ with NcoI. Plasmid pKDlZ.047A was linearized
by treatment with Neal and ligated with the MIR gene using T4 ligase resulting in the 8.0-

kb plasmid, pSC4.295.

-66-

 

6.7-kb
lNcol digest
Neal 13“) Neal
LEW
taIB
i) Ncol digest
l i ii) CIAP treatment

 

Figure 41. Preparation of Plasmid pSC4.295A.

-67-

 

The SA producing capability of SP1.1/pSC4.295 was determined under
fermentation conditions and the results were not encouraging. Accumulation of only
32 g/L SA was observed after 48 h (Figure 42) (entry 3, Table 4), 'which was identical to
that obtained with SP1.1/pKD12.112A, and hence the fermentation was terminated at that
stage. This result indicated that talB overexpression had negligible effect in increasing
carbon flow into the common pathway. Activity of DAHP synthase remained high

throughout the run and was not considered as a factor in the low SA production (entry 3,

 

 

 

 

Table 2).
35 30
A 30 ... ‘ A ‘ A " 25
3 25 - ’3
I i r 20 5’
o 20 -. E‘
m“ .- 15 =
I 15 -. 8
D l- 10 a
<' 10 1' "O
(D
5 4- A l 1 ‘ S -- 5
0 fi .\ S R § 5 _ 0
0 12 18 24 30 36 42 48
time (h)
Figure 42. SP1.1/pSC4.295A fed-batch fermentation time course.
A Dry cell wt. M» I SA mm m DHS(g/L); l l QA(g/L).

-68-

Production of SA as a Function of Increased PEP availability

With E4P levels no longer being a limiting factor in SA synthesis, the next aspect
examined was availability of PEP. Intracellular concentrations of PEP can be improved
by increasing in vivo PEP synthase activity levels.‘““'e This was realized by plasmid
localization of the pps gene (Figure 43). Digestion of pKL1.87B99 with EcoRI and
Hindlll afforded the 3.0-kb pps gene, which was subsequently treated with Klenow
fragment. Plasmid pKD12.138A was digested with NcoI and the 8.9-kb fragment treated
with Klenow fragment. Ligation of the pps gene into linearized pKD12.138A afforded
pKD15.07lB. Transcription of the pps gene is in the orientation opposite to that of the
tktA gene.

Coupling pps amplification with that of tktA had a dramatic effect on SA
production. Strain SP1.1/pKD15.071B synthesized 66 g/L of SA in 23% (mol/mol) yield
(Figure 44) (entry 4, Table 4). The overall yield taking into account DHS and QA
accumulation was calculated to be 29% indicating an increase in carbon being shuttled
into the common pathway. Surprisingly, DAHP synthase activity levels were measured
to be higher than those observed for SP1.1/pKD12.138A (entry 1, Table 3). This was not
expected considering that transcription of pps in addition to the tktA gene should subject

the system to further metabolic stress and hence lower DAHP synthase activity.

-69-

pKD12.138A

 

      
 

9.9-kb
pKL1 .87B
EcoRl/
Hindlll digest
Hindlll 3.0-kb EcoRl
pps UfifldflXShwm
i) Ncol digest
Klenow treatment il) Klenow treatment
iii) CIAP treatment
Ligate
EcoRl
l‘ EcoRl
Kpnl
pKD1 5.071 B
1 1 .9-kb
Kpnl
(Smal)

(Hindll)

Figure 43. Preparation of Plasmid pKD15.07 1B.

-70-

 

on
O
O.)
O

 

 

.L A
3 70 ‘ -. 25
60 «- ‘ "‘
3’ A A "
5 50 ._ .. 20 g
a)" 40 1" ’ 15 E
I 30 -- 3
2 - 10 a
(D 20 "" 5 U
A . .t
10 -- \ g N‘ \
‘I \
0 q ‘ A \— g- >— t- :— §I 5' _ O
0 12 18 24 30 36 42 48 54 60
time (h)
Figure 44. SP1.1/pKD15.071B fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l l QA(g/L).

The primary effect of pps overexpression is to nullify PEP limitation caused by
PTS-mediated glucose uptake. The same effect can be realized by utilization of a non-
PEP dependent mode of glucose transport. Zymomonas mobilis is known to utilize a
facilitated diffusion system (Figure 24). Ingram and co-workers have successfully
utilized the Z. mobilis glucose facilitator gene designated glf, to complement glucose
transport in E. coli strains lacking a functional native glucose uptake system?“ In order
to study the effect that facilitated diffusion would have on SA production, the glf gene

was localized on plasmid pSC5.112 under the control of the tac promoter (Figure 45).

-71-

 

pKD12.138A

9.94m
Hinalll Xba'

BamHl/
Hindlll/
Xbal digest

Hindlll 2.2-”, BamHl

 

9,, Pm (Hindll)(Smal)

i1) Klenow treatment

1) Noel digest
iit) CIAP treatment

1 Klenow treatment

pscs.1 123 aroE
1 1 .1-kb

 

(Smal)
(Hindlll)

Figure 45. Preparation of Plasmid pSC5.112B.

-72-

The 2.2—kb nglf fragment was obtained by digestion of pTC32573d with BamHI,
Hindlll and X baI followed by treatment with Klenow fragment. Digestion of
pKD12.138A with NcoI excised the 1.0-kb B—lac gene and the remaining 8.9—kb fragment
was treated with Klenow fragment. Ligation of the ngIf fragment into linearized
pKDlZ.l38A afforded pSC5.1 12B.

Use of the glucose facilitator system to increase PEP levels had a positive impact
on SA production. After 60 h, SP1.1/pSC5.112B had synthesized 70 g/L of SA in
addition to 19.4 g/L DHS and 6 g/L QA (Figure 46) (entry 5, Table 4). The result was
nearly the same as that observed when overexpression of PEP synthase was utilized to
improve PEP influx into the common pathway. However, since PEP synthase is a
heavily regulated enzyme, its applicability as a long term solution is questionable and use

of the facilitated diffusion system might provide an attractive alternative.

 

 

 

80 30
A
A 70 J. ‘ ‘ 'l I '4- 25
.1 60 -- A ‘ A A
a, .1
5 50? I. - 20 35
m- 40 -- ‘1 ~15 3
E 304- 3
<‘ " 1° 5‘
(’3 20 4 ‘ \ S
10 1' I Q § * \‘ ‘s‘ W‘ 8 § F5
‘
0. . a- s- s. 5. s. b. >- '-o
0 12 18 24 30 36 42 48 54 60
time (h)
Figure 46. SP1.1/pSC5.112B fed-batch fermentation time course.
A Dry cell wt. (g/L); [-3 SA (g/L); m DHS(g/L); I i QA(g/L).

-73-

Table 3. DAHP synthase activities (umol/minlmg) for non-PEP limited biocatalysts.

 

 

 

 

DAHP synthase specific activities
Entry no. Strain [SA] (g/L)
12 h 24 h 36 h 48 h
1 SP1.1/pKD15.071B 66 0.16 0.32 0.26 0.20
2 SP1.1/pSCS.1123 70 0.076 0.21 0.19 0.18

 

 

The results discussed earlier with SP1.1/pSC5.112B involved the use of the
glucose facilitator protein in the presence of a functional PTS system. Examination of
the overall impact that facilitated diffusion by itself has on SA production entailed the use
of an E. coli host in which the native PTS system has been compromised. This was
accomplished by disruption of three genes involved in PTS-mediated glucose uptake
namely, ptsH, ptsl, and crr (Figure 21) to furnish the strain SP1.1pts. Deletion of these
genes renders the strain defective not only in glucose transport, but also in glucose
phosphorylation. Hence, effectiveness of utilizing the glucose facilitator protein is
contingent upon providing the requisite glucose kinase activity. Plasmid pTC325, which
was used as the source of the glf locus, also carries the glk gene encoding for the
Z. mobilis glucose kinase enzyme (Figure 45). Digestion of pTC325 with BamHI and
Xbal excises the 3.9-kb Pm glf glk fragment, which was reacted with Klenow fragment.
Plasmid pKDlZ.138A was digested with NcoI and the 8.9-kb fragment treated with
Klenow fragment. Ligation of the two DNA segments afforded the 12.8-kb plasmid,

pSC6.09OB (Figure 47).

-74-

     
 
 

pKD12.138A
9.9-kb

pTC325 Ncol I,
. Kpnl
BamHl/ (Hindlll
leal digest (Smal)
Xbal 3.9-kb BamHI
i) Ncol digest
Klenow treatment ii) Klenow treatment
iil) CIAP treatment
Ligate
EcoRl

      
  

pSC6.0908
1 2.8-kb

  

(Smal)

(”00" (Hindlll)

(Hindll)

Figure 47. Preparation of Plasmid pSC6.090B.

-75-

Plasmid pSC6.09OB was transformed into SP1.1pts and this strain was evaluated
under fermentation conditions. SP1.1pts/pSC6.090B produced 71 g/L of SA after 60 h in
27% (mol/ mol) yield (Figure 48) (entry 6, Table 4), and appeared to be only slightly
better than SP1.1/pKDlS.071B or SP1.1/pSC5.1 12B. But a closer examination of the
production profile indicated that this biocatalyst had synthesized 54 g/L of SA after only
30 h as opposed to 32 g/L for SP1.1/pKD15.071B and 37 g/L for SP1.1/pSC5.112B.
Unfortunately, this rapid accumulation rate was not sustained and the end result was
similar to those obtained with the aforementioned biocatalysts. Possible reasons for the
decline in production will be discussed later in this Chapter. The effectiveness of
pSC6.09OB in a system with a functional PTS system was also investigated by
transforming the plasmid into SP1.1. Fermentation with this strain however provided
only 46 g/L SA after 60 h in 21% (mol/mol) yield (Figure 49) (entry 7, Table 4). A
distinguishing feature of this fermentation was the accumulation of 19 g/L of acetate.
The low level of SA accumulation may be a direct result of the acetate being toxic to the
cells.

Acetic acid is a by-product of glucose metabolism, and its level can affect the
production of the fermentation process by slowing the bacterial growth and/or inhibiting
recombinant protein biosynthesis.100 Acetate accumulation presumably occurs due to
overloading of the tricarboxylic acid (TCA) pathway by fast oxidation through
glycolysis. Under high glucose concentrations, excess acetyl CoA flows into the TCA
cycle which ultimately translates into higher acetate formation. Strain SP1.1/pSC6.09OB
possesses an intact FT S system as well as an amplified facilitated glucose uptake

mechanism. Intake of glucose by the cell would therefore be at an unprecedented level.

-76-

As opposed to SP1.1/pSC5.112B, which also utilizes facilitated diffusion coupled with

the PTS system, strain SP1.1/pSC6.09OB carries the glk gene which appears to make

some difference.

 

 

 

 

80 30
A 70 ‘r ‘ 1- 25
—l 60 -. A ‘
5’ ‘ -- 20
<1: 50 1-
0
g 40 -11- "" 15
3. 3° " -- 10
(D 20 A § 5 S - ‘1 -_ 5
10 " s 5 § § * V 8 i
0 q \- K- 5' \I \I \I 5' \I _ 0
0 12 18 24 30 36 42 48 54 60
time (h)
Figure 48. SP1.1pts/pSC6.090B fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l i QA(g/L).
50 30
45 «-
A 40 «l- ‘ " 25
.1 A ‘
o " 1
(if :5 “ I o "' 15
D- 12 ° -- 10
a ,0 ° .
5 .. ‘ ~ ‘0. ‘ N S ‘ 5 § -. 5
. ‘1 \ § \ ~ ‘ ‘
0 q . o .0- \_ \- L I \l . - \I 5' _ 0
0 12 18 24 30 36 42 48 54 60
time (h)
Figure 49. SP1.1/pSC6.090B fed-batch fermentation time course.
A Dry ceu wt. (gIL); I SA (g/L); m DHS(g/L); l l QA(g/L)

' 5 acetate (gIL).

 

 

-77-

dry cell wt. (g/L)

acetate, dry cell wt. (g/L)

Plasmid pSC6.162A was constructed to study the effect of stacking the pps and
glf genes in the same system. Localization of the 2.2-kb nglf fragment from pTC325
into pKLl.87A resulted in pSC6.142B, in which the ngIf fragment was transcribed
opposite to the pps gene (Figure 50). The 5.2-kb Pmcglfpps portion was subsequently
excised out of pSC6.142B by BamHI/HindIII digestion, treated with Klenow fragment
and inserted into the NcoI site of pKDl2.138A to generate pSC6.162A(Figure 51).

Plasmid pSC6.162A was transformed into SP1.1 and the resulting strain was
cultured under fermentation conditions. Coupling overexpression of the glucose
facilitator enzyme with that of PEP synthase did not give any increment in SA production
over the strains overexpressing either one of the two enzymes. After 60 h, 67 g/L of SA
in 26% (mol/ mol) yield was obtained along with a total yield of 34% (mol/mol) (Figure

52) (entry 8, Table 4). No benefit with respect to rate of production was accrued either.

-73-

PTC325 pKL1.87B

 

meHV

Ifimiw

Xbal digest

Hindlll 2.2-”, BamHI
9” PMC

i) Salt digest

Klenow treatment ii) Klenow treatment
iii) CIAP treatment

 

Hindll

Figure 50. Preparation of Plasmid pSC6.142B.

-79-

 

; pKD12.138A
pscs.14213 i 99'“
Ncol
BamHl/
lHindlll digest (Hindlll) (’Spn’iil)
Hindlll 5.2-kb BamHI
pps glf Ptac (Hindlll)(Smal)

i1) Klenow treatment
iii) CIAP treatment

1 Klenow treatment

1 i) Ncol digest

     

pSC6.162A
14.1 -kb

(NCO') (Hinoill)

Figure 51. Preparation of Plasmid pSC6.162A.

-30-

on
O
N
01

 

 

 

 

7o -- ‘ A ‘ ‘ ‘
g 60 -- ‘ 20 3
v 3)
<1: 501- 15 "T
o E
05 40 -- —
I 713
D 30" ~10 ;
<. 20 '1' A U
U) ‘ 5
10 -- ‘1 \ ‘ : § §
0 A I__ - s 3- §I :- gl 5- 5l 5' , O
0 12 18 24 30 36 42 48 54 60

time (h)

Figure 52. SP1.1/pSC6.162A fed-batch fermentation time course.
A Dry cellwt.(g/L); - SA (g/L); N DHS(8/L); .[ i QA(g/L).

Plasmid pSC6.301A was constructed to evaluate the impact of PEP synthase
overexpression coupled to that of the glucose facilitator and glucose kinase proteins.
Excision of the pps gene from pKLl.87B by digestion with BamHI and Hindlll was
followed by treatment with Klenow fragment. Plasmid pSC6.0908 was linearized by
reaction with Xbal and the 12.1-kb fragment was treated with Klenow fragment.
Ligation of the two fragments resulted in the 15.1-kb plasmid pSC6.301A (Figure 53).

Transcription of the pps was in the same orientation as that of tktA.

-31-

     

pSC6.0908
12.8-kb

pKL1.87B
BamHl/
Hindlll digest
Hinalll 3.0-kb BamHI \. ’_ . ’ glint!”

 

(Hindlll) (Hindlll)

i1) Klenow treatment

i) Xbal digest
iii) CIAP treatment

Kpnl

psce.301A (Smal)

15.8-kb

(Smal)
(Xbal)

 

(Hindlll) (Xbal)

Figure 53. Preparation of Plasmid pSC6.301A.

-32-

 

This plasmid was transformed into SP1.1 as well as SP1.1pts and both strains
were run on the fermentor. Neither biocatalyst proved to be a good SA producer as
illustrated by in Figures 54 and 55. In fact, accumulation of large amounts of acetate was
observed in both fermentations, with SP1.1pts/pSC6.301A producing the highest amount
of acetate (30 g/L) recorded so far among strains designed to synthesize SA. It is very

likely that the high acetate concentrations compromised the health of the cells resulting in

 

 

 

poor production of SA.

45 35
40 JP . -r 30 :1“
3 351. ‘ 25 E
g 30-1 ‘ A . ' '5'
O 251- ~- 20 8
5 201- 1 . -15 5
Ci 15-+ 1 _ ‘ a;
< 1 ° . -- 10 *5
(D 10 + ‘ ‘ 1 , ~ ,5
1 . I ‘ ‘ 1 . 1‘ 1 ‘ q1.. 5 0
5 -. 1‘ . § § § § § ca

01 . .. 3 5°- ‘ 5- 5| ' \I \l \l 5' r0

01218 24 so 35 42 4e 54 60
time(h)

Figure 54. SP1.1pts/pSC6.301A fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); ® DHS(g/L); l l QA(g/L)
‘ ? acetate (glL).

