Structure-function relationships in a f3-1,4

Structure-function relationships in a f3-1,4-glycanase (Cex) from
Cellulomonas fimi: identification of catalytic residues
by
Alasdair Muir MacLeod
B.Sc. University of Western Ontario, 1986
M.Sc. Laurentian University, 1989
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11
ABSTRACT
The objective of this study was to identify and examine the roles of catalytic
residues in the active site of a 13-1,4-glycanase (Cex) from the bacterium Cellulomonas
fimi. The use of the Gram positive bacterium, Streptomyces lividans, as a host for the
expression of the cex gene was explored. The cex gene was successfully expressed in
S. lividans from the aph promoter of p1J680-cex. The polypeptide was efficiently
exported to the culture medium. Yields of recombinant polypeptide were about 6
mg/L following 40 hours of growth. Cex produced by S. lividans had a molecular
mass greater than that produced by E. coli due to glycosylation. Similar to the native
enzyme from C. fimi, the glycosylation protected the enzyme from a C. fimi protease,
particularly when the enzyme was bound to cellulose.
Cex hydrolyses E3-1,4-glycosidic bonds in cellulose with retention of anomeric
configuration, releasing 13-cellobiose. On the basis of amino acid sequence
alignments, Glu 233 was proposed as the catalytic nucleophile in Cex. The kinetic
parameters determined for Glu 233 mutants generated by site-directed mutagenesis
were consistent with this interpretation. Similarly, Glu 127 was proposed as the
acid/base catalyst in Cex. The kinetic parameters were determined for Glu 127
mutants generated by site-directed mutagenesis, using a range of soluble
cellobiosides and glucosides with differing requirements for acid catalysis. The
results were consistent with a role of Glu 127 as acid/base catalyst. In the presence
of azide, a new product, f3-cellobiosyl-azide was formed with uncharged Glu 127
mutants and catalytic activity was restored to wild-type levels. The results suggest
azide occupied a vacant anionic site consistent with the removal of the acid/base
catalyst. The approach used in this study could be applied to the identification of
the acid/base catalyst in other {3-l,4-glycanases, particularly in the absence of threedimensional structure information.
111
TABLE OF CONTENTS
ABSTRACT
ii
LIST OF TABLES
vi
LIST OF FIGURES
vii
LIST OF ABBREVIATIONS
ix
ACKNOWLEDGMENTS
xi
1. INTRODUCTION
1
1.1 Structure-function relationships
1
1.2 Cellulose and cellulases
1
1.3 Cellulomonasfimi f3-1,4 glycanase (Cex)
3
1.4 Catalytic mechanisms for hydrolysis of a 13-1,4 glycosidic linkage
5
1.5 Identification of catalytic residues
6
1.6 Expression of the cex gene
15
1.7 Objectives
17
2. MATERIALS AND METHODS
18
2.1 Buffers, enzymes and media components
18
2.2 Bacterial strains, plasmids and phage
18
2.3 Media and growth conditions
19
2.4 Recombinant DNA techniques
21
2.4.1 Production and isolation of single-stranded DNA
21
2.4.2 Site-directed in vitro mutation of cex
22
2.4.3 Sequencing of DNA
23
2.5 Detection of cex gene expression
2.5.1 MUCase and PNPCase assays
24
24
iv
2.5.2 Binding to Avicel
.
2.6 Detection of protein
24
25
2.6.1 SDS-PAGE and Western Blotting
25
2.6.2 Glycoprotein detection
26
2.6.3 N-terminal amino acid sequencing
26
2.6.4 Determination of protein concentration
26
2.7 Production and purification of recombinant Cex
26
2.7.1 Production of Cex in Streptomyces lividans TK64
26
2.7.2 Production of Cex and Cex mutants in E. coli
27
2.7.3 Purification of Cex
27
2.8 Proteolysis of Cex
28
2.8.1 Substrate-bound Cex
28
2.8.2 Cex in solution
29
2.9 Enzyme kinetics
29
2.9.1 Determination of steady-state kinetic parameters
29
2.9.2 Kinetics of inactivation of Cex mutants with 2F-DNPC
30
2.10 Characterization of the products of enzymatic hydrolysis
31
2.10.1 Thin layer chromatography
31
2.10.2 H
1
-NMR spectrometry of the products
of enzymatic hydrolysis
32
2.11 Protein Mass determination
32
2.12 Amino acid sequence alignments and database searches
32
3. RESULTS (part 1): expression of the cex gene in Streptomyces lividans
33
3.1 Construction of p1J702-cex and p1J680-cex
33
3.2 Production of Cex by S. lividans TK64[pfJ702-cex I
33
3.3 Production of Cex by S. lividans TK64[p1J680-cex.]
39
V
3.4 Glycosylation of Cex by S. lividans
.
4. RESULTS (part 2): identification of catalytic residues in Cex
44
49
4.1 Wild-type Cex: kinetics
49
4.2 The putative catalytic nucleophile in Cex: Glu 233
54
4.2.1 Generation of mutants at position 233
4.2.2 Determination of kinetic parameters for Glu 233 mutants
4.3 Identification of the acid/base catalyst
4.3.1. Generation of mutants at position 127
4.3.2 Kinetic characterization of mutants of Glu 127
55
60
63
63
65
4.3.3 Effects of sodium azide on reaction rates
72
4.3.4 Effects of sodium azide on products of hydrolysis
78
4.4 Asp 123: another conserved acidic residue
5. DISCUSSION
85
91
5.1 Expression of the cex gene in Streptornyces lividans
91
5.2 Catalysis
94
5.2.1 The catalytic nucleophile
94
5.2.2 The acid/base catalyst
95
6. APPENDIX
101
6.1 Determination of kcat and Km: an example
101
6.2 Interpretation of kcat, K
1 and kcat /IKm
104
6.3 Nucleotide and amino acid sequence of Cex
106
7. REFERENCES
108
vi
LIST OF TABLES
Table
1.1
Family F of 13-1,4-glycanases
2.1
Bacterial strains
19
2.2
Plasmids, phagemid and phage
20
2.3
Oligonucleotides used in site-directed mutagenesis of cex
23
2.4
Wavelengths monitored and extinction coefficients for hydrolysis
of aryl cellobiosides and aryl glucosides
30
3.1
N-terminal processing of native and recombinant Cex
43
4.1
Kinetic parameters for hydrolysis of cellobiosides and glucosides
by wild-type Cex
53
.