-33-

 

 

 

 

 

 

45 30
40 " e 25 3
g 35 -- ‘ ‘ ' g
I 30 .. . ‘- 20 ‘_§
0 25 ~r- L 15 '8
g3 20 4- ’ . a
0 'o
q 15 " o ‘i' 10 m"
< . w iii
U) 10 _+ ‘ . ‘ ‘ ~ ‘1‘- 5 45
1 \ \ \ \ o
5 " .. 4 § \ \ \ § <5
0 q , . E c- b- \I bl 5. 5| \I L0
0 12 1 8 24 30 36 42 48 54 60
time (h)
Figure 55. SP1.1/pSC6.301A fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l J QA(g/L)
acetate (g/L).
Table 4. Titers and yields for SA production by SP1.1 based biocatalysts.
Entry no. Strain [SA] (g/L) SA yield [DHS] (g/L) [QA] (g/L) Total yield
(mol/mol) (mol/mol)
1 SP1.1/pKD12.112A 38 12% 6 2 15%
2 SP1.1/pKD12.138A 52 18% 1 1 4 24%
3 SP1 .1/pSC4.295A 32 16% 6 0 19%
4 SP1.1/pKD15.071B 66 23% 16 4 29%
5 SP1.1/pSCS.1128 7O 24% 19 6 32%
6 SP1 .1 pts/pSC6.0908 71 27% 15 5 34%
7 SP1.1/pSC6.0908 46 21 % 16 3 28%
8 SP1 .1/pSCS.162A 67 26% 17 5 34%
9 SP1.1pts/pSCG.301A 40 15% 9 3 19%
10 SP1.1/pSC6.301A 41 16% 12 2 21%

 

 

-34-

Examination of Lowered Osmotic Stress on SA Production

By this stage, it was fairly surprising that the production levels of most of the
strains peaked out at close to 70 g/L SA after 60 h. For example, stacking the glf and pps
genes was expected to give at least a slight increment over the strains overexpressing the
individual genes. But as indicated in Table 4, that was not the case. It was also
surprising how the productivity of SP1.1pts/pSC6.09OB suddenly leveled off after a
sudden burst in SA synthesis. If this construct had sustained the high level of SA
production for a slightly longer period, it could have accumulated greater than 71 g/L SA.
It was entirely possible that the barrier of approximately 70 g/L was the maximum
production capability of the biocatalysts designed so far. Obtaining higher SA titers and
yields would then involve optimizing the activity of enzymes like DAHP synthase,
transketolase, shikimate dehydrogenase, etc. Additional copies of the genes encoding for
these enzymes could be localized on a new plasmid or on the genome. Utilization of
stronger promoters could also be investigated. But the possibility also existed that some
of the strains could indeed synthesize more SA but were restricted in their ability to do so
by the prevailing conditions in the fermentor. One factor could be that the high
concentrations of SA itself might be impeding further production. It was considered
prudent to discount the latter possibility before constructing new plasmids or strains.

Strain SP1.1/pKD15.071B was selected to establish whether high external SA
concentration was responsible for the limitation on product synthesis. As was discussed
earlier, this biocatalyst had the capability to produce 66 g/L of SA after 60 h (entry 4,
Table 4). For the purpose of this investigation SP1.1/pKDlS.O71B was run under normal

fermentation conditions and at 18 h and 24 h into the experiment, 10 g each of SA was

-35-

added into the fermentor. If external SA concentrations had no influence whatsoever on
the efficiency of the biocatalyst, then SP1.1/pKDlS.07lB would accumulate at least 86
g/L SA after 60 h. On the other hand, if SA concentration did indeed have a deleterious
effect on SA production, then the fermentation would result in accumulation of only 70
g/L SA after 60 h. As illustrated in Figure 56, this experiment yielded 71 g/L of SA after
60 h. This experiment provided evidence that high SA accumulation was indeed limiting
its own biocatalytic synthesis. Hence, construction of a new plasmid or strain to afford
higher SA titers would not be useful unless the mechanism for this limitation is

understood and solved.

80

70 4

60 .4 ‘

50 -~ I

40 -

30 -

20 -- * '

10 4t ‘ ‘ Q ‘ S S
0 J A u D- E- t. 5. :n t t. _ 0

01218 24 30 36 42 48 54 60
time(h)

(D
O

 

D
D
N
01

N
O

a:
dry cell wt. (g/L)

+10

SA, DHS, QA (g/L)

O1

 

 

 

Figure 56. Time course for fed-batch fermentation experiment involving addition of
20 g SA to SP1.1/pKD15.071B culture.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l l QA(g/L)

Several plausible explanations can be offered to account for the restrictive effect
of high SA concentrations on the biocatalyst. The simplest explanation is that beyond a

certain concentration, SA acts as an inhibitor to one of the common pathway enzymes or

-86-

to an enzyme essential to the cell metabolism. Another possibility is that the cell
machinery is still functional and is still capable of SA synthesis. But the high levels of
product accumulation could have shut down the necessary export systems required by the
cell, which in turn would shut down SA production. The option which was finally

‘ In fermentation

investigated as the most likely cause, was that of osmotic stress.'0
processes with high product yields, it is known that cells are exposed to high osmotic
stress from the medium components and the product. This stress can inactivate the cells
and limit product accumulation. The use of osmoprotectants in such fermentations is
fairly widespread. Compounds like betaine, proline, choline and ectoine are zwetterionic
solutes, which can stimulate the growth rate of bacteria in media of high osmolality
(Figure 57).'02 Betaine has been reported to be the osmoregulant of choice for E. coli.

Therefore, for the purpose of this study betaine was used to study the effect of lowered

osmotic stress on a SA producing system.

02

”IO

(gt-I3 9H3 N
H30 'Nt—C H2C02 H3C “NLCHchon 4 )"CO; 0
l l 'N‘
CH3 CH3 H3O +CH3 H3C H’Q‘H
betaine choline L-proline ectoine

Figure 57. Common osmoprotectants used in microbial fermentations.

Strain SP1.1/pKD15.07lB was cultured under normal fermentation conditions
with 10 mM betaine added as an osmoprotectant. No improvement was seen in the titer
or yield of SA after 60 h (Figure 58) (entry 2, Table 5). The concentration of betaine was

hence increased to 100 mM and the fermentation repeated. In this instance, the titer of

-37-

SA declined compared to normal runs indicating that high concentrations of betaine
might be detrimental (Figure 59) (entry 3, Table 5). A couple of aspects of this
fermentation are worth nothing though. The amount of DHS produced was the highest
ever measured and hence the total yield was quite high at 38% (mol/mol). Also, at the
end of the run, it became clear that the cells had taken up approximately 30 mM betaine,
which gave an indication of what the optimum betaine concentration might be for use in
the fermentation. Therefore, a third fermentation was set up containing 30 mM betaine
and performed in an identical fashion as before. After 60 h, SP1.1/pKD15.071B had
accumulated 71 g/L SA in 26 (mol/mol) yield, which is the best result observed so far for
this strain (Figure 60) (entry 4, Table 5). The conclusion can therefore be drawn that
even though osmotic stress does play some role in suppressing SA production, other

factors exist that need to be investigated.

 

 

 

 

7O 25
60 -- ‘ A A r . . .
j 50 .. . A -- 20 j
:2 3’?
q 40 .. E
g 30» «10 S
(‘5 20 4- A ‘ Q ‘ “5 5

104 .. ‘ ‘ N
0 A . .. ' R S. 3. SI SI SI SI SI _ 0
o 12 18 24 30 36 42 48 54 60

time (h)

Figure 58. SP1.1/pKD15.071B fed-batch fermentation time course with 10 mM
betaine.
A Dry cell wt. (g/L); - SA (g/L); mDng/L); l §QA(g/L)

-33-

 

 

 

 

60 35
A 50 -- A A -- 30
_l
22 40 .1 ‘ ‘ " 25
O 30 - 20
(:2 0 o o o . . ‘ i 15
Q 20 4- ‘ ' t; 5 h 10
at) \ ‘ < i
10 .. \ \ \
1 \ ~ \ \ 4.. 5
c < i ‘ t ‘ s
O ._‘ I l 1LT \— 5- \- t. kl bl \l _ 0
O 12 18 24 30 36 42 48 54 60
time (h)

Figure 59. SP1.1/pKD15.071B fed-batch fermentation time course with 100 mM

betaine.

 

 

 

A Dry cell wt. (g/L); I SA (g/L); m DHS(g/L); 1 QA(g/L)
~ betaine(g/L).
80 30
7o --
j 6 ‘ "' 25
§ 0 .. ‘ A .. 20
< 50 -- 4
O
(I? 40 .4 - 15
9 30¢ 10
< 20 ..
CD ‘ ~ . w
10-- - ‘ § § § § “5
\' § ‘ \ \ \ \
0 4 \‘l \l \I 2. . I \I 5. 5' L 0
O 12 18 24 30 36 42 48 54 60
time (h)

Figure 60. SP1.1/pKD15.07lB fed-batch fermentation time course with 30 mM

betaine.

A Dry cell wt. (g/L);

- SA (g/L); m DHS(g/L);

-39-

QA(g/L)

dry cell wt. (g/L)

dry cell wt. (g/L)

Table 5. Effect of varying betaine concentrations on production of SA by

 

 

 

SP1.1/pKD15.0718.
Entry no. Strain [SA] (g/L) SA yield [DHS] (g/L) [QA] (g/L) Total yield
(mol/mol) (mol/mol)
1 SP1.1/pKD15.071B 66 23% 16 4 29%
0 mM betaine
2 SP1 .1/pKD15.071 B 66 22% 14 6 28%
10 mM betaine
3 SP1.1/pKD15.071B 57 25% 25 6 38%
100 mM betaine
4 SP1.1/pK015.071B 71 26% 18 5 34%
30 mM betaine

 

 

-90-

Comparison of SA Production Capabilities Between

E. coli B and E. coli K12

The growth rate of E. coli B has been reported to be much more rapid as
compared to E. coli K12 (RB791).103 A faster and more rigorous growth pattern may
result in faster accumulation of SA and the higher cell mass generated could also result in
more SA production. Also, in E. coli K12, DAHP synthase is precedented to be labile to
proteolytic activity during the Stationary phase of growth. E. coli B on the other hand is
known to be naturally deficient in the Lon proteases.93 It is believed that the Lon
proteases are the major proteases catalyzing the endoproteolytic cleavage of proteins in
the cell. Strains disrupted in the Ian gene produce several phenotypic changes including
increased sensitivity to UV and ionizing radiation, overproduction of
mucopolysaccharide, reduced lysogenization of bacteriophages lambda and P1, and most
importantly, reduced protein degradation.93d Examination of the enzyme activity levels
for the biocatalysts derived from RB791 illustrates how DAHP synthase activity levels
tend to fade away after 24 h into the fermentation run (Tables 2 and 3). Given the lack of
proteolytic activity during stationary phase in E. coli B, use of a biocatalyst derived from
this strain could help avoid this problem, making it quite an attractive alternative to E.
coli K12.

An E. coli B derivative EB1.1, capable of synthesizing SA, was constructed
carrying the identical deletions as in SP1.1. Plasmid pKD12.112A was transformed into
EB1.1 and the Strain was tested under fermentation conditions. Production levels of SA
(39 g/L) were disappointingly the same as that observed with SP1.1/pKD12.112A albeit

with a higher yield of 17% (mol/mol) (Figure 61) (entry 1, Table 6).

-91-

.5
01
(A)
O

 

40 .. 25
‘ n-

: 35 -- ‘ A
E 30 -_ -- 20 3
g 25 J- A t;
g; 15 75
D o

.. 10
at) 5

-5

-o

 

 

   

 

01218 24 30 36 42 48 54 60
time(h)

Figure 61. EBl.l/pKD12.112A fed-batch fermentation time course.
A Dry cell wt. (g/L); I SA (g/L); N DHS(g/L); | J QA(g/L)

The 0D,,00 values recorded for the EBl.l/pKD12.112A cell density during the
course of the fermentation were abnormally higher than those observed for any of the
SP1.1 or SP1.1pts runs. However, for EB1.1 a conversion factor of 0.29 g/L/OD,500 had
to be applied in order to arrive at an accurate estimate of the dry cell weight (g/L). This
value was an average value obtained by centrifugation (16000 x g, 10 min) of 20 mL
aliquots of fermentation culture taken every 6 h and drying the washed, harvested cells to
a constant weight at 100 °C. The corresponding value for RB791 derived strains was
0.43 g/L/ODéoo. The net result was that even though the OD600 values measured for E.
coli B fermentations were very high, the final dry cell weight values were the same as
those for the corresponding E. coli K-12 runs.

The fermentation with EBl.l/pKDl2.138A did not Show any improvement over

EBl.l/pKD12.112A, which was unexpected (Figure 62) (entry 2, Table 6). Experiments

-92-

with SP1.1 had shown that amplification of transketolase activity improved the SA yield
(entries 1 and 2, Table 4) and titer Significantly, but no such effect was seen with EB1.1

(entries 1 and 2, Table 6) and in some cases SP1.1 proved to be a better biocatalyst.

 

 

 

 

40 30
A 35 -r A -- 25
-' 30 -- A A
3, .1
< 25 fl - 20 g
0 .
(£- 20 -- A - 15 g
15 - 0
2 -- 10 g.
a) 10 " 4 5
5 «- . \- N R Q Q \ "
\
‘ 'l-u I ' g :- §- §_ §- t. t." _ 0
0 12 18 24 30 36 42 48 54 60
time (h)
Figure 62. EBl.l/pKD12.l38A fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l l QA(g/L)

In order to verify that that the biocatalyst was not limited in PEP, EB1.1 was
transformed with the pps encoding plasmid pKD15.07lB. Fed-batch fermentation with
this strain also proved to be disappointing and produced only 44 g/L SA in 18% yield
after 60 h (Figure 63) (entry 3, Table 6). This result seemed to hint that an SP1.1 derived

host was superior to one based on EB 1 . 1.

-93-

 

 

01
O
00
O

 

 

45 -- A 2
._ 5
g 40 -- ‘ j
v 35 " e -- 20 a
< 30 T ‘1’
o ‘ ‘ E
D .. o
- i. - 10
3‘; 15 5
1g “ \ ‘ : -r- 5
1' \ \
0 fl \ \I b. g- SI §- >u §I F 0
0 12 18 24 30 36 42 48 54 60
time (h)
Figure 63. EB 1.1/pKD15.07 1B fed-batch fermentation time course.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); l I QA(g/L)

Table 6. Titers and yields for SA production by EBIJ based biocatalysts.

 

 

 

Entry no. Strain [SA] (g/L) SA y/ield [DHS] (g/L) [QA] (g/L) Total yield
(mo mol) (mol/mol)

1 EBt.1/pKD12.112A 39 17% 8 2 21%

2 EB1.1/pKD12.138A 37 22% 9 2 28%

3 EB1.1/pKD15.071B 44 18% 1O 2 22%

 

 

Comparison of the values in Table 7 indicate that DAHP synthase specific activity
levels for the E. coli B strain (entries 1 and 2, Table 7) are indeed much higher than the
corresponding values for the E. coli K12 phenotype (entries 3 and 4, Table 7). More
significant are the transketolase activity values reported in Table 8. Native E. coli B

transketolase specific activity (entry 1, Table 8)) was measured to be higher than

-94-

overexpressed transketolase activity in E. coli K12 (entry 4, Table 8). These higher

specific activities however did not translate into enhanced SA levels.

Table 7. Comparison of DAHP synthase activities (umol/minlmg) between EB1.1

and SP1.1 biocatalysts.

 

 

DAHP synthase specific activities

 

 

Entry no. Strain [SA] (g/L)
2 h 24 h 36 h 48 h
1 EB1.1/pK012.112A 39 1.51 1.13 0.99 0.84
2 EB1.1/pKD12.138A 37 0.82 0.80 0.49 0.41
3 SP1.1/pK012.1 12A 38 0.37 0.35 0.37 0.31
4 SP1.1/pKD12.138A 52 0.068 0.19 0.13 0.096

 

 

Table 8. Comparison of transketolase activities (umol/min/mg) between EB1.1 and

SP1.1 biocatalysts.

 

 

Transketolase specific activities

 

 

Entry no. Strain [SA] (g/L)
12 h 24 h 36 h 48 h
1 EB1.1/pKD12.112A 39 0.073 0.17 0.34 0.39
2 EB1.1/pKD12.138A 37 0.11 0.50 0.73 0.53
3 SP1.1/pKD12.112A 38 0.024 0.10 0.14 0.18
4 SP1.1/pKD12.138A 52 0.11 0.17 0.21 0.35

 

 

-95-

Discussion

A) Comparison of Titers and Yields

Efficiency of a biocatalyst can be evaluated by comparing the yield of SA
produced to the theoretical maximum yield (mol of SA produced/ mol of glucose
consumed). Determining the theoretical maximum yield for biocatalytic synthesis of SA
begins with balancing the E4P and PEP inputs with the products and by-products formed
(Eq. (1)). The input of E4P and PEP required for SA synthesis is then equated with the
amount of glucose required to form these substrates (Eq. (2a)). In order to account for
the operation of the PTS-mediated glucose uptake system, a pyruvic acid term is added
(Eq. (2a)). A coefficient of 2.33 is required to balance the number of carbon atoms in the
glucose input to the number of carbon atoms formed in PEP, E4P and pyruvic acid (Eq.
(2b)). Hence, synthesis of 1 mole of SA requires the consumption of 2.33 moles of
glucose resulting in a theoretical maximum yield of 43% (mol of SA produced] mol of

glucose consumed).

(1) PEP + E4P —> 2H3PO4 + H20 + SA

(2a) xglucose —-> PEP + E4P + xpyruvic acid

(2b) x6(C) —" 3(0) + 4(0) + x3(C)

Use of the non-native, facilitated diffusion system for glucose transport eliminates
the conversion of 1 mol of PEP to 1 mol of pyruvate. Exclusion of the term for pyruvic

acid from equation (2b) leads to a coefficient of 1.17 and hence a maximum theoretical

-96-

yield for SA of 86% (mol of SA produced/ mol of glucose consumed) in the absence of
PT S-mediated glucose uptake.