11
4.2
Kinetic parameters for hydrolysis of p-nitrophenyl-13-D cellobioside
(PNPC) by Cex and Cex P233 mutants
62
4.3
Mass spectrometry of purified Cex and E127 mutants in the
absence and presence of 2,4-DNPC
66
Kinetic parameters for hydrolysis of various substrates by Cex and
E127 mutants
68
4.5
HNMR spectra for 13-cellobiosyl-azide and -glucosyl-azide
1
82
4.6
Products of hydrolysis with wild-type and E127 mutants in the
presence or absence of sodium azide
84
Kinetic parameters for hydrolysis of cellobiosides by D123A
88
4.4
4.7
vu
LIST OF FIGURES
1.1
Cellulose and its hydrolysis
1.2
The exoglucanase/xylanase Cex from Cellulomonns firni
4
1.3
Stereochemical classification of f3-1,4 glycanases
5
1.4
Proposed mechanism for hydrolysis of glycosides by Cex
7
1.5
2F-DNPC: a mechanism-based inactivator of Cex
8
1.6
Cellobiosides with differing requirements for acid catalysis
13
3.1
Construction of p1J702-cex
35
3.2
Construction of p1J680-cex
37
3.3
Production of exoglucanase by S. lividans TK64[p1J702-cexl
38
3.4
Production of exoglucanase by S. lividans TK64[p1J680-cex]
40
3.5
Identification of extracellular Cex in S. lividans [p1J680-cexl
cultures
42
3.6
Glycosylation of Cex by S. lividans
45
3.7
Sensitivity of Cex from S. lividans to the protease from C. fimi
48
4.1
Alignment of family F catalytic domains
50
4.2
Construction of pTZI8R-cex
57
4.3
Generation of pUC12-1.lcex(PTIS) encoding Cex mutants
59
4.4
Purification of Cex E233D and E233Q
61
4.5
Purification of Cex E127A, E127G and E127D
64
4.6
Inactivation of Cex EI27A by 2F-DNPC
71
4.7
PNPCase activity of Cex and E127 mutants in the presence of
various concentrations of sodium azide
74
.
2
viii
4.8
4.9
4.10
4.11
4.12
Kinetic parameters for hydrolysis of PNPC by Cex E127 in the
presence of various concentrations of sodium azide
75
Kinetic parameters for hydrolysis of 2,4-DNPG by Cex E127 in the
presence of various concentrations of sodium azide
76
Kinetic parameters for hydrolysis of 2,4-DNPC by Cex E127 in the
presence of various concentrations of sodium azide
77
Proposed mechanism for hydrolysis of glycosides by Cex E127A
in the presence of azide
79
Hydrolysis of PNPC by wild-type Cex in the presence and absence
of azide
80
Hydrolysis of PNPC by Cex E127A and E127G in the presence and
absence of azide
81
Time course of hydrolysis of PNPC by Cex E127A in the presence
of azide
82
4.15
Purification of Cex D123A
86
4.16
PNPCase activity of Cex D123A in the presence and absence of
azide
87
Hydrolysis of PNPC by Cex D123A in the presence and absence
of azide
90
4.13
4.14
4.17
ix
LIST OF ABBREVIATIONS
2,4-DNPC
2”,4”-dinitrophenyl f3-D-cellobioside
2,4-DNPG
2’,4’-dinitrophenyl f3-D-glucoside
2F-DNPC
2”,4”-dinitrophenyl 2-deoxy-2-fluoro 13-D cellobioside
2F-DNPG
2”,4”-dinitrophenyl 2-deoxy-2-fluoro 13-D glucoside
4-BrPC
4” -bromophenyl f3-D-cellobioside
A
absorbance
aa
amino acid
aph
gene encoding aminoglycoside phosphotransferase
Apr
ampicillin resistance
bp
base pair
Cex
C. fimi exo 3-1,4-glycanase
CMC
carboxymethylcellulose
DMSO
dimethyl sulfoxide
EDTA
ethylenediamine tetra-acetic acid
IPTG
isopropyl-f3-D-thiogalactoside
kb
kilobase pairs
kcat
enzyme turnover number
kDa
kilodaltons
K
equilibrium binding constant
k
inactivation rate constant
Km
Michaelis cons Iant
I<rnr
kanamycin resistance
lacZpo
E. coli lacZ gene promoter and operator
LB
Luria-Bertani medium
me!
gene encoding tyrosinase (melanin)
x
Mr
relative molecular mass
MUC
methylumbelliferyl 13-D-cellobioside
NMR
nuclear magnetic resonance
PEG
polyethylene glycol
PFU
plaque-forming units
PNPC
paranitrophenyl 13-D-cellobioside
PNPG
paranitrophenyl -D-glucoside
PTIS
portable translation initiation site
R2YE
regeneration yeast extract
SDS-PAGE
sodium dodecyl sulfate-polyacrylamide gel electrophoresis
TE
tris-EDTA
Thr
thiostrepton resistance
TLC
thin-layer chromatography
TYP
tryptone, yeast extract, phosphate
maximal velocity of enzyme-catalysed reaction
xi
ACKNOWLEDGMENTS
I wish to extend my thanks to my supervisor Dr. Tony Warren for providing
continual advice and encouragement throughout this study. I also thank Drs. Doug
Kilburn, Robert Miller and Neil Gilkes for their guidance.