Fermentations producing SA also accumulate moderate amounts of DHS and QA.
Therefore, in order to arrive at an accurate estimate of carbon flow into the pathway
leading to SA, accumulation of DHS and QA must also be considered when calculating
the yield of the fermentation. No biocatalytic strain investigated so far has come close to
the maximum theoretical yield of 43% (mol/ mol) for SA. This is true irrespective of
whether or not the strain was dependent on the native PT 8 system for glucose uptake.
Draths and co-workers have reported104 the use of SP1.1/pKDlZ.112A under slightly
different fed—batch fermentation conditions, details of which are described in chapter 4.
The fermentation produced 18.6 g/L SA along with 7.8 g/L quinic acid QA and 4.4 g/L of
DHS after 42 h. This fermentation suffered from a serious drawback involving the
concomitant formation of QA in large amounts. Efforts have since been made to rectify
this problem and have been discussed in detail in Chapter 4.

Under the fermentation conditions described in this chapter, SP1.1/pKD12.112A,
produced 38 g/L SA in 12% (mol/ mol) yield. As expected, E4P was indeed a limiting
factor in SA synthesis and as observed with SP1.1/pKDlZ.l38A, amplified expression of
tktA gave an increment in SA yield (18%) and titer (52 g/L). The total yield taking into
account DHS and QA was 24%. Alleviation of PEP limitation however gave the biggest
boost to SA production. The highest titer and yield obtained in a PT S-dependent strain
was with SP1.1/pSC5.112B. This strain overexpressed the glf locus from Z. mobilis and
produced 70 g/L in 24% yield. The total yield for this system was 34%. Strain

SP1.1/pKD15.071B, overexpressing for pps, also gave comparable values within

-97-

experimental error. The strain affording the best results was SPl.1pts/pSC6.0908, which
produced 71 g/L SA in 27% (mol/ mol) yield. Accumulation of 15 g/L DHS and 5 g/L

QA was also observed resulting in a total yield of 34%.

B) Effects of Increased PEP Availability

The intracellular concentrations of E4P and PEP have been established to be
limiting factors for directing more carbon into the common pathway after DAHP
synthase has been overexpressed. Li and co-workers have recently reported the
biocatalytic synthesis of DHS and demonstrated the pronounced effect of tktA
overexpression even in absence of pps overexpression.76 Liao and co-workers have
established that overexpression of pps improves carbon flow into the common pathway
only in presence of amplified tktA expression, but the reverse does not hold true.°”°°
Hence, in a biocatalytic environment possessing ample PEP concentrations and
overexpressed, feedback insensitive DAHP synthase, E4P availability is a critical factor
limiting aromatic amino acid biosynthesis. As expected, overexpression of transketolase
did benefit SA synthesis even in the absence of PEP synthase overexpression. All studies
done thereafter involved investigating the effect of increased intracellular PEP
concentration in the presence of overexpressed transketolase.

Although many enzymes including PEP carboxylase, PEP kinase, and PEP
carboxykinase utilize PEP as a substrate, one of the biggest drains on PEP availability for
aromatic amino acid biosynthesis, is the PTS system for glucose uptake.71 Under the

fermentation conditions employed, the cells remain under a very glucose-rich

environment throughout the course of the fermentation. It is therefore quite reasonable to

-93-

expect the availability of PEP flowing into the common pathway to be quite minimal.
Out of all the various strategies employed for increasing in vivo PEP availability,
amplified expression of PEP synthase is particularly effective. Overexpression of PEP
synthase allows for PTS-generated pyruvate to be recycled back to PEP. The other
option of course is to utilize a non-PTS dependent mode for glucose transport, which
would hence allow most of the PEP to be channeled into the common pathway. Both
alternatives were examined to improve shikimate production and both strategies gave
results complimentary to each other. Overexpression of E. coli PEP synthase or Z.
mobilis glucose facilitator protein in SP1.1 gave an increment of approximately 20 g/L
SA over the corresponding PEP limited strain SP1.1/pKD12.138A. The result obtained
with facilitated diffusion was quite heartening, given that PEP synthase is a heavily
regulated enzyme. In fact, its overexpression has been reported to inhibit growth of E.
coli. Glucose uptake by facilitated diffusion on the other hand is a low affinity, high
velocity transport and does not utilize the PTS system nor does it alter PEP synthase
expression levels. The optimum utilization of facilitated diffusion was however realized
when the native PT S glucose uptake system was no longer functional and glucose uptake
was totally dependent on facilitated diffusion. Biocatalyst SP1.1pts/pSC6.090B gave the
best results in terms of SA yield, titer, and rate of production among all SA producing
strains. Between 24 h and 30 h, this biocatalyst accumulated SA at the rate of 5 g/L/h as
opposed to approximately 2 g/L/h for SP1.1/pKD15.07lB and SP1.1/pSC5.112B.
However, the effects of pps and glf overexpression were not additive and no
improvement was observed when these genes were used simultaneously in the same

biocatalyst as observed with SP1.1/pSC6.162A, SP1.1/pSC6.301A and

-99-

SP1.1pts/pSC6.301A. This result may indicate that in vivo PEP concentration was now
sufficiently high, but some other enzyme may now be rate limiting. The likelihood of
high SA concentration inhibiting its own synthesis cannot be discounted either and in fact
results have been obtained consistent with this possibility. Research is currently

underway to investigate how to navigate this problem.

C) E. coli B v/s E. coli K12

DAHP synthase and transketolase activities have been proven to dictate the
efficiency of any biocatalyst designed to produce a common pathway
intermediate/metabolite. Under fed-batch fermentor conditions, biocatalysts derived
from E. coli K12 (RB791) typically suffer from a steady reduction in enzyme activity
once the cell growth reaches stationary phase. Loss in enzyme activity has also been
shown to correspond to decline in the rate of production. This phenomenon has been
observed with fermentations producing SA. Examination of the values reported in tables
3, 4 and 5, indicates that except for SP1.1/pKD12.1 12A and SP1.1/pSC4.295A, DAHP
synthase activity levels were quite low for the other fermentations. The metabolic burden
associated with overexpressing the additional genes took an obvious toll on the strain. It
is quite possible that higher SA yields might be obtained if a biocatalyst was designed
which was capable of maintaining high and steady enzyme activity levels even in
stationary phase. A host strain that could be designed to serve this purpose is E. coli B.
This microbe is known to be naturally deficient in a number of proteases, which could

very well be the solution to achieving higher DAHP synthase activity.

-100-

The biocatalyst EB1.1 is identical to SP1.1 in all respects except that it is derived
from E. coli B instead of RB791. Examination of fermentation results obtained with
EB1.1 indicates that DAHP synthase and transketolase activities were indeed much
higher than the SP1.1 counterpart. In fact native transketolase activity levels in EB1.1
were higher than overexpressed levels in SP1.1. However, the stable and high enzyme
activities did not translate into improved SA titers or yields and EB1.1 proved to be an
inferior biocatalyst to SP1.1. It is quite possible that the export system of E. coli B is
inferior to that of RB791. A maximum barrier of approximately 70 g/L SA has been
observed with RB791 derived catalysts. This barrier maybe closer to 40 g/L for strains
constructed from E. coli B. These results illustrate that although in vivo enzyme
activities play an important role in determining the yield and titer of SA, they are not the
sole governing factor. There might be other controlling aspects involved with SA
production, which might not be in sync with the high enzyme activities in order to derive
any benefit from them. Biosynthesis is a process that involves a delicate balancing of a
range of cellular processes. Optimum results are obtained when all these processes
function in a synchronous fashion with matching efficiencies. We seem to have struck

the right chord in RB791, but we still need to probe for the right tune in E. coli B.

-101—

Synthesis of Phenol from Shikimic Acid

The credentials of SA as an industrially relevant chemical have already been well
established?“92 Providing further credence to its potential as a valuable starting material
comes with the establishment of a synthetic route to phenol from SA. Annual worldwide
production of phenol is estimated to be about 5.2 x 108 kg and is entirely based on

5 The reactivity of the aromatic ring in phenol as well as the

synthetic processes.IO
presence of a functionalizable hydroxy group makes phenol one of the most widely used
chemicals in industry. Commercial uses of phenol include application in phenolic resins,
synthesis of bisphenol A, caprolactam, aniline, and alkylphenols.105

The leading commercial route to phenol is via the oxidation of cumene, which

5 Alkylation of benzene with

accounts for 95% of the global phenol production.lo
propylene under Friedel-Crafts conditions affords cumene. Oxidation of cumene yields
cumene hydroperoxide which upon subsequent cleavage results in phenol and acetone.
The process utilizes carcinogenic, toxic materials and most importantly involves peroxide
intermediates adding the risk of violent explosions to the already hazardous operation.

SA produced by fed-batch fermentations in high titers and yields from glucose
can be transformed into phenol by acid catalyzed dehydration.40 In a high pressure,
stainless steel reaction vessel, SA was dissolved in degassed, carbonated water. The
vessel was sealed and submerged in a sand bath and the temperature of the sand was
slowly raised to 350 °C at a rate of 1.5 °C/min. The reaction vessel was heated at 350 °C
for 30 min, allowed to cool to RT and the aqueous reaction mixture extracted with ether.

Concentration of the organic layer gave a residue which when subjected to Kugelrohr

distillation under reduced pressure afforded pure phenol as white crystals.

-102-

The residue remaining behind after the phenol distilled over mainly consisted of
m-hydroxybenzoic acid. This residue was dissolved in boiling water and returned to the
reaction vessel. A small amount of copper powder was added, the vessel sealed and the
temperature raised to 350 °C at the rate of 1.5 °C/min. After 3 h at 350 °C, the vessel
was cooled to RT and subjected to the identical work-up and purification protocols as

described earlier. The overall yield of phenol from both reactions was 51% based on the

starting SA used.

 

 

 

  

acetone
groggene, solid )
ca
[2 3 4 talyst ’
200-260 °C i) 02, 80—130 ° H0
400-600 psi ii) 302, 60-100 °C
benzene cumene phenol
C02H cozH
"0” Fig.1 H 0* 350°C
—>, a 3 ’ > O +
; OH Ho‘ 5 OH HO HO
HO 6H 6H
D-glucose SA phenol MHB

i l

H20, Cu
350 °C

Figure 64. Comparison of phenol synthesis from benzene and D-glucose.

This conversion represents a novel scheme to convert glucose to phenol via the
interrnediacy of SA. Its overall impact in providing for a safer and environmentally
benign process for producing phenol is quite significant. Development of this route
exemplifies how biocatalysis can be utilized as a very valuable tool for the safe, efficient,

and cheap production of a wide range of chemicals.

-103-

W

INVESTIGATIONS INTO SHIKIMIC ACID-QUINIC
ACID EQUILIBRATION

Introduction

During the early stages of development of a SA producing biocatalyst, a mixture
of SA, QA and DHS was routinely seen when the strains were cultured under fed-batch
fermentation conditions. For example, Draths and co—workers have reported results
discussing the synthesis of SA from D-glucose in SP1.1/pKD12.112.‘°‘ This strain
produced 25.8 g/L SA along with 8.1 g/L quinic acid (QA) and 6.5 g/L of 3-

dehydroshikimic acid (DHS) after 48 hours (Figure 64) (entry 1, Table 9).

 

30 30
A254 ~25
320" 405
g .
”.15.- 15:

Q)
I o
910“ «10 a
as -°

5- -5

or -0

 

 

 

Figure 65. SP1.1/pKD12.112A fed-batch fermentation time course. Glucose limited

conditions, K, = 0.1.

A Dry cell wt. (gIL); - SA (g/L); m DHS(g/L); i l QA(g/L).

-104-

It is important to mention that these results were obtained under glucose-limited
fed-batch fermentation conditions, which are vastly different from the glucose-rich
conditions described in Chapter 3 of this thesis. QA is a secondary metabolite of the
common pathway for aromatic amino acid biosynthesis, produced by the reduction of
DHQ by shikimate dehydrogenase.104 The accumulation of QA is a serious drawback
because it represents a drain on realizable SA yields and titers. More importantly QA co-
purifies along with SA and unless the SA is well in excess of the QA, it is not trivial to
obtain pure SA.106

DHS on the other hand most likely arises due to the feedback inhibition of
shikimate dehydrogenase by SAPS" As highlighted in Chapter 3 of this thesis,
accumulation of as much as 20 g/L DHS has been observed in SA producing
fermentations. Although, it does not pose any hindrance in terms of SA purification, its
production co-relates directly to a loss in SA titer and yield. Development of an efficient
process for the industrial production of SA would therefore entail understanding and
eliminating the source of the QA and DHS contamination.

Before the advent of the excess glucose conditions described in Chapter 3 of this
thesis, fermentations typically employed limited glucose concentrations. It was under
these glucose-limited conditions that QA contamination was routinely observed. Karen
Draths and Dave Knop were successful in eliminating QA formation via catabolite
repression by increasing glucose concentrations.‘°“'107 This indicated that formation of
QA is dependent on uptake of a compound from the external medium, which under the

increased glucose levels is no longer possible. Since SA was in abundance in the

-105-

fermentation broth and since a metabolic pathway exists for the conversion of SA to QA,
it was quite possible that uptake of SA was the source of QA contamination.

In this study, several experiments were performed to validate the theory of QA
arising due to uptake and in vivo processing of SA. Results were obtained that were
consistent with this line of thought. Increased glucose concentrations and use of a
glucose analogue dramatically shifted the SA/QA ratios in favor of SA. A variation of
SP1.1 was also constructed in which the gene proposed to be responsible for shikimate
uptake was knocked out. However, use of this strain did not eliminate SA/QA
equilibration. Hence, although fed-batch fermentation conditions have been developed
which eliminate SA/QA equilibration, the problem still remains to be solved at the

genetic level.

Fed-Batch Fermentor Conditions

The fed-batch fermentor equipment was the same as that described in Chapter 3
except that the baffle cage was not used. Fermentations were run at 33 °C, pH 7.0 and
the dissolved oxygen (D.O.) level was maintained at 10%. Initial glucose concentration
was kept at 20 g/L. The major differences in the fermentation conditions used in Chapter
3 and the studies described in this Chapter lie in the operating parameters used for
controlling the oxygen level.

In the first stage, the stirrer was ramped up to 940 rpm instead of 750 rpm. Once
the stirrer reached this preset limit, it was maintained at this speed and the airflow was
ramped up from 0.06 LIL/min to 1.0 LIL/min in order to keep the DO. level at 10%. In

the third stage, at an impeller rate of 940 rpm and airflow of 1.0 LIL/min, the DO. levels

-lO6-

were maintained steady at 10% by oxygen sensor controlled glucose feeding for the
remainder of the fermentation. Glucose levels were therefore maintained at the bare
minimal level required to maintain healthy cell growth and at no stage in the fermentation
was there any excess glucose present. At the beginning of the third stage, D.O. levels did
fall below 10% due to residual initial glucose still present. This lasted for approximately
30 min until all the glucose was consumed and the controlled glucose feeding was
started. The fermentations were typically run for 48 h after which oxygen sensor control
of glucose became difficult to control and indiscriminate addition of glucose would take

place.

SA/QA Eguilibration: Is Uptake of SA the
Source of QA Contamination?

The observation of SA/QA equilibration was first reported by Draths and co-

 

workers in 1999.104 The strain SP1.1/pKD12.112A produced SA and QA in a molar ratio
of 3.521 after 48 h (entry 1, Table 9). The synthesis of QA by SP1.1/pKD12.112A was
extremely surprising, given the absence in E. coli of quinate dehydrogenase, an
oxidoreductase which interconverts DHQ and QA. Given the similarity in structure
between DHQ and DHS, the possibility arose that aroE encoded shikimate
dehydrogenase could catalyze the reduction of DHS to SA as well as that of DHQ to QA.
Incubation of DHQ with shikimate dehydrogenase resulted in oxidation of NADPH and
concomitant formation of QA.104 The observed rate was approximately one tenth of the
rate for shikimate dehydrogenase catalyzed conversion of DHS to SA. Considering these

facts, a reasonable strategy to eliminate QA contamination would be to minimize

-107-

cytosolic concentrations of DHQ. This could be accomplished by overexpressing aroD
encoded DHQ dehydrogenase. However, even amplified expression of DHQ dehydratase
did not reduce the level of SA/QA equilibration under fed-batch fermentor conditions.’04
This result indicated that QA formation may not result via de novo biosynthesis but rather

from equilibration of initially synthesized SA (Figure 65).

 

HO, C02H H,O C02H
AroB AroD AroE
: ------- (D ....... H
HO OH
R = P03H2; DAHP DOHHS Intracellular SA
R = H; DAH : ‘
: DHIAroE : uptake
1 1
Ho“ C02H 02H
HO"'©\OH Ho'" ; OH
OH . 6H
QA SA in culture
supernatant

Figure 66. The two possible pathways leading to QA production.

Although E. coli does not have the capability to catabolize SA, it has been
reported to possess a transport system specific for SA uptake.108 It is quite possible that
SA transport in E. coli may be an evolutionary vestige of a previous ability to catabolize
SA as a sole source of carbon for growth and metabolism. When E. coli cells are offered
multiple carbon sources, the carbohydrate-metabolizing enzymes are expressed in a
hierarchical fashion. Glucose is the preferred substrate and its presence blocks the
transport of other sugars as well as other less favored carbon sources. Glucose
catabolism is also known to reduce the rate of expression of genes encoding the catabolic

enzymes for other sources of carbon. This phenomenon is termed catabolite

-108-

repression.”109 Uptake of SA could therefore be hindered by high glucose concentrations
in the fermentor.