My appreciation also goes to Dr. Loida Carlson of the UBC Biotechnology lab
who helped me with the Streptomyces expression. Dr. Steve Withers, Karen Rupitz,
Dr. Thisbe Lindhorst and Dee Tull of the UBC Chemistry Department provided
valuable discussions regarding the kinetic studies of Cex. In addition, Dr. Withers
provided substrates crucial to the success of this study.
Members of the cellulase lab, too numerous to mention, provided an enjoyable,
enriching and stimulating environment in which to work. In particular, I’d like to
thank Shen Hua, John Coutinho, Emily Kwan and Edgar Ong, former members of
the Wesbrook room 206 family, for their mentorship. Patti Miller was indispensable
to the running of the lab and for informing me when new flavours appeared at
Baskin-Robbins. I am also grateful to Helen Smith for her original sense of humour
and for providing the opportunity on many occasions to discuss science while
descending Pocalolo or Exhilaration at Whistler/Blackcomb. I thank David for
listening to a lot of stuff that may have seemed dry at times.
I appreciate the financial support received from PENCE and from NSERC for a
postgraduate scholarship.
1
1. Introduction
1.1 Structure-function relationships
Modern molecular biological techniques are providing new tools with which
to study the structure and function of biocatalysts, leading to new insights into the
mechanisms of catalysis. Such information is making possible the engineering of
novel biocatalysts with desired properties, including enhanced catalytic efficiency,
altered substrate specificity and increased pH stability or thermostability. There is
little doubt that a greater understanding of the structure-function relationships in
enzymes, including an understanding of the roles that particular amino acid
residues play in catalysis, will facilitate the design of new enzymes.
1.2 Cellulose and cellulases
The study of structure-function relationships in enzymes that will hydrolyse
plant biomass has received considerable attention over the last twenty years partly
because of the economic potential of these enzymes in industries such as fuel
production and pulp and paper (Yang et a!., 1992). Cellulose is the most abundant
carbohydrate available from plant biomass (Beguin and Aubert, 1994).
Cellulose
molecules are linear polymers of 13-1,4-linked D-glucose residues. Native cellulose from
plant cell walls consists of individual cellulose chains, each consisting of up to 10,000
glucose residues, which are arranged in parallel and are hydrogen bonded to each other
to form an insoluble crystalline matrix (Blackwell, 1981; Figure 1.1 this study). Within
the matrix, there are regions of high crystallinity and regions more amorphous in
2
A
B9°9x
Crystalline
region
4
Amorphous
region
Adsorption of cellulases
Endoglucanases
Cellobiohydrolases
4
o.oo
3-glucosidases
Figure 1.1. Cellulose and its hydrolysis (Adapted from Beguin and Aubert, 1994):
A) Molecular structure of cellulose: 13-1,4 linked D-glucopyranose polymer (only 3
residues shown). B) Synergistic model for cellulose hydrolysis. Shaded glucopyranose
residues indicate the reducing ends of the cellulose chains.
3
nature. Considerable effort has been directed towards the understanding of structurefunction relationships in enzymes which hydrolyse this substrate.
Many microorganisms, both bacteria and fungi, are capable of hydrolysing
cellulose. It is dear that the hydrolysis of cellulose presents a challenge to
microorganisms in terms of both the accessibility and the insolubility of the substrate.
Although there are a variety of strategies employed in the hydrolysis of cellulose to
glucose by microorganisms, most rely on the secretion of a number of cellulolytic
enzymes which act synergistically (Beguin and Aubert 1994). The generally accepted
model for hydrolysis of cellulose is that three major classes of enzymes are involved
(Figure 1.1). Endoglucanases (EC 3.2.1.4) hydrolyse f3-1,4 linkages in the internal
regions of a cellulose chain, probably initiating attack at the amorphous regions of the
cellulose fibril. This hydrolysis results in the liberation of oligosaccharides of various
sizes which can then be acted on by exoglucanases or cellobiohydrolases (EC 3.2.1.91).
The exoglucanases or cellobiohydrolases hydrolyze oligosaccharides generally from the
non-reducing ends to liberate cellobiose units. Cellobiose can then be hydrolysed into
glucose by the f3-glucosidases (EC 3.2.1.21).
1.3 Cellulomonas fimi 3-1,4-glycanase (Cex)
Cellulomonas fimi is a Gram positive, coryneform, mesophilic bacterium capable
of growth on cellulose. To date, several cellulase genes from this organism have been
cloned, sequenced, and expressed in E. coli (Gilkes et a!., 1984a; Wong et a!., 1986; O’Neill
et a!., 1986; Moser et a!., 1989; Coutinho et a!., 1991; Meinke et a!., 1991; 1993; 1994). These
include four endoglucanases, CenA, CenB, CenC, and CenD, an exoglucanase/xylanase
4
(Cex) and a cellobiohydrolase (CbhA). A seventh cellulose-binding polypeptide
(Cbpl2O) is currently being characterized (Shen et a!., 1994).