The rate of glucose addition to the fermentor is controlled by the proportional-
integral-derivative (PID) setting gain (K). The fermentation with SP1.1/pKD12.112A,
which resulted in a 3.5 :1 ratio of SA/QA employed a normal PID setting of K. = 0.1.
Adjusting the PID setting to Kc = 0.8 increased glucose availability, but subjected the
cells to a much more aggressive feeding regimen. When a PID setting of K. = 0.1 was
employed, the glucose feeding rate and dissolved oxygen level usually stabilized a couple
of hours after the third phase was initiated and did not fluctuate much for the remainder
of the experiment. With a PID setting of Kc = 0.8 however, the glucose feed response
was so rapid that a pulsed addition regimen set in, inducing a pulsed dissolved oxygen
profile. Dissolved oxygen stayed at 0% until a glucose pulse was consumed and then
started increasing rapidly. As soon as the oxygen level reached 10%, the glucose pump,
which so far was inactive, switched on and its output ramped up swiftly. Glucose
addition continued at a rapid rate until the dissolved oxygen peaked between 20 - 30%.
A dramatic decline in the dissolved oxygen level was then observed due to the excess
glucose now present in the medium. The glucose addition also decreased at a
commensurate rate and turned off completely at 10% dissolved oxygen level. The
dissolved oxygen kept dropping until it was close to 0%. It remained steady at this level
until all the glucose was consumed and then the cycle was repeated.

The effect of this pattern of glucose addition and dissolved oxygen level Was a
drastic reduction in the formation of QA by SP1.1/pKD12.112A throughout the

fermentation period (Figure 66).104 Since the only difference between this experiment

~109-

and the one run with Kc = 0.1 was the level of glucose, the observed decrease in QA
formation could be attributed to glucose induced catabolite repression of SA uptake. The
overall SA titer was much lower due to the metabolic burden imposed on the cells by the

oscillatory, critically damped control scheme (entry 2, Table 9).

 

   

16 25
A

A14» ‘ A ‘20
312+ A ’ g,
2?in .152
01 8. E
(D =
g 64 "108
<' 4. 5
c0 2 ..5

0} .0

 

 

0 12 18 24 30 36 42
time (h)

Figure 67. SP1.1/pKD12.112A fed-batch fermentation time course. Glucose limited
conditions, K, = 0.8.
l

A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); ., l QA(g/L).

This result reinforces the theory that QA contamination occurs due to uptake of
SA from the fermentation broth and its in vivo processing to QA. However, use of this
pulsed glucose addition scheme cannot be used for an industrial scale process. Lowering
of the metabolic capacity of the cells resulted in loss of dissolved oxygen control and
unregulated glucose addition, necessitating shutting down of the fermentation after 42 h.
Furthermore, increase in the PID setting could suppress QA formation only in
SP1.1/pKDlZ.112A and had no effect on fermentations involving SP1.1/pKD12.138A

(entry 5, Table 9). This strain takes advantage of transketolase overexpression and is

-110-

capable of producing high titers and yields of SA as described in Chapter 3. However,
QA contamination negates any potential application of SP1.1/pKD12.138A on an
industrial scale. A universally applicable method was therefore sought after for the

suppression of QA synthesis.

HO

HO O
HO

HO
OCH3

methyl-a-D-glucopyranoside

Figure 68. Structure of methyl-a-D-glucopyranoside (MGP).

The compound methyl-a-D-glucopyranoside (MGP) is a non-hydrolyzable
glucose analogue which is known to be a substrate for the PT S system of glucose

‘09“‘0 It has also been shown to exhibit the same catabolite

transport (Figure 67).
repressive tendencies as glucose. MGP undergoes phosphorylation while being
transported across the cell membrane, akin to glucose. However, unlike phosphorylated
glucose, the phosphorylated MGP cannot be catabolized by the cell. It is therefore
dephosphorylated by the cytosolic phosphatases and channeled out of the cell. Export of
free MGP has been postulated to occur as an exchange for phosphorylated MGP.
Extracellular concentrations of MGP would therefore be expected to remain constant and
hence a continuous addition of MGP is not needed.“°" A concentration of 1 mM MGP

was found to be saturating in terms of MGP transport and phosphorylation. The most

important ramification of this process was that high glucose levels were no longer needed

-111-

to maintain a catabolic repressive effect on Shikimic acid uptake. Hence, a normal PID
setting of Kc = 0.1 could be employed, ensuring that the cells remained viable for longer
durations of time.

Strain SP1.1/pKD12.112A was cultured under normal fermentation conditions in
the presence of 1 mM MGP. No quinic acid formation was observed even after 48 h
using a PID setting of Kc = 0.1 (Figure 68). This result is in stark contrast to that
observed when the same strain was cultivated under identical fermentation conditions in
the absence of MGP, when a 3.521 ratio of SA/QA was observed. This method of
suppressing QA synthesis was also more appealing than employing a Kc = 0.8 as
evidenced by the higher SA yield and titer (entry 3, Table 9), as also the improved

dissolved oxygen control up to 48 h.107

 

 

 

 

 

 

 

30

S.

I 20 r»

O A

0)“ 15-

I

0, 1o

(7‘;
L L
0'—‘—r“: . . . .

0 12 18 24 30 3

6 42 48
time (h)

Figure 69. SP1.1/pKD12.112A fed-batch fermentation time course. Glucose limited
conditions, Kc = 0.1, lmM MGP.
A Dry cell wt. (g/L); I SA (g/L); m DHS(g/L); | l QA(g/L).

-112-

Strain SP1.1/pKD12.138A gave a SA/QA ratio of 1.6:1 after 48 h under normal
fed-batch fermentation conditions at a PID setting of K, = 0.1 (Figure 69) (entry 4, Table
9). Increasing the gain to K = 0.8 worsened the situation giving a SA/QA ratio of 1.421
with much lower yields and titers after 42 h (entry 5, Table 9). By contrast, conducting a
fed-batch fermentation with SP1.1/pKD12.138A with a PID setting of KC = 0.1 in the
presence of 1 mM MGP reduced the QA contamination to trace amounts (Figure 70).
After 48 h, this fermentation afforded 34.5 g/L SA in 19% yield with a 14.1:1 ratio of
SA/QA (entry 6, Table 9), thereby allowing the impact of transketolase overexpression

on yield and titer to be realized.'°7

 

 

 

 

 

 

 

30 25
A
A

g 25" a a ‘ -- 20 ,-
.- 20.. . . g
g 1" 15 E-
05 15-- =
g --10 8
.. 10-- b
< 'D
a) 5‘ "5

o- L-.o

36 42 48

 

time (h)

Figure 70. SP1.1/pKD12.138A fed-batch fermentation time course. Glucose limited
conditions, K, = 0.1, 0 mM MGP.
‘ Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); i ' QA(g/L).

-ll3-

b
0
(D
O

 

A35.“- ‘ ‘ ‘ ‘ ‘ ""25
3304 A ,-
E’25_1 +205
0 .
C”'20-- 15%
I
0.15" ~10:
5,10.- 0
5..
0.

 

 

 

0 12 18 24 30 36 42 48
time(h)

Figure 71. SP1.1/pKD12.138A fed-batch fermentation time course. Glucose limited
conditions, K, = 0.1, 1 mM MGP.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); I | QA(g/L).

Table 9. Titers and yields of SA produced under various glucose-limited conditions.

 

 

 

Entry no. Strain Kc [MGP] [SA] (g/L) [GA] (g/l.) [DHS] (97L) SA/ QA SA yield Total yield
(mM) ratio (mol/mol) (mol/mol)
1 SP1.1/pK012.1 12A 0.1 0 25.8 8.1 6.5 35:1 13% 20%
2 SP1.1/pK012.112A 0.8 0 13.7 1.2 4.6 12:1 10% 14%
3 SP1.1/pKD12.112A 0.1 1 27.3 0.0 5.3 - 15% 18%
4 SP1 .1/pKD12.138A 0.1 0 28.0 19.3 10.8 1.621 14% 29%
5 SP1.1/pKD12.138A 0.8 0 11.7 9.4 5.7 1.4:1 8% 18%
6 SP1.1/pKD12.138A 0.1 1 34.5 2.8 8.8 14.1 :1 19% 25%

 

 

-114-

Experimental Evidence Supporting QA Formation Via SA Uptake

As illustrated in Figure 65, there are two possible avenues to consider as sources
of the QA contamination. It could be arising via de novo biosynthesis or it could occur
by uptake of SA from the culture supernatant and its in vivo processing to QA. Both
mechanisms utilize the ability of aroE encoded shikimate dehydrogenase to reduce DHQ
as well as DHS to QA and SA respectively. The fact that lowering the intracellular
concentrations of DHQ by overexpression of aroD encoded DHQ dehydratase had no
effect on QA accumulation weakens the argument in favor of de novo synthesis.
Information gleaned from results discussed earlier in this Chapter, implicate uptake of SA
as the culprit leading to QA formation. Moreover, if QA formation were occurring via de
novo synthesis, then accumulation of some DHQ would be expected. Biocatalysts
designed to biosynthesize QA, namely QP1.1/pKD12.138A, routinely produce DHQ as a
contaminant in fed-batch fermentations. This phenomenon is not observed during SA
producing fermentations.

In order to establish the mode of formation of QA with some degree of confidence
strain SC1.0 was constructed (Figure 71). This biocatalyst was derived from E. coli
RB79lserA also known as JWF 1. The aroL and aroK encoded isozymes of shikimate
kinase were rendered inactive by successive Pl phage-mediated transductions of
aroLzzTnIO and aroKzszR resulting in strain SC1.1. A final P1 phage-mediated
transduction of aroBzzTnS resulted in loss of aroB encoded DHQ synthase activity. P1

phage was propagated from JBS.95b

-115-

Strain construction Selections used for identifying correct phenotype

 

M9/glu M9/glu/ser
RB791 serA' —-— +++
l aroL::Tn1 0
LB/Tc M9/glu/ser
RB791 serA' aroL::Tn10 +++ +++
larchCmR
LB/Tc/Cm M9/glu/ser
RB791 serA' aroL::Tn10 aroK::CmR +++ - — -
laroBz:Tn5
LB/Tc/Cm/Kn
RB791 serA' aroL::Tn10 aroK::CmR aroB::Tn5 +++
$01 .0

Figure 72. Construction and selection of SC1.0.

The overall effect of this operation was that the entire common pathway in SC1.0
was confined between SA and DHQ (Figure 72). Most importantly, this strain lacked the
capability for SA and QA synthesis, which was critical for the success of the
investigation. SA could be manually added to a fermentation culture of SC1.0
transformed with an appropriate plasmid. If any QA formation were to be observed
during the fermentation, it would have to occur via import of SA into the cell and its in

vivo conversion to QA.

AroF
54" area A D A E Arok
+ i» DAHP —XM3-> DHQ -£* DHS i» SA sap
PEP Arol.
lAroE uptakeT

QA SA

Figure 73. The modified shikimate pathway in SC1.0.

-ll6-

Plasmid pKD12.138 was transformed into SC1.0 and fed-batch fermentations
were conducted with this strain. The first fed-batch fermentation performed with
SCl.O/pKD12.138A was under normal conditions with no addition of SA. This run was
used as the control experiment and confirmed that the biocatalyst indeed was not able to
synthesize SA and QA. It did however succeed in accumulating 68 g/L of DAH in the
culture supernatant after 48 h. Observation of DAH is an artifact of the loss of DHQ
synthase activity resulting in formation of DAHP, which gets dephosphorylated and
exported out the cell as DAH.

Once it had been established that SCl.O/pKDlZ.138A was incapable of in vivo
synthesis of SA or QA, the main experiment was proceeded with. SCl.O/pKD12.138A
was cultured under normal fermentation conditions and at 18 h into the run, 10 g of SA
was added to the fermentation. The fermentation was continued for 48 h and aliquots
were removed every 6 h. Analysis of the fermentation samples indicated that 62 g/L of
DAH had been produced. However, no QA was observed and the original amount of SA
added into the fermentation (10 g) was measured to be unchanged. It was quite possible
that the elevated levels of DAH/DAHP being synthesized by the strain interfered with SA
uptake. Production of DAH/DAHP could be minimized by shutting the glucose feed
once the cells attained maximum growth and entered stationary phase of growth.
Depriving the cells from glucose might also serve a dual purpose. In the absence of
glucose the cells would be starved for a carbon source and since SA would most likely be
the only one present in substantial amounts, chances of its uptake would be enhanced.

The fermentation with SCl.O/pKD12.l38A was repeated under normal cultivation

conditions. In this experiment, 10 g of SA was added at 12 h instead of 18 h to provide

-ll7-

the cells sufficient time to uptake the SA. At 18 h, the glucose pump was turned off and
the fermentation was continued for 48 h. Analysis of fermentation samples confirmed
that shutting down the glucose feed did indeed lower DAH/DAHP synthesis dramatically.
Only 7 g/L of DAH was measured after 48 h as opposed to 62 g/L observed in previous
fermentations. However, the negative aspect of the experiment was the complete absence
of any QA formation whatsoever. Attention was also given to the fact that deprivation of
glucose drastically affected the growth of the cells. A maximum dry cell weight of 14
g/L at 24 h was measured for this strain when the glucose pump was shut down at 18 h.
The maximum dry cell weight recorded for SC1.0/pKD12.138A was 24 g/L at 30 h when
the glucose feed was continued with for 48 h. Two likely scenarios therefore emerged to
account for absence of SA/QA equilibration. It was quite likely that the sickness of the
cells caused by lack of glucose prevented uptake of SA. This was indeed a strong
possibility because SA/QA equilibration was normally observed to set in when cell
growth was at peak efficiency. The other possibility was to accept that QA formation
was not a result of SA uptake but due to the presence of a de novo biosynthetic route. A
conclusive interpretation could therefore not be drawn from this body of work.

A close examination of the problems associated with the study discussed earlier,
highlighted the importance of sustaining good biocatalyst growth but without
accumulation of any product which might interfere in the uptake mechanism. An
alternative approach was therefore designed. Since inactivation of DHQ synthase
coupled with overexpression of arol‘“FBR encoded DAHP synthase was responsible for
accumulation of DAH/DAHP, it was decided to instead inactivate DAHP synthase. The

common pathway would then be corralled between DAHP and SA instead of DHQ and

-ll8-

SA (Figure 73). An added advantage of this approach was that SP1.1 could be used as
the host strain, since the native E. coli DAHP synthase isozymes are feedback inhibited
by the aromatic amino acids. Regular addition of aromatic amino acids during the
fermentation would therefore ensure suppression of genomic DAHP synthase. Glucose
addition could therefore be continued throughout the fermentation period thereby
maintaining healthy cell growth without accumulation of any product. The plasmid
required for the investigation would have to incorporate all the genes present in
pKD12.138A except for aroFFBR, which encodes for feedback insensitive DAHP

synthase.

AroF

E4P } {—A'OG
E k
+ AroH DAHP Aro§l DHQ AroD DHS AroE SA Aro S3P

PEP AroL
lAroE ”MST

QA SA

Figure 74. Effect of inactivating DAHP synthase in SP1.1.

The aroFFBR locus in pKD12.138A possesses a BglII restriction site which is a
unique site in the plasmid. Digestion of pKDlZ.l38A with BglII resulted in a linear
plasmid which was treated with Klenow fragment. Ligation of the linear plasmid
afforded pSC5.214A, which was identical to pKDlZ.138A in all respects except for the
presence of the inactive arol‘i‘FBR gene (Figure 74). Plasmid pSC5.214A was transformed
into SP1.1 and this strain was used in the fermentation experiments designed to decipher

the mode of formation of QA.

-ll9-

  
  

   

pKD12.138A aroE
9.9-kb

Ncol

(Hindlll)

(Hindlll) (Smal)

ii) Klenow treatment

i) Bglll digest
iii) Ligation with T4 ligase

ECORI

pSCS.214A aroE
9.9—kb

Ncol

(Hindi!)

(Hinolll) (Smal)

Figure 75. Preparation of Plasmid pSC5.214A.

-120-

The first fed-batch fermentation performed with SP1.1/pSC5.214A was under
normal conditions and provided information as to its production capabilities. After 48 h,
accumulation of 24 g/L SA and 3.5 g/L DHS was recorded (Figure 75). No QA
production was observed. No additional aromatic amino acids were added apart from the
initial dosage required to sustain cell growth and hence native DAHP synthase remained

active during the fermentation.

 

 

 

 

 

4o 25
A 35 ~- A ‘
a 30 -- g
I; 25 T ‘7
O
(D. 20 “P E
76

I
D 15 it o
<' L ‘ 5
(D 10 4

5 e

o ‘ . . . l .

0 12 18 24 30 36 42 48
time (h)

Figure 76. SP1.1/pSC5.214A fed-batch fermentation time course. Glucose limited
conditions, no extra aromatic amino acids added.

A Dry cellwt.(g/L); - SA (g/L); m DHS(g/L); l f i QA(g/L).

The next experiment conducted with SP1.1/pSC5.214A was to observe the effects
of maintaining a finite concentration of aromatic amino acids in the fermentation at all
times. Apart from the aromatic amino acids added at 0 h for normal cell growth, 0.7 g
phenylalanine, 0.7 g tyrosine, and 0.35 g tryptophan were added at 18 h and 30 h into the
fermentation. After 48 h, analysis of the fermentation samples indicated that no SA,

DHS or QA were produced implying the complete suppression of native DAHP synthase.

-121-

It was also worth noting that the cells exhibited robust growth and hence the problems
associated with SCl.O/pKDl2.l38A could be avoided.