Cex, 443 amino acids long, comprises an N-terminal catalytic domain (315 amino
acids) and a C-terminal cellulose binding domain (107 amino acids) separated by a
proline-threonine rich linker (Figure 1.2). The domains in Cex retain their respective
catalytic and cellulose-binding properties when separated proteolytically (Gilkes et a!.,
1988, 1989). Cex contains three disulfide linkages, two within the catalytic domain and
a third within the cellulose binding domain (Gilkes, et at., 1991a). When expressed in E.
coli, Cex has a molecular mass of 47 kDa. Native Cex from C. fimi is a glycoprotein with
a slightly greater molecular mass of 49 kDa. The glycosylation is not necessary for the
catalytic function of the enzyme (Langsford et at., 1984). When the enzyme is bound to
cellulose, however, the glycosylation affords it protection from a protease present in
culture supernatants of C. fimi (Langsford et at., 1987).
42aa
316aa
U
ss
leader
catalytic domain
lO7aa
U
ss
ss
PT
linker
20 aa
47.1 kDa
I
celuose
binding
domain
Figure 1.2. The f3-1,4-glycanase Cex from Cellulomonasfirni
aa = amino acid; ss = disulfide bond; PT = proline/threonine
Cex was originally classified as a cellulase, however, it has significant activity on
xylan, a 3-1,4 linked polymer of D-xylose (Gilkes et at, 1984b). Xylan, although
chemically similar to cellulose, is structurally more complex (Settineri et at., 1965).
5
Xylan molecules may be twisted and the backbone substituted with arabinose,
glucuronic acid or methyiglucuronic acid. Although the activity of Cex on insoluble
cellulose is quite low relative to other cellulases, it has significant activity against a
range of soluble cellobiosides, glucosides and xylobiosides such as p-nitrophenyl f3-D
cellobioside (PNPC) and 2’,4’-dinitrophenyl f3-D glucoside (2,4-DNPG) and p
nitrophenyl [3-D xylobioside (PNPXX) (Gilkes et a!., 1991 a; this study).
1.4 Catalytic mechanisms for hydrolysis of a 13-1,4-glycosidic linkage
The hydrolysis of a 13-1,4-glycosidic linkage results in either the retention or
inversion of the configuration at the anomeric carbon (CI) (Sinnott, 1990). 13-1,4-.
glycanases can thus be classified according to their stereospecificity. Those enzymes for
which the products of hydrolysis retain the [3-configuration (i.e. [3—> [3)are termed
“retaining” enzymes. In contrast, those which catalyse the hydrolysis of this linkage
with inversion of configuration at the anomeric carbon (i.e. f3 —> a) are termed
“inverting” enzymes (Figure 1.3).
HO
}
0
{
Retaining
HO\.oR
.
Inverting
OH
Figure 1.3. Stereochemical classification of [3- 1,4 glycanases. Only one
glucose residue is shown for simplicity. The site ofbond cleavage is
indicated by
.
6
Cex is a “retaining” enzyme. It hydrolyses 3-1,4-g1ycosidic bonds in cellulose and
xylan, two residues in from the non-reducing terminus, with release of f3-cellobiose or
xylobiose (Withers et at., 1986). Hydrolysis therefore occurs with retention of
configuration at the anomeric carbon. It is generally accepted that hydrolysis of a l-1,4glycosidic bond by a retaining enzyme, such as Cex, involves the formation and
hydrolysis of a covalent c-D-glycopyranosy1-enzyme intermediate via oxocarbonium
ion-like transition states (Koshland et al., 1953; Withers et al., 1988) as shown in Figure
1.4. A number of amino acids must be involved in binding and stabilizing the transition
state complexes. Two amino acids, the catalytic nucleophile and the acid/base catalyst
play particularly important roles. In the formation of the glycosyl-enzyme intermediate
(the glycosylation step) the acid/base catalyst donates a proton to the glycosidic oxygen
which facilitates bond cleavage by stabilizing the leaving group. The glycosyl-enzyme
intermediate is stabilized by the formation of a covalent bond between the carboxyl
group of the catalytic nucleophile and the anomeric carbon (Cl) of the sugar. The
second step (the deglycosylation step) involves the hydrolysis of the glycosyl-enzyme
intermediate. In this step, the acid/base catalyst acts as a general base by accepting a
proton from water. The resulting hydroxyl group then acts to displace the catalytic
nucleophile resulting in a product which retains the j3-configuration at the anomeric
carbon.
1.5 Identification of catalytic residues.
The catalytic nucleophile in a retaining 13-1,4-glycanase, such as Cex, can be
“trapped” using a mechanism-based inactivator such as 2 “,4” dinitrophenyl 2-deoxy-2fluoro f3-D cellobioside (2F-DNPC; see Figure 1.5). The basis for such trapping is that
the presence of the fluorine on carbon 2 destabilizes the transition states for both the
7
‘A/B
7’
o
O1
Nuc
Nuc
-ROH
‘A/B
7’
Glycosylation
o
0HO
glycosyl-enzyme
intemdiate
Deglycosylation
Nuc
0
2
+H
A/B
oc
/\
7’
o
Hoo:
O\fO
Nuc
Nuc
Figure 1.4. Mechanism for hydrolysis of glyco sides by Cex. The acid/base
catalyst is represented by an “A/W. The catalytic nucleophile is represented by “Nuc”
Only one glucose residue is shown for simplicity. Adapted from Koshland et al., 1953.
8
formation and hydrolysis of the glycosyl-enzyme intermediate (refer to Figure 1.4)
resulting in the slowing down of both steps of the reaction. With the presence of an
activated leaving group on the substrate such as dinitrophenol the requirement for acid
catalytic assistance is reduced facilitating an increase in the rate of the first step
(glycosylation).
2
NO
2F-DNPC
Figure 1.5. 2F-DNPC: a mechanism-based inactivator of Cex
The net result is the accumulation or HtrappingI of the glycosyl-enzyme intermediate
(Withers et al.,1990). The covalently-bound catalytic nucleophile can then be identified
in proteolytic digests of the protein, using mass spectrometry to compare the masses of
the peptides with those released from the native (non-inactivated) enzyme. Using the
mechanism based inactivator 2 ‘,4 dinitrophenyl 2-deoxy-2-fluoro 3-D glucoside (2FDNPG), the catalytic nucleophile in an Agrobacteriurn 13-glucosidase (Abg), a retaining
glucanase from another family of enzymes, was identified as G1u358 (Withers and
Street, 1988; Withers et al, 1990).
No reliable strategies have been devised for the unequivocal labeling of
the acid/base catalyst in a retaining glycanase. Group specific reagents such as
carbodiimides can be used to label carboxylic acid residues with unusually high pKa
13-
9
values; however, these frequently label at multiple sites. Affinity labels such as glycosyl
epoxides have also proven unreliable in this regard (Legler, 1990). X-ray
crystallographic analysis of enzyme or enzyme/inhibitor complexes often reveals
residues that are suitably positioned to act as catalytic residues. Hen egg-white
lysozyme (HEWL), a retaining 13-glycanase, was the first enzyme for which the threedimensional structure was determined by X-ray crystallography (Blake et al., 1965).
Two acidic residues, Asp 52 and Glu 35 were determined to be correctly positioned in
the active site to act as the stabilizing anion/nucleophile and the acid/base catalyst
respectively. Although the mechanism is still a subject of investigation, it was
essentially derived de novo from examination of the three-dimensional structure.
However, it is estimated that three-dimensional structural information is available for
only 4% of those proteins for which the amino acid sequence is known (Rost et al., 1993).
Although some proteins are clearly related in amino acid sequence to proteins of known
three-dimensional structure, for 5 of every 6 new genes sequenced, there is no
homologous structure in current data banks (Rost et al., 1993). The catalytic domain of
Cex has been crystallized (Bedarkar et al., 1992) but the structure has yet to be solved.
To date, the genes encoding more than 170 cellulases and xylanases from various
microorganisms have been cloned and sequenced. Currently, these enzymes can be
grouped into about a dozen families (A to L) on the basis of amino acid sequence
similarities in their catalytic domains (Henrissat et al., 1989). To date, three-dimensional
structures have been published for only a few of these enzymes. Many of these families
contain enzymes from bacteria and fungi, and some contain both endo- and exo -f3-1,4glycanases. Endoglucanases and exoglucanases clearly have active sites of somewhat
different topologies. However, members of a given family have been shown to exhibit
the same stereospecificity (Gebler et a!., 1992b). Therefore, the enzymes within a
particular family hydrolyse 13-1,4-glycosidic bonds by the same mechanism (i.e. with
10
retention or inversion) suggesting that their catalytic residues are likely to be
conserved. Cex belongs to family F of 3-1,4-glycanases (Henrissat et at., 1989). In 1989,
twelve members could be assigned to this family on the basis of amino acid sequence
alignments. Currently, twenty enzymes can be assigned to the family, sixteen of
bacterial origin and four of fungal origin (Table 1.1). Family F-members are primarily
categorized as 3-1,4-xylanases although activity against PNPC or methylumbelliferyl
-
cellobioside (MUC) has been reported for several of them (Grepinet et at., 1988; Luthi et
at., 1990; Lin et at., 1991; Shareck et at., 1991; Haas et at., 1992). In addition, low activity
against carboxymethylcellulose (CMC) has been reported in some instances (Luthi et at.,
1990). Clearly, family F enzymes have a mixed specificity for both xylan and cellulose
in contrast to the low molecular weight family G xylanases, which are not active against
cellulose. In addition to Cex, which was the first family F enzyme to be crystallized
(Bedarkar et at, 1992), two more enzymes in the family have been crystallized: XynA
from Pseudomonas fluorescens (Pickersgill et at., 1993) and, very recently, Clostridium
thermocellum XynZ (Souchon et at., 1994). To date, however, no three-dimensional
information has been published for any member of this family. Furthermore, prior to
the initiation of this study, no catalytic residues had been identified in any member of
this family. However, in families such as family F, which contains a large number of
enzymes, amino acid sequence alignments can serve to pinpoint potential catalytic
residues in the absence of three-dimensional structure information. Acidic residues,
particularly aspartates and glutamates, are the most likely to act as catalytic
nudeophiles and acid/base catalysts (Sinnott, 1990; Zvelebil and Sternberg, 1988).
Several such residues are conserved throughout family F (see Results, Figure 4.1).
Site-directed in vitro mutagenesis has proven to be a valuable tool in probing the
structure-function relationships of enzymes and has contributed greatly to an
understanding of the role of specific residues in catalysis. Mutation of “critical” residues
11
previously targeted by chemical modification alone has often proved the residues not to
have the roles proposed for them (Schimmel 1990). Even in cases where the threedimensional structure is known, site-directed mutagenesis experiments have been very
revealing in confirming or contradicting existing beliefs derived from such information
regarding the roles amino acids play in catalysis. For example, studies involving the
Table 1.1
Family F of 3-1,4-g1ycanases’
Enzyme
Organism
XynA
XynA
XynA
XynB
XynA
Ce1B
ORF4
Cex
Xyn
XynX
XynZ
Xyn
Xyn
XynA
XynB
XynA
XynA
Xy1A
Xyn
XynA
Aspergillus kawachii
Bacillus sp. strain C-125
Butyrivibrio fibrisolvens
Butyrivibriofibrisolvens H17c
Caldocellum saccharolyticurn
Caldocellum saccharolyticurn
Caldocellum saccharolyticum
*Cellulomonasfjmi
Clostridium stercorarium (str F9)
Clostridium thermocellum
*Clostridium thermocellum
Cryptococcus albidus
Penicillium chrysogenum
*pseudomonas fluorescens
Pseudomonasfluorescens
Ruminococcusfiavefaciens
Streptomyces lividans
Thermoanaerobacter
saccharolyticum (B6A-RI)
Thermoascus aurantiacus
Thermophilic bacterial sp rt8.84
type
# amino acids
total cat
Acc#
f
b
b
b
b
b
b
b
b
b
b
f
f
b
b
b
b
328
368
378
635
312
1011
312
443
387
1087
809
311
353
585
555
1092
436
328
368
350
347
312
347
312
315
387
?