SP1.1/pSC5.214A was cultured under fed-batch fermentation conditions identical
to those described above with the periodic addition of aromatic amino acids. The main
difference was the addition of 10 g SA at 12 h into the fermentation. No QA acid
formation was observed until 24 h into the experiment. Traces of QA were then
measured to have been formed between 24 h and 30 h followed by a sudden burst in QA
production with a concomitant depletion in SA amounts between 30 h and 36 h. After
48 h, a final SA/QA ratio of 0.76:1 was measured providing irrefutable evidence that
involvement of SA uptake and its in vivo processing was responsible for the formation of

QA. In addition, 2.5 g/L of DHS was also recorded to have been formed after 48 h.

 

 

 

 

 

 

 

 

8 35

7J- ‘ ‘ A ]- 30
A 6.,. A
S, -- 25
IE 5* ‘ -- 20 3
00).. 41. A E
E 3i “15 8
(.- 2. A "10 '5
U) .

1-- --5

o A . .‘ . . o

0 12 18 24 30 36 42 48

time (h)

Figure 77. SP1.1/pSC5.214A fed-batch fermentation time course. Glucose limited
conditions; aromatic amino acids added at 0 h, 18 h and 30 h; 10 g SA added at 12 h.

A Dry ceuwt (em; I SA (g/L); m DHS(g/L); l IQA(g/L).

-122-

Establishment of the role of SA uptake in the formation of QA provided an
avenue to explore which could lead to elimination of SA/QA equilibration. Extensive
research has been conducted on identifying the genetic basis for SA uptake. In 1998,
Pittard and co-workers cloned and sequenced a 1.3—kb DNA fragment labeled shiA,
which was shown to be responsible for the ability of E. coli to import SA.'” Introduction
of a mutation into the shiA locus rendered the cell unable to transport SA.

The strain JP11123 carrying the .i'hiAzzKanR locus had been previously constructed

' P1 phage-mediated transduction of shiAzzKanR from

by Pittard and co-workers.”
JP11123 into SP1.1 resulted in SP1.1shiA which should now be unable to transport SA
and hence be incapable of producing QA. Plasmid pKDl2.138A was transformed into
SP1.lshiA and the resulting biocatalyst SP1.1shiA/pSC5.214A was tested under normal
fed-batch fermentation conditions. As described earlier, at 0 h, 18 h, and 30 h into the
fermentation, 0.7 g phenylalanine, 0.7 g tyrosine, and 0.35 g tryptophan were added to
inhibit native DAHP synthase. 10 g SA was added to the culture at 12 h into the
fermentation. The fermentation was allowed to complete 48 h and the samples analyzed.
Knocking out the shiA locus in SP1.1 had no effect on SA/QA equilibration (Figure 78)
and the result obtained was essentially the same as that observed with SP1.1/pSC5.214A.
Since this strain lacks the capability to synthesize QA but production of this compound is

still observed, the most likely explanation is that there may be multiple proteins

responsible for SA uptake.

-123-

 

 

 

 

 

 

9 35

8 “30
a“ ‘ -
:6 “253
0 5 20;
3’3 4 --15§
0.3 a
< “10.:3
a) 2

1 "5

o 4.0

48

 

Figure 78. SP1.1shiA/pSC5.214A fed-batch fermentation time course. Glucose
limited conditions; aromatic amino acids added at 0 h, 18 h and 30 h; 10 g SA added
at 12 h.

A Dry cell wt. (g/L); I SA (g/L); m DHS(g/L); l l QA(g/L).

Comparison was also made between SP1.1/pKD12.138A and
SP1.13hiA/pKD12. 138A under fed-batch fermentation conditions with a Kc = 0.1 setting,
in the absence of MGP. These conditions produced 28 g/L SA and 19.3 g/L QA in a
molar ratio of 1.6:] with SP1.1/pKDlZ.138A (Figure 69). Introduction of the shiA
phenotype gave no discemable advantage and in fact, SP1.1shiA/pKDl2.l38A gave a

worse SA/QA ratio of 1.1:1 (Figure 79).

-124-

 

A 20 ‘ wi- 25
_J A j
E ‘ (e 20 El
E§ 1551 AF‘ ‘ E;
- 15 _
(i) 10» 7’
0 A --10 ‘3’.
<f 'o

(D 5.],- 5

0 0

 

 

 

 

 

 

 

 

 

 

$4., , , 1 ,
0 12 18 24 30 36 42 48
time (h)

Figure 79. SP1.1shiA/pKDlZ.l38A fed-batch fermentation time course. Glucose
limited conditions, K, = 0.1, 0 mM MGP.
A Dry cell wt. (g/L); - SA (g/L); m DHS(g/L); i l QA(g/L).

Discussion

Before the emergence of glucose-rich fed-batch fermentor conditions described in
Chapter 3 of this thesis, biocatalysts were routinely cultured under conditions limited in
glucose. A resulting manifestation of these conditions was the observation of QA being
co-synthesized along with SA. Normal fed-batch fermentations were run by maintaining
the glucose feed pump proportional-integral-derivative (PID) gain setting of Kc = 0.1.
The ramification of this value on the fermentation conditions was that the glucose
concentration throughout the fermentation period, was just sufficient to sustain healthy
growth of the cells and ensure a 10% dissolved oxygen value. The input of glucose into
the fermentation was exactly matched by its consumption by the cells and at no given

time could excess glucose be measured to be present in the culture supernatant. When a

-125-

PID gain setting of K = 0.8 was adopted, the only experimental parameter being changed
was the rate of glucose feeding. Instead of a steady feeding pattern, a pulsed glucose
addition was assumed resulting in a pulsed dissolved oxygen profile. The overall effect
was to maintain a higher effective concentration of glucose in the fermentation, which
translated into disappearance of QA formation. The disappearance of QA formation
hinted at the involvement of an uptake phenomenon, which was now catabolically
repressed by the increased glucose levels.

Suppression of QA production via the utilization of a PID setting of Kc = 0.8 came
with its share of penalties. The pulsed feeding regimen caused early loss of dissolved
oxygen control resulting in unregulated addition of glucose, forcing shutdown of the
fermentation at 42 h. The approach was found to succeed only for SP1.1/pKDlZ.112A
and not be viable for strains capable of producing higher amounts of SA namely,
SP1.1/pKD12.138A. In addition, the feeding regimen was detrimental to the health of
the cells affecting yields and titers of SA. Hence this method was not considered as the
ideal route to pursue and alternative options were considered.

The use of MGP was based on the notion that at K. = 0.8, glucose induced
catabolite repression prevented uptake of SA, thereby accounting for the disappearance of
QA. Various reports have acknowledged the ability of MGP to rrrimic glucose in various
situations including its ability to effect catabolite repression on non-carbohydrate
substrates. Use of MGP would permit use of normal fermentation conditions with a PID
setting of K = 0.1, thereby ensuring normal cell growth and higher SA titers. An
addition of 1 mM MGP at the beginning of the fermentation was found to be sufficient to

last the entire duration of the fermentation. Strains SP1.1/pKD12.112A and

-126-

SPl.l/pKDl2.138A were tested under these conditions and both gave positive results.
Good dissolved oxygen control was maintained throughout the fermentation and at the
end of 48 h no QA was detected in the culture supernatant. Apart from the fact that both
strains produced SA free from QA contamination, it was important to acknowledge that
they did so in reasonable titers and yields. The observation that maintaining a finite
carbohydrate concentration was key to suppressing QA formation lent credence to the
theory of SA uptake being the source of the problem. These results also represented the
first instance of fermentation conditions being available for industrial production of SA.

Research conducted to this point provided results consistent with QA synthesis
occurring due to import of SA by the cell followed by in vivo processing via the common
pathway. They by no means discounted the possibility that QA could also arise by de
novo biosynthesis. It was therefore important to test for accumulation of QA within a
system in which the capability for de novo synthesis of QA was not functional. The
biocatalyst SP1.1/pSC5.214A perfectly suited these requirements and provided a means
to lock the common pathway between DAHP and SA with no possibility for de novo
synthesis. Under normal fermentation conditions, SP1.1/pSC5.214 was able to exhibit
SA/QA equilibration when the only possibility for QA formation was via uptake of added
SA from the culture medium.

SA transport has for long been hypothesized to be mediated by multiple loci in the
E. coli genome.'°8‘”' Recently the major locus purported to be responsible for SA uptake
named shiA has been cloned and sequenced.”' The KanR gene was inserted within the
shiA locus and the resulting shiAzzKanR fragment was introduced into the E. coli genome

by homologous recombination. The resultant strain JP11123 exhibited a marked inability

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to import SA from the external medium as compared to the parent strain AB2826.
Restoration of SA uptake capability in JP11123 was observed upon introduction of a
plasmid carrying wild-type shiA.

Transduction of the shiA::KanR locus into SP1.1 was intended to provide a strain
devoid of any SA/QA equilibration tendencies. However, the ability of
SP1.13hiA/pSC5.214 to produce both QA and DHS from exogenously supplied SA
contradicts the notion that mutants deficient in shiA activity are unable to transport SA.
However, as stated before, multiple loci have been held responsible for SA transport in
the past. Evidence for the shiA' phenotype of JP11123 was obtained by monitoring for
uptake of [”C]-SA. These experiments were performed for 90 seconds at cell densities of
approximately 0.11 g/L.”' On the other hand, fermentations performed with
SP1.1shiA/pSC5.214A were run for 48 h and SA/QA equilibration was observed only
after 36 h at cell densities of approximately 25 g/L. The longer fermentation times and
greater then ZOO-fold cell density could very well allow for sufficient amplification of
residual uptake activity resulting in SA/QA equilibration. It is therefore entirely possible
that residual SA uptake activity associated with the remaining loci might be responsible
for the results observed with SP1.1shiA/pSC5.214A.

Even though use of fermentation conditions described in Chapter 2 and 3 have
now been successfully implemented to circumvent the problem of SA/QA equilibration,
the fact remains that the problem still persists at a genetic level. Another problem that
hasnt received sufficient attention is that of DHS accumulation. Even under glucose-rich
conditions, amounts of DHS accumulating in SA fermentations are quite substantial (15-

20 g/L, Table 4, Chapter 3). Part of the problem lies in the feedback inhibition of

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shikimate dehydrogenase by SA. But at least some fault rests on the import of SA by the
cell and its conversion to DHS by the reverse reaction of shikimate dehydrogenase.
Attenuation of this problem could result in the ideal situation of a biocatalyst

synthesizing pure SA devoid of major contaminants in high titers and yields.

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CHAPTER 5

EXPERIMENTAL

General Methods

General Chemistry
All reactions sensitive to air and moisture were carried out in oven and/or flame
dried glassware under positive argon pressure. Air or moisture sensitive reagents and
solvents were transferred to reaction flasks fitted with rubber septa via syringes or
cannula. Unless otherwise specified, all reactions were carried out at room temperature.
Solvents were removed using either a Bttchi rotary evaporator at water aspirator pressure
or under high vacuum (0.5 mm Hg). Hydrogenation was performed on a Parr

hydrogenation apparatus under 50 psi of hydrogen.

Reagents and Solvents
HMPA was dried and stored over activated Linde 4 A molecular sieves under
nitrogen. CHzCl2 and Et3N were distilled from calcium hydride under nitrogen. THF was
distilled under nitrogen from sodium benzophenone ketyl. Water used in purifications
was glass distilled deionized. All other reagents and solvents were used as available from
commercial sources or purified according to published procedure. Organic solutions of

products were dried over anhydrous MgSO4.

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Chromatography

Radial chromatography was carried out with a Harrison Associates Chromatotron
using 1, 2 or 4 mm layers of Silica gel 60 PF254 containing gypsum (E. Merck). Silica gel
60 (40-63 pm, E. Merck) was used for flash chromatography. Analytical thin—layer
chromatography (TLC) utilized precoated plates of silica gel 60 F-254 (0.25 mm,
Whatman). TLC plates were visualized by immersion in anisaldehyde stain (by volume:
95% ethanol, 3.5% sulfuric acid, 1% acetic acid and 2.5 % anisaldehyde) followed by
heating.

Diethylaminoethyl cellulose (DEAE) was purchased from Whatman, Dowex 50
(H*) from Sigma, hydroxylapatite (HA) from Bio-Rad, and Dymatrex Red A from
Amicon. Depending on its use, DEAE was recycled and cleaned with either
1M Et3NH”HCO3' (TEAB) or 0.5 M KC1 followed by exhaustive washing with water.
TEAB buffer was prepared by bubbling CO2 gas through an aqueous solution of Et3N
until the pH reached 7.5 at 0 °C. Dowex 50 (H*) was cleaned before use by treatment
with bromine at pH 14, extensive rinsing with water followed by washing with
6N aqueous hydrochloric acid, and finally washing with water until the pH of the eluent

tested neutral. All aqueous chromatographic purifications were carried out at 4 °C

Spectroscopic and Analytical Measurements
1H NMR and 13C NMR spectra were recorded on a Varian VX-300 FT-NMR
spectrometer. Chemical shifts for lH NMR spectra are reported in parts per million
(ppm) relative to internal tetramethyl silane (Me4Si, 8 = 0.0 ppm) with CDCl3 as the

solvent, and to sodium 3-(trimethylsilyl) propionate-—2,2,3,3-d4 (TSP, 6 = 0.0 ppm) when

-l3l-

D20 was the solvent. The following abbreviations are used to describe spin multiplicity:
s (singlet), d (doublet), t (triplet), m (unresolved multiplet) and dd (doublet of doublets).
Chemical shifts for 13C NMR spectra are reported in ppm relative to CDCl;,
(8 = 77.0 ppm) or internal standard acetonitrile (CH3CN, 5 = 3.69 ppm) in D20. Fast
atom bombardment (FAB) mass spectra were obtained on a double-focusing Kratos
MS50 mass spectrometer employing glycerol as the matrix. Elemental analysis were
performed by Atlantic Microlab Inc. (Norcross, GA). UV and visible spectra were
recorded on a Perkin-Elmer Lambda 3B spectrometer or a Hewlett Packard 8452A diode

array spectrometer.

Enzyme Assays

General Information

3-deoxy-D-arabino-heptulosonic acid (DAH) was biosynthesized using the E. coli
strain AB2848/pKLl.87A/pMF52A. The supernatant from 1 L of culture was passed
through 500 mL Dowex 50 (Hi) and the clear eluent adjusted to pH 8.0 using freshly
prepared 1 M lithium hydroxide. After the solution was concentrated down to dryness,
the oily residue was stirred with 1 L methanol at 4 °C for l h. The white precipitate was
filtered off and the filtrate concentrated under reduced pressure. The residue was
dissolved in water, passed down Dowex 50 (H‘) and the eluent concentrated to dryness to
give DAH. The conversion of DAH to 3-deoxy-D-arabino-heptulosonic acid 7-
phosphate (DAHP) followed the same protocol as reported by Frost and Knowles.”2

DHQ synthase was purified in the Co*2 form using E. coli RB791/pJB14

according the procedure of Frost and co-workers.“3 Purification of DHQ synthase in the

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Zn+2 form followed the same procedure but with replacement of CoCl2 in all the buffers
with 75 11M ZnSO4.7HZO. DHQ dehydratase was purified from E. coli AB2848/pKDZOl
according to the protocol established by Chaudhuri and co-workers.86 Concentrations of
phosphorous containing compounds were determined by the method developed by Avila
and Ames.“" Protein solutions were concentrated by ultrafiltration using either PM-lO
Diaflo membranes or Centricon concentrators from Amicon.

Cell lysis was achieved by two passes through a French pressure cell (SLM
Aminco) at 16000 psi and the cellular debris was separated from the lysate by
centrifugation (48000 x g, 20 min, 4 °C). Protein concentration was quantified using the

Bradford dye-binding procedure with assay solution from Bio-Rad.

DAHP synthase assay

DAHP synthase activities were measured using the procedure reported by
Schoner.“5 Harvested cells were resuspended in SOmM potassium phosphate (pH 6.5)
containing 10mM PEP and 0.05mM CoClz. After the cells were disrupted by passage
through a French press, the lysate was diluted in a solution of 50 mM potassium
phosphate (pH 7.0), 0.5 mM PEP and 250 mM 1,3-propanediol. A dilute solution of E4P
was concentrated to 12 mM and the pH was adjusted to 7.0 with KOH. Two solutions
were prepared and incubated separately at 37 °C for 5 min. The first solution (1 mL, pH
7.0) contained 6 mM E4P, 12 mM PEP, 1 mg ovalbumin and 25 mM potassium
phosphate. The second solution was 0.5 mL of diluted lysate. The two solutions were
mixed (time = 0) and incubated at 37 °C. At regular time intervals, 0.15 mL samples

were removed from the reaction mixture and quenched with 0.1 mL of 10%

-l33-

 

trichloroacetic acid (w/v). Precipitated protein was removed by centrifugation and the
DAHP level in each sample was determined using the thiobarbiturate assay.

To a 0.1 mL aliquot of DAHP containing sample was added 0.1 mL of 0.2 M
NaIO4 in 8.2 M H3PO4 and the mixture incubated at 37 °C for 5 min. The reaction was
quenched by addition of 0.8 M NaAsO2 in 0.5 M NaZSO4 and 0.1 M H2804 (0.5 mL) and
vortexed until a dark brown color appeared. This solution was finally treated with 3 mL
of 0.04 M thiobarbituric acid in 0.5 M NaZSO4 (pH 7.0) and heated at 100 °C for 15 min.
The samples were cooled and the pink color extracted into 4 mL of cyclohexanone. The
aqueous and organic layers were separated by centrifugation (2000 x g, 15 min) and the
absorbance of the organic layer measured at 549 nm (e = 68000 L mol’1 cm“). One unit
of DAHP synthase activity was defined as the formation of 1 umol of DAHP per min at

37 °C.