315
311
353
345
272
332
436
D14847
P07528
P23551
X61495
M34459
A43802
M34459
L11080
D12504
M67438
M22624
JS0734
S31307
Xl 5429
P23030
P29126
M64551
b
f
b
1127
p.sq
685
?
?
?
M97882
P23360
L18965
Acc# = GenBank or swissprot database accession number; cat = catalytic (family F)
domain; p.sq. = partial (protein) sequence; type: f = fungal, b = bacterial. ? = unknown;
*
= enzyme (or catalytic domain) has been crystallized; 1
As of Mar, 1994.
12
replacement of the putative catalytic nucleophile in T4 lysozyme (Asp2O) with Cys
(Hardy and Poteete, 1991) or with Ala (Rennell et al., 1991) revealed the mutant
retained 80% of its activity, casting into doubt the previous assignment of this residue as
the catalytic nucleophile.
The determination of Michaelis-Menten parameters for hydrolysis of various
substrates by mutants of Cex generated by site-directed in vitro mutagenesis can
provide insight into the roles that various amino acids play in catalysis. Wild-type Cex
will hydrolyse aryl glucosides and aryl cellobiosides and such as p-nitrophenyl -D
cellobioside (PNPC), 2”, 4”-dinitrophenyl f3-D-cellobioside (2,4-DNPC) and 4”bromophenyl 3-D-cellobioside (4-BrPC). These well-defined soluble substrates contain
aglycon moieties with different leaving group abilities (Figure 1.6). Those of high pKa,
therefore poor leaving group ability, will have a greater requirement for acid catalytic
assistance for hydrolysis of the 3-1,4-g1ucosidic linkage than those of low pKa. In effect,
hydrolysis of those substrates with a high pKa will depend on the presence of a
functional acid/base catalyst. In contrast, those of low pKa should have a much
reduced requirement for the presence of a functional acid catalyst for departure. A
detailed examination of the kinetic parameters of mutants of Cex with these substrates,
therefore, will aid in the identification of the acid/base catalyst. kcat values will yield
information on how particular mutations effect the overall rates of hydrolysis of these
substrates. Based on knowledge of the mechanism of Cex, examination of the Km and
kcat/Km values will yield information on how particular mutations effect the overall
rates of the glycosylation or deglycosylation steps. The generally-accepted double
displacement mechanism for Cex (see Figure 1.4) can be expressed as follows:
13
SUBSTRATE
pKa of aglycon unit
2
NO
OH
3.96
HOOZ©
HO
HO
2,4-DNPC
7.18
PNPC
Br
OH
9.34
4-BRPC
Figure 1.6. Cellobioside substrates with differing requirennts for acid catalysis.
The site of bond cleavage is indicated by
?tt
14
>glycosylation
deglycosylation
3
k
E.S
E+S
E-G
-ROH
where,
E
S
E.S
E-G
P
=
=
=
=
=
E+P
÷H20
concentration of unbound enzyme
concentration of unbound substrate
concentration of enzyme-substrate complex
concentration of glycosyl-enzyme intermediate
concentration of unbound product
The value of K the apparent dissociation constant for all the enzyme-bound species,
can be expressed as:
Km
=
[El [SI
or
[all enzyme bound species I
[El [SI
[E.S] + [E-GI
[2]
It can be seen from equations [1] and [21 that mutations which reduce the rate of
deglycosylation will result in an accumulation of the glycosyl-enzyme intermediate
[E-G] and therefore result in a reduction in Km. Similarly, a reduction in the rate of
glycosylation would be expected to increase the Km. These assumptions are valid
provided deglycosylation is the rate-limiting step as shown in the Appendix. The effect
of a mutation on the rate of the glycosylation step can also be estimated by examination
of kcat/Km which presumably reflects the rate of the formation of the glycosyl-enzyme
intermediate from the free enzyme and substrate. Another measure of the effect of a
mutation on the glycosylation step of the reaction can be made by examination of the
15
rate of the inactivation of the mutant relative to the wild-type using the mechanismbased inactivator 2F-DNPC. Essentially, the reaction proceeds only through the
glycosylation step as discussed earlier. That is, k
3 <<k
, resulting in the accumulation
2
of a stable glycosyl-enzyme intermediate. A more detailed interpretation of kcat, Km
and kcat/Km in terms of the rate constants shown in equation [1], and an explanation of
the assumptions made above is presented in a separate section (see Appendix).
In addition to analysing the kinetic parameters with the substrates as described
above, the ability to restore activity to a mutant generated by site-directed mutagenesis
could yield information into the role the particular amino acid residue plays in catalysis.
Cupples et at. (1990) reported that the anionic nucleophile sodium azide enhanced the
rate of hydrolysis of o-nitrophenyl r3-D glucoside (ONPG) by E461 mutants of E. coli
-
galactosidase, another retaining glycosidase. It has subsequently been shown by use of
a mechanism-based inactivator that the nucleophile in E. coli f3-galactosidase is E537. It
is suggested that a probable role for E461 is as the general acid/base catalyst in this
enzyme (Gebler et a!., 1992a). Since [3-galactosidase hydrolyses f3-1,4-glycosidic linkages
by the same (retaining) mechanism as Cex (Gebler et a!., 1992b) it is likely that sodium
azide, or other anionic nucleophiles could enhance the rate of such a mutant in Cex.
1.6 Expression of the cex gene
A necessary first step for the study of structure-function relationships of proteins
is to have a system in place which facilitates high level production and isolation of the
protein product under investigation. The cex gene has been successfully subcloned and
expressed in E. coli strains from the E. coli lacZ promoter using pUC-based vectors. In
E. coli, Cex is produced as a soluble polypeptide, exported to the periplasm, correctly
processed and disulfide bonded (Gilkes et a!., 1.991a). The protein can be purified from
16
the periplasm or from whole-cell extracts of E. coli by affinity chromatography on
cellulose (Gilkes et a!., 1991a; this study). Typical yields of purified wild-type Cex
obtained from overnight batch cultures of E. coli JM1O1 [pUC12-1.lcex (PTIS)] are about
5-20 mg/L of culture. E. coli currently remains the microbial system of choice for the
production of heterologous proteins from cloned genes. It would be useful however, to
employ a host strain which would secrete the protein directly into the culture medium.
Gram-positive bacteria have a single cell membrane. Consequently, proteins which are
exported across the cell membrane are secreted directly into the culture medium.
Streptomyces species are Gram-positive Actinomycetes which are well noted for their
production of antibiotics and extracellular enzymes (Brawner et a!., 1991). Some
naturally-occurring extracellular enzymes secreted by Streptomyces include
polysaccharidases such as cellulases (Nakai et al., 1988) and xylanases (Shareck et a!.,
1991). Of the approximately 500 species of Streptornyces known to date, the best
characterized is S. coelicolor. As a host for the production and secretion of heterologous
polypeptides, however, S. lividans is the most widely used (Hopwood et a!., 1985;
Ghangas and Wilson, 1989; Steiert et a!., 1989; Taguchi et a!., 1989; Anne and Van
Mellaert, 1993). Unlike other Streptornyces species, S. lividans is useful as a host for the
expression of foreign DNA because it apparently lacks an endonuclease restriction
system (Kieser and Hopwood, 1991). In addition, cloning vectors and protocols for
efficient protoplast transformation (> i0
7 transformants/.ig DNA) have been fairly well
established (Hopwood et a!., 1985). S. lividans may be particularly well suited as a host
for the expression of C. fimi DNA. C. fimi is distantly related to Streptomyces lividans
(Stackebrant and Woese, 1981). The genomes of C. firni and S. lividans are extremely
rich in G+C, each with an average of about 75%. Hence, both organisms have an
extremely biased codon usage of G or C in the third position. Based on the above, S.
lividans appeared to be an attractive host for expression of the C. fimi cex gene.
17
1.7 Objectives
The overall objective of this study was to contribute to a better understanding of
the structure-function relationships in the exoglucanase/xylanase (Cex) from
Cellulomonas fimi. The approach was, firstly, to explore the use of Streptomyces lividans
as a host for the expression of the C. fimi cex gene. The main part of the study involved
the characterization of mutants of Cex generated by site-directed in vitro mutagenesis.
Residues were targeted on the basis of amino acid sequence alignments of family F
1-
1,4-glycanases and on knowledge of the types of residues likely to be involved in
hydrolysis of a 13-1,4-glycosidic linkage by a retaining enzyme. Detailed kinetic analyses
were carried out on various substrates in an attempt to identify both the catalytic
nucleophile and the acid/base catalyst in Cex. Following the initiation of this study, the
nucleophile in Cex was identified as Glu 233 by use of the mechanism-based inactivator
2F-DNPC as mentioned in section 1.5. (Tull et a!., 1991). Therefore, the major thrust of
this study became identification of the acid/base catalyst.
18
2. Materials and Methods
2.1 Buffers, enzymes and media components.
Buffers and solutions used in this study were prepared as described by
Sambrook et a!. (1989). All buffer chemicals were obtained from Sigma. Restriction
endonucleases, polymerases, ligase and nucleotides were from Pharmacia or New
England Biolabs (NEB). SequenaseTM was from United States Biochemical (USB). CF-I
cellulose was from Whatman. Avicel (type PH-i 01) was from FMC International.
Substrates 2”,4”-dinitrophenyl 13-D-cellobioside (2,4-DNPC), 2’,4’-dinitrophenyl -D
glucoside (2,4-DNPG), 4” -bromophenyl 13-D-cellobioside (4-BrPC), and 2 “,4”
dinitrophenyl 2-deoxy-2-fluoro 13-D cellobioside (2F-DNPC) were a gift from Dr. Steve
Withers, Dept. of Chemistry, UBC. 13-cellobiosyl-azide and f3-glucosyl-azide were
synthesized by Dr. Thisbe Lirtdhorst, Dept. of Chemistry, UBC. Cellobiose, p
nitrophenyl (3 -D-glucoside (PNPG) and p-nitrophenyl [3 -D-cellobioside (PNPC) were
obtained from Sigma. All media components were obtained from Difco.
2.2 Bacterial strains, plasmids, and phage.
The bacterial strains, plasmids and phage used in the study are described in
Tables 2.1 and 2.2. Bacterial stocks were maintained at -70°C in LB medium containing
10% DMSO. Plasmid DNA was stored in TE buffer or water at -20°C. Phage were
stored in TYP medium at 4°C.