Transketolase activity measurement

Transketolase activity was assayed using the method of Paoletti.69a Harvested
cells were resuspended in 50 mM potassium phosphate (pH 7.5), 1 mM MgC12, and 0.2
mM DTT and subjected to lysis using a French press as described earlier. A dilute
solution of E4P was concentrated to 10 mM and the pH adjusted to 7.0 with KOH. The
assay mixture (1 mL) contained 200 mM triethanolamine (pH7.6), 5 mM MgC12, 0.1 mM
thiamine pyrophosphate, 0.4 mM NADP, 0.4 mM B-hydroxypyruvate, 0.2 mM E4P, 8
units glucose-6-phosphate dehydrogenase, and 4 units phosphoglucose isomerase. The
solution was allowed to equilibrate at room temperature for 5 min and the absorbance at

340 nm was monitored for several minutes. After all the unreacted D-glucose 6-

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phosphate from the E4? synthesis had reacted, an aliquot of the diluted lysate was added
and the reaction monitored at 340 nm for 20-30 min. One unit of tranketolase activity is

defined as the formation of 1 umol of NADP (e = 6220 L mol'l cm") per min.

Bacterial Strains
E. coli DHSOt [F ’ endAI hst17(r‘Km*K) supE44 (hi-1 recAI gyrA relAI
0801acZAM15 A(lacZYA-argF)U,69], RB791 (W3110 lacL81") and JWFl (RB791 serA)
were obtained previously in this laboratory. E. coli B was purchased from ATCC.
JP11123 (thi-I argE3 his-4 proAZ galK2 lacYI tsk29 aroD362 ~shiA354zzKanR) was
obtained from the laboratory of Professor J Pittard. TP2811 (F xyl argHI AlacX 74 aroB
ilvA A(ptsH ptsI crr)::KanR) was obtained from Sophie Lévy at the Institut Pasteur,

France. Plasmid pTC325 was obtained from Professor L. Ingram.

Storage of Bacterial Strains and Plasmids
All bacterial strains were stored at -78 °C in glycerol. Plasmids were transformed
into either DH50t or JWFl for Storage. Glycerol freezes were prepared by adding
0.75 mL of culture to 0.25 mL of sterile 80% (v/v) glycerol. The solutions were mixed,

allowed to stand at room temperature for 2 h, and then stored at -78 °C.

Culture Medium

All solutions were prepared in distilled, deionized water. LB medium contained (1
L) Bacto tryptone (10 g), Bacto yeast extract (5 g), and NaCl (10 g). M9 salts (1 L)

contained NazHPO4 (6 g). KHZPO4 (3 g). NH4C1 (1 g). and NaCl (0.5 g). M9 minimal

-l3S—

medium contained D-glucose (10 g), MgSO4 (0.12 g), and thiamine hydrochloride (0.001
g) in l L of M9 salts. M9 medium (1 L) was supplemented where appropriate with L-
phenylalanine, L-tyrosine, L-tryptophan, and L-serine to a final concentration of 40 ug/mL
and with p-hydroxybenzoic acid, potassium p-aminobenzoate, and 2,3-dihydroxybenzoic
acid to a final concentration of 10 pg/mL. Antibiotics were added where required to the
following final concentrations: chloramphenicol (Cm), 20 ug/mL; ampicillin (Ap), 50
ug/mL; and kanamycin (Kan), 50 ug/mL. Inorganic salts, D-glucose, and MgSO4
solutions were autoclaved separately while thiamine hydrochloride, amino acids,
aromatic vitamins, casamino acids, and antibiotic stock solutions were sterilized through
0.22-um membranes. Solid medium was prepared by addition of 1.5% (w/v) Difco agar
to medium solution.

Fermentation medium (1L) contained KZHPO4 (7.5 g), citric acid monohydrate
(2.1g), ammonium iron (III) citrate (0.3g), concentrated H2804 (1.2 mL), L-tyrosine
(0.7 g), L-phenylalanine (0.7 g) and L-tryptophan (0.35g). The pH of the medium was
adjusted to 7.0 with concentrated NH4OH prior to autoclaving. The following
supplements were added to the medium immediately prior to initiation of the
fermentation: D-glucose (20 g or 30 g), MgSO4 (0.24 g), p-hydroxybenzoic acid (0.01 g),
2,3-dihydroxybenzoic acid (0.01 g), potassium p-aminobenzoate (0.01 g) and trace
minerals including (NH4)6(M07024)-4HZO (0.0037 g), ZnSO4-7HZO (0.0029 g), H3BO3
(0.0247 g), CuSO4-5HZO (0.0025 g) and MnC12-4H20 (0.0158 g). Solutions of D-glucose
and MgSO4 were autoclaved separately while aromatic vitamins and trace minerals

solutions were sterilized through 0.22 pm membranes.

-l36-

 

General Fed-Batch Fermentor Conditions

Fermentations were conducted in a B. Braun M2 culture vessel with a 2 L
working capacity. The vessel was modified using a stainless steel baffle cage containing
four 1/ " x 4" baffles for all fermentations except those using KL3/pKL4.l30B.
Environmental conditions were supplied by a B. Braun Biostat MD controlled by a DCU-
1. Data was acquired on a Dell Optiplex Gs+ 5166M personal computer utilizing B.
Braun MFCS/W in software. PID control loops were used to control temperature, pH, and
glucose addition. The temperature was maintained at 33 °C, and the pH was maintained
at 7.0 by addition of NH4OH or 2 N H2804. Glucose was added as a 60% (w/v) solution
for all Strains. Dissolved oxygen (D.O.) was monitored using a Mettler-Toledo 12 mm
sterilizable O2 sensor fitted with an Ingold A-type O2 permeable membrane. D.O. was
maintained at 10% air saturation throughout the course of the fermentations. Antifoam
(Sigma 204) was pumped into the vessel manually as needed.

Each inoculant was initiated by introduction of a single colony into 5 mL of M9
medium and grown at 37 °C with agitation for 12-24 h until the culture was turbid. After
this time, the starter culture was transferred to 100 mL of M9 medium and grown for an
additional 12 to 24 h at 37 °C and 250 rpm. After an appropriate ODf,00 was reached (1.0-
3.0), the inoculant was transferred to the fermentation vessel. The initial glucose
concentration in the fermentation was 30 g/L for all fermentations discussed in Chapter 3,
and 20 g/L for those described in Chapter 4. Three staged methods were used to maintain
D.O. levels at 10% air saturation during the course of the run. With the airflow at an
initial setting of 0.06 LIL/min, D.O. concentration was maintained by increasing the

impeller speed from its initial set point of 50 rpm to its preset maximum of 750 rpm.

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When the impeller reached its preset maximum, the mass flow controller then maintained
D.O. levels by increasing the airflow rate from 0.06 LIL/min to a preset maximum of 1.0
LIL/min. Once the air flow rate leveled off at 1.0 L/L/min at a constant impeller speed of
750 rpm, glucose was manually pumped into the fermentor so as to maintain an average
glucose concentration of 25 g/L throughout the run. At this stage, the DO. level was
maintained at 10% by allowing the impeller speed to vary from 750 rpm to 1600 rpm.
Fermentations were typically run for 60 h.

The fermentations in Chapter 4 did not employ the baffle cage and were run under
Slightly different conditions. During the first two stages the stirrer was allowed to ramp
up to 940 rpm and the air flow up to 1.0 LIL/min. At this stage dissolved oxygen levels
were maintained for the duration of the fermentation at 10% by oxygen sensor-controlled
glucose feeding. Under these conditions, at no stage during the fermentation was there
any excess glucose present in the fermentation culture. Fermentations were typically run

for 42-48 h.

Analysis of Fermentation broth
Samples (5 mL) of fermentation broth were taken at indicated intervals. Cell
densities were determined by dilution of fermentation broth with water (1:100) followed
by measurement of absorption at 600 nm (ODwo). Dry cell weight (g/L) was obtained
using a conversion coefficient of 0.43 g/L/OD600 for RB791-derived strains and 0.29
g/L/OD600 for strains derived from E. coli B. The remaining fermentation broth was
centrifuged using a Beckman microcentrifuge to obtain cell-free broth. Solute

concentrations in the cell-free broth were determined by 1H NMR (Figure 80). For lH

-l38-

 

NMR quantitation of solute concentrations, solutions were concentrated to dryness under
reduced pressure, concentrated to dryness on additional time from D20, and then
redissolved in D20 containing a known concentration of the sodium salt of 3-
(trimethylsilyl)propionic—2,2,3,3-d4 acid (TSP) purchased from Lancaster Synthesis Inc.
Concentrations were determined by comparison of the integrals corresponding to each
compound with the integral corresponding to TSP (8 = 0.00 ppm) in the 1H NMR.
Compounds were quantified using the following resonances: Shikimic acid (6 4.4, m, l
H) (Figure 81), DHS (8 4.3, d, 1 H) (Figure 82), quinic acid (8 4.2, m, 1 H) (Figure 83).
Concentrations of Shikimic acid and quinic acid derived from their respective 1H NMR
integral values tended to be overestimated and precise concentrations of Shikimic acid
and quinic acid were calculated by application of the following formulae: [SA]actual =
0.67523 x [SA]NMR + 0.26658; [QMacml = 0.65887 x [QA]NMR — 0.48161. These
equations were obtained by dissolving various known amounts of Shikimic acid or quinic
acid in 1 mL of D20 containing 10 mM TSP and recording their 1H NMR spectra. The
concentration of SA or QA in each sample that was estimated from 1H NMR, was plotted
against the known concentration of Shikimic acid or quinic acid for that sample resulting
in the calibration curve. All ]H NMR spectra were recorded on a Varian VXR-300 FF-

NMR spectrometer (300 MHz).

-l39-

 

 

 

 

 

in

 

 

 

 

 

 

 

 

Figure 80. Crude 1H NMR of fermentation broth.

-140-

 

A

' " T 7

T 1' T T'Y’j

T— Tj—T 1'

 

 

 

Figure 81. 1H NMR of Shikimic acid.

 

-141-

 

. A.-

 

 

 

41.)

r j— T 1'— l j

 

V’ T T

5‘1

 

PM

ka
pn-

 

v7 v

 

 

 

 

 

 

 

Figure 82. 1H NMR of 3-dehydroshikimic acid.

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Jr-
;.

 

 

JL-

 

 

 

 

 

 

 

 

 

 

Figure 83. 1H NMR of quinic acid.

-l43-

 

I T fl I Y

 

Genetic Manipulations

General Information

Recombinant DNA manipulations were performed according to procedures
outlined in Sambrook and co-workers.116 Restriction endonucleases were purchased from
Gibco BRL and New England Biolabs. T4 DNA ligase, Klenow fragment and dNTP’s
were purchased from Gibco BRL while calf intestinal alkaline phosphatase was
purchased from Roche Molecular Biochemicals. Depending on the selection available, E.
coli DH50t or JWFl were used for all plasmid constructions. Electrophoresis grade
agarose was purchased from Gibco BRL. Prior to use, phenol was distilled and mixed
with 0.1% (w/v) 8-hydroxyquinoline. This was followed by extraction with 1 M Tris-
HCl (pH 8.0) (twice) and finally extraction with 0.1 M Tris-HCl (pH 8.0) until the pH of
the aqueous layer was greater than 7.6. Phenol was stored under an equal volume of 0.1
M Tris-HCl (pH 8.0) at 4 °C. SEVAG was prepared by mixing chloroform and isoamyl
alcohol (24:1 v/v). TE buffer contained 10 mM Tris-HCl (pH 8.0) and lmM NazEDTA
(pH 8.0). Endostop solution (10X concentrated) was made up of 50% glycerol (v/v), 0.1
M NazEDTA, pH 7.5, 1% sodium dodecyl sulfate (SDS) (w/v), 0.1% bromophenol blue
(w/v) and 0.1% xylene cyanole FF (w/v) and stored at 4 °C. Prior to use, 1 mL of 10X
Endostop was mixed with 0.12 mL of DNAase-free RNAase. DNAase-free RNAase was
prepared by dissolving 10 mg RNAase in lmL of 10mM Tris-HCl (pH 7.5) and 15 mM
NaCl. The solution was heated at 100 °C for 15 min to inactivate the DNAase and stored

at -20 °C.

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Large Scale Purification of Plasmid DNA

Purification of plasmid DNA on a large scale followed a modified alkaline
alkaline lysis procedure described in Sambrook and co-workers.“6 A 2 L Erlenmeyer
flask containing 500 mL of LB with appropriate antibiotics, was inoculated with a single
colony and the culture was shaken (250 rpm) at 37 °C for 14 h. In cases where the
plasmid did not possess an antibiotic marker, 500 mL of M9 minimal medium was used
and the culture grown for 24 h under the conditions described earlier. The cells were
harvested by centrifugation (4000 x g, 5 min, 4 °C) and resuspended in 10 mL of cold
GETL solution [50 mM glucose, 20 mM Tris-HCl (pH 8.0), 10 mM NazEDTA (pH 8.0)]
into which 50 mg of lysozyme had been added immediately prior to use. The suspension
was kept at room temperature for 5 min after which it was treated with 20 mL of 1 %
sodium dodecyl sulfate (w/v) in 0.2 N NaOH. The thick suspension was gently mixed,
kept on ice for 15 min and then treated with 15 mL of cold 3 M KOAc (prepared by
mixing 60 mL of 5 M KOAc, 11.5 mL of glacial acetic acid and 28.5 mL of water). The
mixture was shaken vigorously resulting in a thick white precipitate. The sample was
stored for 10 min on ice. The cellular debris was removed by centrifugation (48000 x g,
20 min, 4 °C) and the supernatant transferred equally to two clean centrifuge tubes.
Isopropanol (0.6 volume) was added to each tube and they were kept at room temperature
for 15 min. The precipitated DNA was recovered by centrifugation (20000 x g, 20 min, 4
°C), the supernatant discarded and the DNA pellet carefully rinsed with 70% ethanol.

The DNA pellet was dried, dissolved in 3 mL TE and transferred to a 15 mL
Corex tube. To this solution was added 3 mL of cold 5 M LiCl and after thorough

mixing, the high molecular weight RNA was removed by centrifugation (12000 x g, 10

-145-

nrin, 4 oC). The clear supernatant was transferred to a fresh Corex tube and treated with
6 mL of isopropanol followed by gentle mixing. The precipitated DNA was collected by
centrifugation (12000 x g, 10 min, 4 °C), rinsed with 70% ethanol and dried. The dried
DNA pellet was dissolved in 0.5 mL of TE containing 10 pg of DNAase-free RNAase,
transferred to a 1.5 mL microcentrifuge tube and stored at room temperature for 30 min.
To this solution was added 0.5 mL of 13% PEG-8000 (w/v) in 1.6 M NaCl. The solution
was mixed well and centrifuged (microcentrifuge, 5 min, 4 °C) to recover the precipitated
DNA. The supernatant was discarded and the pellet dissolved in 0.4 mL of TE. The
sample was sequentially extracted with phenol (0.4 mL), phenol and SEVAG (0.4 mL
each) and finally SEVAG (0.4 mL). After the extraction was complete, the aqueous
portion was mixed with 0.1 mL of 10 M NH,OAc followed by addition of 1 mL of 95%
ethanol. The sample was mixed well and left at room temperature for 5 min resulting in
precipitation of DNA. After recovery of the DNA by centrifugation (microcentrifuge,
5 min, 4 °C), the pellet was dissolved in 0.2-0.4 mL of TE.

In order to determine the DNA concentration, an aliquot (10 uL) of the DNA
solution was diluted to 1 mL in TE and the absorbance at 260 nm was measured relative
to TB. DNA concentration was then calculated based on the fact that the absorbance at

260 nm of 50 ug mL'1 of plasmid DNA is 1.0.

Small Scale Purification of Plasmid DNA
A Single colony was inoculated into 5 mL of LB containing appropriate
antibiotics and grown overnight at 37 °C. In cases where the plasmid did not possess an

antibiotic marker, 5 mL of M9 minimal medium was used and the culture grown for 24 h.

-146-

Cells were harvested from 3 mL of culture in a 1.5 mL microcentrifuge tube by
centrifugation. The cell pellet was resuspended in 0.1 mL of cold GETL solution into
which lysozyme (5 mg mL“) was added immediately before use. The suspension was
incubated on ice for 10 min and was then treated with 0.2 mL of 1 % sodium dodecyl
sulfate (w/v) in 0.2 N NaOH. The mixture was shaken gently and kept on ice for 5-10
min. To this sample was added 0.15 mL of cold 3 M KOAc solution and the mixture
shaken vigorously resulting in formation of a thick white precipitate. The sample was
stored on ice for 5 min following which the precipitate was removed by centrifugation
(microcentrifuge, 20 min, 4 °C). The supernatant was transferred to a fresh
microcentrifuge tube and extracted with phenol and SEVAG (0.25 mL each). The
aqueous layer was transferred to a fresh microfuge tube and mixed well with 1 mL of
95% ethanol. The mixture was kept at room temperature for 5 min and the precipitated
DNA was centrifuged (15 min, room temperature) into a pellet. The DNA pellet was
rinsed with 75% ethanol, dried well and redissolved in 50-100 itL of TE. No
concentration determination was done and the sample was used directly for restriction

endonuclease analysis.

Restriction Enzyme Digestion of DNA

Restriction enzyme digests were performed in buffers provided by Gibco BRL or
New England Biolabs. A typical restriction enzyme digest contained 0.8 pig of DNA in
8 [LL of TE, 2 “L of restriction enzyme buffer (10X concentration), 1 [LL of bovine serum
albumin, 1 uL of restriction enzyme and 8 [LL TE. Reactions were incubated at 37 °C for

1 h, terminated by addition of 2.0 uL of 10X Endostop and analyzed by agarose gel

-l47-

electrophoresis. For cloning experiments, the reaction was terminated by addition of
1 [LL of 0.5 M NazEDTA (pH 8.0), followed by extraction with phenol and SEVAG
(0.1 mL each). DNA was precipitated by addition of 0.1 volume of 3 M NaOAc (pH 5.2)
followed by thorough mixing and addition of 3 volumes of 95% ethanol. Samples were
mixed and kept at -78 °C for 3 h. Precipitated DNA was recovered by centrifugation (15
min, 4 °C), treated with 0.1 mL of 70% ethanol and again centrifuged (15 min, 4 °C).

DNA was dried and redissolved in TE.

Agarose Gel Electrophoresis

Agarose gels were run in TAE buffer made up of 40 mM Tris-acetate and 2 mM
EDTA (pH 8.0). Gels typically contained 0.7% agarose (w/v) in TAE buffer. Addition
of ethidium bromide (0.5 pig mL") to the agarose allowed for visualization of DNA under
ultra-violet exposure. Two standards were used to determine the size of DNA fragments:
A DNA digested with Hindlll resulted in bands at 23.1-kb, 9.4-kb, 6.6-kb, 4.4-kb, 2.3-kb,
2.0-kb, and 0.6-kb; while A DNA digested with HindIII and EcoRI gave fragments at

21.2-kb, 5.1-kb, 5.0-kb, 4.3-kb, 3.5-kb, 2.0-kb, 1.9-kb, 1.6-kb, 1.4-kb, 0.9-kb, and 0.8-kb.

Isolation of DNA from Agarose

Two methods were used for isolating DNA from agarose gels. The first method
involved cutting out the band of agarose containing the required DNA from the gel and
chopping it thoroughly with a razor. The agarose was then transferred to a 0.5 mL
microfuge tube packed tightly with glass wool and having an 18 gauge hole at the

bottom. The tube was centrifuged for 5 min using a Beckman microfuge to extrude the

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DNA solution from the agarose into a 1.5 mL microfuge tube. The DNA was
precipitated out using 3 M NaOAc and 95% ethanol as previously described.
The second method involved use of the Zymoclean DNA isolation kit purchased

from Zymogen.

Treatment of DNA with Klenow fragment

DNA fragments with recessed 3’ termini were modified to blunt end fragments by
treatment with the Klenow fragment of E. coli DNA polymerase I. Digested DNA (4-6
ug) in 120 uL of TE was treated with 6 [L of a solution containing 25 mM of each of the
desired dNTP’S, followed by addition of 6 units of the Klenow fragment. The reaction
was incubated at room temperature for 30 min. Klenow reactions were quenched by
addition of 15 [1].. of 10X Endostop in cases where the DNA fragment needed to be
purified by agarose gel electrophoresis, or in case of vector plasmid was taken forward

directly for treatment with calf intestinal alkaline phosphatase.

Treatment of Vector DNA with Calf Intestinal Alkaline Phosphatase

Linearized plasmid vectors were dephosphorylated to prevent self-ligation. The
plasmid (4-6 pg) after digestion was dissolved in TE (120-140 uL) and treated with 14
ML of dephosphorylation buffer and 3 [LL of calf intestinal alkaline phosphatase (3 units).
The reaction was incubated at 37 °C for l h and quenched with 1 uL of 0.5 M NazEDTA
(pH 8.0) followed by heating at 75 °C for 15 min. The DNA was extracted with phenol
and SEVAG (0.1 mL each), precipitated out as described previously and redissolved in

TE.

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Ligation of DNA

Molar ratios of insert to vector were typically maintained at 3 to 1 during
ligations. A typical ligation reaction contained 0.1 pig vector DNA, 0.05 to 0.2 ug insert
in a total volume of 7 uL. To this was added 2 uL of T4 ligation buffer (5X
concentration) and 1 [LL of T4 DNA ligase (2 units). The reaction was incubated at 16 °C

for at least 10 h and then used to transform competent cells.

Preparation and Transformation of Competent Cells

Competent cells were prepared using a procedure modified from Sambrook and
co-workers.117 A single colony was inoculated into 5 mL LB containing the necessary
antibiotics. After overnight growth 1 mL of the culture was transferred to a 500 mL
Erlenmeyer flask containing 100 mL LB with the necessary antibiotics. The cells were
cultured in a gyratory shaker (250 rpm, 37 °C) until an absorbance of 0.4-0.6 was
observed at 600 nm. The entire culture was transferred to a sterile centrifuge bottle and
the cells were collected by centrifugation (4000 x g, 5 min, 4 °C). The cells were washed
with 100 mL of cold 0.9% NaCl (w/v) and then resuspended gently in 50 mL of cold 100
mM CaCl,. The suspension was kept on ice for 30 min and then centrifuged (4000 x g, 5
min, 4 °C). The cells were slowly resuspended in 4 mL of cold 100 mM CaCl2
containing 15% glycerol (v/v). Aliquots (0.25 mL) were transferred to microfuge tubes
and immediately frozen with liquid nitrogen. Competent cells were stored at -78 °C and
used over a period of at least six months without any noticeable loss in transformation

efficiency.

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In order to perform a transformation, frozen competent cells were allowed to thaw
on ice for 5 min immediately prior to use. Either 10 [LL of a ligation reaction or 1-2 uL
of 0.1 mg mL" plasmid solution was added to 0.1 mL of cold competent cells. The
solution was gently mixed and kept on ice for 30 min. The cells were then beat shocked
at 42 °C for 2 min and kept on ice briefly (1-2 min). LB (0.5 mL) was added to the cells
and the sample incubated (no agitation) at 37 °C for 1 h. Cells were pelleted in a
microcentrifuge. If the cells were to be plated onto LB, then the cell pellet was
resuspended in 0.1 mL of LB and the entire volume spread out onto the LB plate
containing the appropriate antibiotics. If the cells were to be plated out onto M9 minimal
medium plates, then the cell pellet was washed once with 0.5 mL of M9 minimal
medium. The cells were resuspended in fresh M9 minimal medium (0.1 mL) and spread
onto the plate. Competent cells which were not transformed with any DNA were also
subjected to the same transformation protocol. These cells were used to confirm the
viability of the competent cells and also to confirm absence of growth on the selective
medium without the required DNA.

Transformation was also performed by electroporation using electrocompetent
cells. In order to prepare electrocompetent cells, instead of the 100 mL culture grown for
normal competent cells, a 500 mL culture was grown. Once an absorbance between 0.5-
0.7 was observed at 600 nm, the cells were kept on ice for 15 min and harvested. The
cells were gently washed twice with cold water (350 mL and 150 mL) and then
resuspended in 20 mL of cold aqueous 10% glycerol (v/v). The cells were centrifuged

(4000 x g, 5 min, 4 °C) and the cell pellet was slowly resuspended with 5 mL of cold

-lSl-

aqueous 10% glycerol. The cells were transferred into microfuge tubes in 0.25 mL
aliquots, immediately frozen using liquid nitrogen and stored at ~78 °C.

The electroporation was performed in Bio-Rad gene pulser cuvettes with an
electrode gap of 0.2 cm. The cuvettes were chilled on ice for 5 min prior to use.
Electrocompetent cells were thawed on ice for 5 min and 0.1 mL of thawed cells was
added to the chilled cuvette. To this was added 1-2 [1L of plasmid DNA (0.1 mg mL")
and the mixture was gently shaken. The Bio-Rad gene pulser was set at 2.5 Kvolts,
25 uF and 200-400 Ohms. The outside surface of the cuvette was wiped clean and it was
placed in the sample chamber. A single pulse was applied, the cuvette was removed and
0.5 mL of LB was added to it. The contents of the cuvette were transferred to a 1.5 mL
microfuge tube. The cells were incubated at 37 °C for 1 h. The transformed cells were

plated in the same manner as in a normal transformation with the same controls.

MR;

Synthetic Procedures

Synthesis of PCA phosphonate 4 and PCA homophosphonate 5

Methyl 3,4-dimethoxy benzoate 10. To a solution of 3,4-dimethoxy benzoic
acid (100 g, 552.5 mmol) in methanol (1000 mL) was added 5 mL of conc. H2804. The
reaction mixture was refluxed for 24 h, to rt and neutralized with saturated aqueous
NaHCO3. The solution was concentrated to dryness and the solid obtained was

partitioned between EtOAc and water. The organic layer was dried and concentrated to

afford the ester 10 as a pink solid (106 g, 98 %): 1H NMR (CDCl,) 5 3.90 (s, 3H), 3.94

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(S, 6H), 6.89 (d, J: 8 Hz, 1H), 7.55 (d, J: 2 Hz, 1H), 7.68 (dd, J: 8, 2 Hz, 1H); 13C NMR
(CDC13) 8 166.7, 152.8, 148.4, 123.4, 122.5, 111.7, 110.1, 58.4, 55.8, 51.8. Anal. Calcd
for CmleO,: C, 61.25; H, 6.17. Found: C, 61.21; H, 6.10.

5-bromo-3,4-dimethoxy methybenzoate 11. A solution of methyl 3,4-
dimethoxy benzoate 10 (25 g, 127 mmol) and NaOAc (15.2 g, 190.5 mmol) in AcOH
(300 mL) at rt was treated slowly with a solution of Br2 (54 g, 337 mmol) in AcOH (100
mL) over a period of 30 min. After the addition was complete, the reaction mixture was
stirred at 70 ° C for 24 h. The reaction mixture was then poured into 1 L of ice water and
kept at 4 °C for 6 h. The brown solid which had precipitated was filtered and crystallized
from EtOAc/hexane to give 34 g of the bromide 11 in 97% yield: 1H NMR (CDCl,) 8
3.90 (s, 6H), 3.92 (s, 3H), 7.10 (S, 1H), 7.42 (s, 3H); 13C NMR (CDCl,) 8 165.8, 151.9,
147.7, 122.7, 116.8, 114.1, 113.8, 56.2, 56.1, 52.2. Anal. Calcd for CIOHHBrO4: C,
43.84; H, 4.05. Found: C, 43.94; H, 4.07.

5-vinyl-3,4-dimethoxy methyl benzoate 12. The bromide 11 (10.5 g, 38.2
mmol) and tetrakis(triphenylphosphine)palladium (0) (0.882 g, 0.764 mmol) were
dissolved in dry HMPA (260 mL) and stirred for 10 min at rt. The reaction mixture was
then heated up to 65 °C and treated with tributyl vinyltin (13.9 g, 43.9 mmol). After
18 h, the reaction was quenched by the addition of saturated aqueous KF, stirred for
15 min and filtered through celite. The filtrate was concentrated to a minimum volume
and the liquid loaded onto a silica gel column. Purification by flash chromatography
(hexane, EtOAc/hexane, 1:4 v/v) afforded the vinylic ester 12 as white crystals (5.3 g,
63%): 1H NMR (CDCl,) 8 3.90 (s, 3H), 3.92 (s, 3H), 3.96 (s, 3H), 5.30 (dd, J: 1, 11 Hz,

1H), 5.57 (dd, J: 1, 17 Hz, 1H), 7.03 (S, 1H), 7.43 (s, 1H), 7.54 (dd, J: ll, 17 Hz, 1H);

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13‘C NMR (CDCl,) 8 167.1, 151.8, 147.8, 135.8, 134.2, 120.2, 114.9, 112.6, 109.2, 55.8,
55.7, 51.8. Anal. Calcd for C,2H,,O4: C, 64.86; H, 6.35. Found: C, 64.79; H, 6.33.

3-formyl-4,5-dimethoxy methyl benzoate 13. Alkene 12 (5.33 g, 24 mmol) was
dissolved in CHzCl2 and cooled the solution to —78 °C. Ozone was then passed through
the reaction flask until the reaction was complete as indicated by TLC. Addition of 50
mL Et3N followed by concentration of the reaction mixture in vacuo gave a colorless oil.
The oil was treated with 300 mL of water and the flask kept at 4 °C for 3 h. White
crystals were obtained which were filtered off to give 4.9 g of the required forrnyl ester
13 in 91% yield: 1H NMR (CDCl,) 8 3.97 (s, 3H), 3.99 (s, 3H), 4.01 (s, 3H), 7.47 (s,
1H), 7.51 (s, 1H), 10.65 (s, 1H); 13C NMR (CDCl,) 8 191.2, 166.2, 152.3, 151.9, 131.2,
125.9, 112.5, 109.5, 56.3, 56.2, 52.6. Anal. Calcd for C,,H,205: C, 58.96; H, 5.40.
Found: C, 58.57; H, 5.49.

Phosphonate Intermediate 15. Formyl ester 13 (1 g, 4.5 mmol) and Nal (1.35 g,
9 mmol) were dissolved in dry AcCN under Ar at rt and TMSCl was added. After
stirring for 10 min the reaction mixture was cooled to 0 °C and tetramethyl disilazane
was added dropwise. After the addition was complete the reaction was stirred at 0 °C for
10 min and then refluxed for 2 h. The solution was then partitioned between EtOAc and
water. The organic layer was dried and concentrated to get a red oil containing the
required iodide 14: 1H NMR (CDCl,) 8 3.92 (s, 3H), 3.93 (s, 3H), 3.94 (s, 3H), 4.96 (s,
2H), 6.86 (s, 1H), 7.48 (s, 1H); l3C NMR (CDCl,) 8 166.4, 152.0, 148.0, 135.4, 120.0,
113.7, 113.3, 56.02, 52.03, 5.2.

The iodide 14 without any purification was dissolved in toluene (20 mL) at rt.

Triisopropyl phosphite (5.5 g, 26.2 mmol) was added and the reaction mixture refluxed

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for 24 h. The solution was concentrated to ~ 2 mL and then treated with 50 mL hexane.
The mixture was kept at -20 °C and the white crystals formed were filtered off to get the
phosphonate intermediate 15 (0.39 g, 23%): 1H NMR (CDC13) 8 1.17 (d, J: 6 Hz, 6H),
1.27 (d, J: 6 Hz, 6H), 3.77 (d, J: 23 Hz, 2H), 3.89 (s, 3H), 3.91 (s, 3H), 3.93 (s, 3H),
4.61 (m, 2H), 6.95 (d, J: 2.4 Hz, 1H), 7.46 (s, 1H); ”C NMR (CDC13) 8 167.2, 151.3,
146.9, 127.8, 121.6, 114.3, 113.2, 70.2 (d, J: 7.0 Hz), 55.8, 51.7, 31.4 (d, J: 138 Hz),
24.0, 23.8, 23.7, 23.6. Anal. Calcd for C17H27O7P: C, 54.54; H, 7.27. Found: C, 54.40;
H, 7.34.

Protocatechuic phosphonate 4. A solution of protected phosphonate 15 (0.39 g,
1.04 mmol) in dry CH2C12 (10 mL) was cooled to —78 °C and treated with BBr3 (9.5 g,
38.1 mmol). The reaction was allowed to warm to rt and stirred for 12 h. The reaction
mixture was cooled to 0 °C and slowly treated with conc. HCl (10 mL). After stirring for
1 h at rt CHCl3 (20 mL) was added, and the aqueous layer separated out and concentrated
to dryness. The residue was dissolved in water, neutralized to pH 7.5 and loaded onto a
DEAE-52 cellulose anion exchange column (25 mL). Washed the column with 50 mL
water followed by 250 mL of 200 mM Et3NH*HCO3’ (pH 7.5). The fractions containing
phosphorous were identified by the methods of Avila and Ames and the buffer removed
by azeotropic distillation of three additions of 2-propanol. The resulting white residue
was dissolved in water and passed down a short column of Dowex 50 (H*). The filtrate
was concentrated to get protocatechuic phosphonate 4 (0.161 g, 62%): 1H NMR (D20) 8
3.5 (d, J: 22 Hz, 2H), 6.85 (s, 1H), 7.38 (s, 1H); ”C NMR (D20) 8 174.4, 150.8, 145.1,
130.9 (d, J: 10 Hz), 124.8 (d, J: 6 Hz), 121.9, 121.4, 34.9 (d, J: 130 Hz); HRMS (FAB)

calcd for C8H8PO7 247.0008, found 246.9998.

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Homophosphonate Intermediate 16. A solution of n-BuLi in hexane (1.6M, 6
mL, 8.9 mmol) was added to a solution of tetraisopropyl methylenediphosphonate (2.9 g,
8.9 mmol) in dry THF (20 mL) at —78 °C. The mixture was stirred for 10 min and a
solution of the forrnyl ester 13 (2 g, 8.9 mmol) was added via cannula. The reaction was
stirred at —78 °C for 10 min and then at rt for 12 h. Saturated aqueous NH4C1 and EtOAc
were added, the aqueous layer extracted with EtOAc (2x) and the combined organic
layers dried and concentrated. Recrystallization from EtOAc/hexane afforded white
crystals of the homophosphonate intermediate 16 (1.17 g, 34%): 1H NMR (CDC13) 8
1.38 (s, 12H), 3.91 (s, 3H), 3.95 (s, 3H), 3.96 (s, 3H), 4.76 (m, 2H), 6.12 (dd, J: 17, 18
Hz), 7.03 (s, 1H), 7.46 (s, 1H), 8.23 (dd, J: 17, 22 Hz, 1H); ”C NMR (CDC13) 8 166.6,
151.8, 149.3, 146.0 (d, J: 8 Hz, 1H), 131.0 (d, J: 25 Hz), 122.0, 117.2 (d, J: 191 Hz),
112.8, 109.3, 70.6 (d, J: 6 Hz), 70.5, 56.0, 52.1, 24.1, 24.0, 23.9, 23.8. Anal. Calcd for
C,8H2,O-,P: C, 55.95; H, 7.05. Found: C, 55.40; H, 6.83.

Homophosphonate Intermediate l7. Ethenyl phosphonate 16(0.5 g, 1.3 mmol)
was dissolved in MeOH (20 mL) and hydrogenated over 10% Pd over C (0.25 g) at 50 psi
H2 pressure for 12 h. The mixture was filtered through celite, and the filtrate
concentrated in vacuo. Homophosphonate intermediate 17 was obtained as white crystals
(0.376 g, 76%): 1H NMR (CDCl,) 8 1.34 (s, 12H), 2.06 (m, 2H), 3.20 (m, 2H), 3.89 (s,
3H), 3.91 (s, 3H), 3.93 (S, 3H), 4.72 (m, 2H), 6.78 (s, 1H), 7.48 (s, 1H); ”C NMR
(CDCl,) 8 166.9, 151.9, 146.8, 137.7 (d, J: 18 Hz), 120.4, 113.5, 113.4, 69.9 (d, J: 6
Hz), 28.8 (d, J: 123 Hz), 27.8 (d, J: 11 Hz), 23.9. Anal. Calcd for C,8H290,P: C, 55.66;

H, 7.53. Found: C, 55.03; H, 7.50.

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Protocatechuic homophosphonate 5. Homophosphonate intermediate 17
(0.466 g, 1.2 mmol) was deprotected and purified as described for phosphonate
intermediate 15 to afford protocatechuic homophosphonate 5 as a white powder (0.277 g,
88%): 1H NMR (D20) 8 2.00 (m, 2H), 3.02 (m, 2H), 6.76 (s, 1H), 7.37 (s, 1H); ”C NMR
(D20) 8 172.9, 151.3, 144.7, 140.3 (d, J: 19 Hz), 122.3, 121.7, 120.6, 31.1 (d, J: 131

Hz), 29.9; HRMS (FAB) calcd for CngoPO7 261.0164, found 261.0158.

Synthesis of catechol homophosphonate 6

Catechol Homophosphonate Intermediate 19. A solution of tetraisopropyl
methylenediphosphonate in dry THF was cooled to —78 °C and treated with n-BuLi in
hexane (1.6M, 37 mL, 54.2 mmol). The mixture was stirred for10 min and a solution of
2,3-dimethoxy benzaldehyde l8 (9 g, 54.2 mmol) in dry THF was added via cannula.
The reaction mixture was stirred at —78 °C for 10 min and then at rt for 18 h. The
reaction was by addition of cold water and the product extracted out with EtOAc (3x).
The combined organic layers were dried and concentrated to get a colorless oil.
Purification by flash chromatography (hexane, EtOAc/hexane, 1:2, 1:1 v/v) gave catechol
homophosphonate intermediate 19 as a colorless oil (11.8 g, 95%): lH NMR (CDCl,) 8
1.33 (s, 3H), 1.35 (s, 3H), 1.37 (s, 3H), 1.39 (s, 3H), 3.86 (s, 3H), 3.87 (s, 3H), 4.72 (m,
2H), 6.35 (dd, J: 18, 19 Hz, 1H), 6.93 (dd, J: 2, 8H2, 1H), 7.06 (dd, J: 8, 8 Hz, 1H),
7.13 (dd, J: 2, 8 Hz, 1H), 7.78 (dd, J: 18, 23 Hz, 1H); ”C NMR (CDC13) 8 153.0, 148.0,

142.1 (d, J: 8 Hz), 129.1 (d, J: 23 Hz), 124.0, 118.8, 117.1 (d, J: 190 Hz), 113.5, 70.3

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(d, J: 6 Hz), 61.1, 55.8, 24.1, 24.0, 23.9, 23.8. Anal. Calcd for C,6H2505P: C, 58.53; H,
7.68. Found: C, 57.91; H, 7.70.

Catechol Homophosphonate Intermediate 20. A solution of catechol
homophosphonate intermediate 19 (0.87 g, 2.7 mmol) was hydrogenated over 10% Pd
over C (0.5 g) at 50 psi H2 pressure for 12 h. The reaction mixture was filtered through
celite and the filtrate concentrated in vacuo. Purification by radial chromatography (2
mm thickness, hexane, EtOAc/hexane 1:2, 1:1 v/v) afforded catechol homophosphonate
intermediate 20 as a colorless oil (0.66 g, 75%): 1H NMR (CDC13) 8 1.32 (s, 3H), 1.33 (s,
3H), 1.34 (s, 3H), 1.35 (s, 3H), 2.01 (m, 2H), 2.90 (m, 2H), 3.84 (s, 3H) 3.85 (s, 3H), 4.72
(m, 2H), 6.79 (m, 2H), 6.99 (dd, J: 8, 8 Hz, 1H); ”C NMR (CDCl,) 8 152.6, 146.9,
135.0 (d, J: 18 Hz), 123.9, 121.4, 110.6, 69.9 (d, J: 6 Hz), 60.5, 55.6, 27.9 (d, J: 138
Hz), 23.9, 23.6 (d, J: 4 Hz). Anal. Calcd for C,6H27OSP: C, 58.17; H, 8.24. Found: C,
57.56; H, 8.24.

Catechol Homophosphonate 6. Catechol homophosphonate intermediate 20
was deprotected and purified in exactly the same manner as described for phosphonate
intermediate 15 to yield catechol homophosphonate 6 as a brown oil (0.643 g, 97 %). 1H
NMR (D20) 8 2.10 (m, 2 H), 2.90 (m, 2H), 6.82 (m, 3H); ”C NMR 8 147.2, 144.6, 132.2
(d, J: 17 Hz), 123.8 (d, J: 27 Hz), 117.0, 29.5 (d, J: 132 Hz), 25.768; HRMS (FAB)

calcd for CngzPOs 219.0422, found 219.0416.

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Enzyme Kinetics

Determination of Inhibition Constants (K) for Inhibitors of DHQ synthase

DHQ synthase activity was assayed in a 1.0 mL solution of 3-(N—
morpholino)propanesulfonate (MOPS) buffer (50 mM, pH 7.5) containing either 50 11M
CoCl2 or 75 uM ZnSO4 depending on the form of enzyme being studied, NAD“ (10 MM),
DHQ dehydratase (1 unit), and varying concentrations of DAHP and inhibitor. After
equilibration at rt, purified DHQ synthase (0.024 units) was added, and the increase in
absorbance at 234 nM was monitored over time. Initial rates were determined by linear

square fits of the progress curves and were used to determine inhibition constants.

HAPTER

Strain Constructions
Strain SP1.1
Construction of SP1.1 began with the homologous recombination of the aroB
gene into the serA locus of E. coli RB791 resulting in RB791 serA::aroB. Construction
of plasmid pKL3.82A which was used for the homologous recombination was described

48

previously. The procedure used for the homologous recombination was based on

48:95” Competent E. coli RB791 was transformed with

methods described elsewhere.
pKL3.82A and plated out onto LB plates containing Cm. Colonies were observed after
incubation of the plates at 44 °C for 12 h. The resulting cointegrates were inoculated into
5 mL LB and grown at 30 °C for 12 h to allow excision of the plasmid from the genome.

Cultures were diluted (1:20,000) in LB and two more growth cycles were performed at

30 °C for 12 h to enrich the cultures for more rapidly growing cells that had lost the

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temperature-sensitive replicon. Cultures were then diluted (1:20,000) into LB and grown
at 44 °C for 12 h to promote plasmid loss from the cells. Serial dilutions of each culture
were spread onto LB plates and incubated at 30 °C for 12 h. The resulting colonies were
screened on multiple plates to select the desired recombination product. RB791
serAzzaroB was isolated based on the following growth characteristics: growth on M9
containing serine; no growth on M9; growth on LB; and no growth on LB containing Cm.

RB791 serA::aroB was then subjected to two successive P1 phage-mediated
transductions to transfer the aroL478::Tn10 and aroK::CmR loci of ALO807 onto the
genome and eliminating shikimate kinase activity. P1 phage was propagated from E.
coli. AL0807, which was generously provided by Professor M. G. Marinus.“
Transductions were carried out using protocols described by Miller.“7 RB791 serA::aroB
aroL::Tn10 aroK::CmR was selected based on the following growth characteristics:
growth on M9 containing aromatic amino acids, aromatic vitamins and serine; no growth
on M9 containing aromatic amino acids and aromatic vitamins; no growth on M9
containing serine; and growth on LB containing Tc and Cm. RB791 serAzzaroB

aroL::Tn10 aroK::CmR was renamed SP1.1.

Strain EB1.1
Construction of EB1.1 followed the 'same protocols as used for the generation of
SP1.1 except that E. coli B (ATCC11303) was used as the starting strain instead of the E.

coli K12 strain RB791.

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Strain SP1.1pts

Strain SP1.1 was subjected to P1 phage mediated transduction to transfer the
A(ptsHptsIcrr)::KanR locus of TP2811 onto the SP1.1 genome. P1 phage was propagated
from TP2811, which was generously provided by Professor Sophie Lévy.94
Transductions were carried out using protocols described by Miller.“7 Following the
transduction, cells were plated out onto LB medium containing only Kan, since no
colonies were obtained when transductions were plated out onto LB medium containing
Kan, Tc, and Cm. Colonies growing on the Kan plates were subsequently screened for
resistance to Tc and Cm. SP1.1pts possessed the following growth characteristics:
growth on LB containing Tc. Cm and Kan; no growth on M9 containing aromatic amino
acids, aromatic vitamins, and serine; and light pink colonies on MacConkey agar
supplemented with 1% glucose. One candidate possessing these growth characteristics
was made competent. Transformation of the candidate with pSC5.112B afforded
colonies, which were incapable of growth on M9 medium containing aromatic amino
acids, aromatic vitamins, and serine, while transformation of the candidate with
pSC6.09OB afforded colonies which grew on M9 medium containing aromatic amino

acids, aromatic vitamins, and serine.

Plasmid pKD12.036A

Construction of this 4.8-kb plasmid started with the PCR amplification of the

P aroE fragment from pIA321,98 followed by digestion of the PCR product with Kpnl.

laC

Plasmid pKL4.20B was linearized by treatment with Kpnl and the two DNA fragments

-l6l-

 

were ligated to generate pKDlZ.036A. Orientation of the PmaroE locus was in the

opposite direction to that of the aroI:FBR gene.

Plasmid pKD12.047A

This 6.7-kb plasmid was constructed by ligation of the serA gene into
pKD12.036A. The serA locus was excised from pD262595b by digestion with Smal and
subsequently treated with Klenow fragment. Plasmid pKD12.047A was digested with

Smal and treated with Klenow fragment. Ligation of the purified fragments resulted in

pKD12.047A in which the serA gene is transcribed in the same orientation as PmaroE.

Plasmid pKDlZ.112A

This 7.7-kb plasmid was created by insertion of the B-lac gene into pKD12.047A.
PCR amplification of the fl-lac gene was accomplished from pUC18 followed by
digestion of the 1.0-kb fragment with NcoI. Plasmid pKDlZ.047A was linearized by

digestion with Neal and the two DNA fragments were ligated to afford pKD12.112A.

Plasmid pKD12.138A

This 9.9-kb plasmid was constructed by inserting the 2.2-kb tktA fragment from
pMF51A72“ into pKDlZ.112A. Digestion of pMFSlA with BamHI followed by treatment
with Klenow fragment yielded the tktA gene with blunt ends. Plasmid pKDlZ.112A was
linearized with Hindlll and treated with Klenow fragment. Subsequent ligation of the
tktA fragment to pKD12.112A resulted in pKD12.138A. The tktA gene is transcribed in

the same orientation as the serA gene.

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Plasmid pKDlS.071B

This 11.9-kb plasmid was assembled by replacing the 1.0-kb ,B-lac gene of
pKD12.138A with the pps fragment from pKL1.87B.99 Plasmid pKL1.87B was digested
with BamHI and Hindlll and the resulting 3.0—kb pps fragment was treated with Klenow
fragment. Following digestion of pKD12.138A with NcoI, the 8.9-kb fragment was
modified to blunt ends using Klenow fragment. Ligation of these two purified fragments
yielded pKD15.071A. The pps gene is transcribed in the opposite orientation relative to

the tktA gene.

Plasmid pSC5.1 1213

This 11.1-kb plasmid was created by replacing the 1.0-kb B-lac gene of
pKD12.138A with the glf fragment from pTC325.73°'d A 2.2-kb fragment containing the

glf locus behind Pm was excised from pTC325 by digestion with BamHI, Hindlll and

Xbal followed by treatment with Klenow fragment. Plasmid pKD12.138A was digested

with NcoI and the resulting 8.9-kb fragment was treated with Klenow fragment. The two
purified fragments were ligated to generate pSC5.112B. The Pmcglf gene is transcribed in

the opposite orientation relative to tktA.

Plasmid pSC6.09OB
This 12.8-kb plasmid was constructed by ligation of the nglfglk cassette into
pKD12.138A. The nglfglk cassette was excised from pTC325 by digestion with

BamHI and Xbal and subsequently treated with Klenow fragment. Plasmid pKD12.138A

was digested with Neal and the 8.9-kb fragment was treated with Klenow fragment.

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Ligation of the purified fragments resulted in pSC6.09OB in which the nglfglk cassette

is transcribed in the same orientation as the tktA gene.

Plasmid pSC6. 142B
This 7.5-kb plasmid was assembled by inserting the glf gene into pKL1.87A.

Plasmid pTC325 was digested with BamHI, Hindlll and Xbal and the resulting 2.2-kb
nglf fragment was treated with Klenow fragment. Plasmid pKL1.87A was linearized
by digestion with Sall subsequently treated with Klenow fragment. Ligation of the ngIf

fragment into pKL1.87A afforded pSC6.142B in which the ngIf gene is transcribed in

the opposite orientation relative to the pps gene.

Plasmid pSC6.162A

This 14.1-kb plasmid was constructed by inserting the 5.2-kb ppstglf cassette
from pSC6.142B into pKDlZ.138A. Plasmid pSC6.142B was digested with BamHI and
Hindlll to liberate the ppst gl f fragment, which was then treated with Klenow

fragment. The 1.0-kb fi-lac gene was excised from pKD12.138A by digestion with Neal
and the resulting 8.9-kb fragment was treated with Klenow fragment. Ligation of the two

purified DNA fragments afforded pSC6.162A. The nglf fragment was transcribed in

the same orientation and the pps gene in the opposite orientation relative to the tktA gene.

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Plasmid pSC6.301A

This 15.8-kb plasmid was created by inserting the pps gene into pSC6.090B.
Plasmid pKL1.87B was digested with BamHI and Hindlll and the resulting 3.0-kb pps
fragment was treated with Klenow fragment. Plasmid pSC6.09OB was linearized by
digestion with Xbal and then treated with Klenow fragment. Ligation of the pps
fragment into pSC6.09OB afforded pSC6.301A. The pps gene is transcribed in the same

orientation relative to the tktA gene.

HA 4

Strains Constructions
Strain SC1.0

Construction of SC1.0 started with the P1 phage-mediated transductions of
aroL478zzTn10 and aroKzszR into JWFl thereby knocking out the aroL and aroK
encoded isozymes of Shikimate kinase. This strain was designated as SC1.1 and it was
selected based on the following growth characteristics: growth on M9 containing
aromatic amino acids, aromatic vitamins and serine; no growth on M9 containing
aromatic amino acids and aromatic vitamins; no growth on M9 containing serine, and
growth on LB containing Tc and Cm.

The construction of SC1.0 was completed by the P1 phage-mediated transduction
of aroB::Tn5 into SC1.1. P1 phage was propagated from JB5, which was already
available in the Frost laboratory.95b Selection of SC1.0 was based on the following
growth characteristics: growth on M9 containing aromatic amino acids, aromatic

vitamins, and serine; no growth on M9 containing aromatic amino acids and aromatic

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vitamins; no growth on M9 containing serine and; growth on LB containing Tc and Cm,

Kan

Strain SP1.lshiA
E. coli SP1.13hiA was constructed by transduction of the shiAzzKanR locus from
JP11123 into SP1.1. The preparation of strain JP11123 has been previously reported by

‘ Transduction of the shiA::KanR locus via P1 phage from

Pittard and co-workers.‘1
JP11123 into SP1.1 followed the procedure described by Miller.”7 Following the
transduction, cells were spread onto LB plates containing Kan. Plates were incubated at

37 °C for 16 h before colonies appeared. SP1.1shiA was identified based on growth on

LB plates containing Tc, Cm, and Kan.

Plasmid pSC5.214A

Plasmid pSC5.214A was constructed via disruption of the aroF‘BR gene in
plasmid pKDl2.138A. This was achieved by utilizing the unique BglII restriction site
present at 0.623-kb within aroE“. Plasmid pKD12.138A was linearized by digestion
with BglII followed by treatment with Klenow fragment. Subsequent ligation afforded

the 9.9-kb plasmid pSC5.214A.

-l66-

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