19
Table 2.1. Bacterial strains
Bacterial strain
Genotype
Reference or source
E. coli JMIO1
supE thi z(1ac-proAB)[F’ traD36 proAB
lacIZAMl5]
Yanisch-Perron et a!.,
1985
E. coli RZ1032
HfrKL16 P0/45 [lysA(61-62)j dutl
ungl thu relAl Zbd-279::TnlO supE44
Kunkel et a!., 1987
S. lividans TK64
pro-2, str-6
Hopwood et a!.,
1985
S. lividans TK24
str-6
Hopwood et a!.,
1985
2.3 Media and growth conditions
Luria-Bertani (LB) medium was described previously (Sambrook et at., 1989).
TYP medium contained the following (per litre): 16 g tryptone, 16 g yeast extract, 5 g
NaC1, 2.5 g 4
HPO adjusted to pH 7.0. Small-scale cultures of E. coli containing pUC
2
K
,
or pTZ-based vectors were grown in shake flasks at 225 rpm at 30°C or 37°C in LB or
TYP supplemented with ampicillin (Amp) at 100 !lg/mL. Generally, small-scale S.
lividans cultures were grown in LB medium in baffled shake flasks at 30°C and 250 rpm.
Flask volumes were limited to 10% of capacity. With pIJ702 or pIJ68O vectors,
thiostrepton (Th) was included at 5 j.tg/mL. S. lividans transformants were selected on
regeneration yeast extract agar (R2YE, Hopwood et at., 1985) overlaid with 0.3%
nutrient agar supplemented with thiostrepton at 50 jg/mL. Solid media contained
1.5% agar.
20
Table 2.2. Plasmids, phagemids and phage
Plasmid, phagemid
or phage
Relevant characteristics
pUC12-1.lcex
contains 2.5 kb C. fimi DNA; expresses O’Neill et at., 1986b
cex from lac promoter; Apr
pUCI2-1.lcex(PTIS)
contains 1.8 kb C. fimi DNA; expresses O’Neill et at., 1986a
cex from inc promoter; contains
portable translation initiation site
(PTIS); Apr
pTZ18R
contains Fl on; Apr
Mead et at., 1986;
Pharmacia
pTZ18R-cex
as above; contains 1.8 kb C. fimi DNA
This study
Streptomyces expression
Hopwood et at.,
1985
p1J702
vector; Th
Reference or source
p1J702-cex
as above, contains entire pUC12-1.lcex This study
plasmid; expresses cex from mel
promoter
p1J680
Streptomyces expression
vector; Thr
Hopwood et at.,
1985
p1J680-cex
as above, expresses cex from aph
promoter; contains entire
pUC12-1.lcex plasmid
M13K07
helper phage for preparation of single- Vieira and Messing,
1987
stranded DNA; Kmr
This study; MacLeod
et at., 1992
21
Growth of E. coli was monitored spectrophotometrically by A600. Growth of
Streptomyces was monitored as follows: Mycelium dry weight was determined by
washing mycelial pellets in distilled water, followed by filtration through Millipore
type HA filters (Bedford, Maryland). The cell mass was dried for about 4 days at 37°C.
2.4 Recombinant DNA techniques
Most recombinant DNA techniques were carried out essentially as described by
Sambrook et a!. (1989). Double-stranded plasmid DNA was isolated from E. coli by the
small-scale alkaline lysis method (Sambrook et al., 1989). Plasmid DNA was isolated
from Streptomyces, by the boiling method essentially as described by Kendall and
Cullum (1984). Restriction endonuclease digestions and ligations were carried out
according to the directions of the manufacturer& in the buffers provided.
Electrophoresis of DNA generally employed 0.8% agarose gels in TBE buffer
(Sambrook et al., 1989). DNA fragments excised from agarose gels were purified using
the Gene Clean II Kit (BiolOl, La Jolla, Ca) according to the manufacturer’s directions.
Streptomyces protoplasts were prepared and transformed as described by Hopwood et
a!. (1985). Preparation and transformation of competent E. coil strains was performed
as described by Hanahan (1983).
2.4.1 Production and isolation of single-stranded DNA.
Single-stranded template DNA was obtained as follows: Single, overnight
colonies of E. coli JM1O1 containing pTZ18R-based constructs were inoculated into 2mLs of TYP medium containing 100 tg/mL ampicillin and 10 PFU/mL M13K07
helper phage. Following 1 h incubation at 35°C, kanamycin was added to 70 j.tg/mL.
22
Cultures were grown overnight at 35°C and 225 rpm. Cells were removed by
centrifugation in a microfuge for 5 mm at room temperature. The phagemid were
precipitated at 4°C with 1.7 M ammonium acetate and 12% (w/v) PEG-6000.
Single-stranded DNA was isolated from the phagemid as follows (D. Trimbur,
personal communication). After centrifugation for 10 minutes at 4°C the supernatant
was removed and the phage pellet was resuspended in 20 iL of 10 mM Tris-HC1, 1 mM
EDTA pH 8.0 (TB). 200 pL of 4 M NaC1O
4 were added and the mixture was incubated
for 5 mm at room temperature. Glass fibre filter disks (GF/C, Whatman) were placed
in the bottom of a microtitre plate in which the wells had been pierced with a hot
needle. The DNA was bound to the glass fibre by vacuum filtration through the
microtitre plate. The bound DNA was washed thoroughly with 70% ethanol and air
dried for 5 minutes. The filter disks were transferred to 0.5-mL Eppendorf tubes
pierced at the bottom, which were in turn placed inside 1.5-mL Eppendorf tubes. The
DNA was eluted from the filter disks with 20 iL of H
0 and captured by centrifugation
2
for 30 s.
2.4.2 Site-directed in vitro mutation of cex
Site-directed in vitro mutation was performed essentially as described by Kunkel
et al., 1987. Single-stranded, uracil-containing DNA templates (pTZ18R-cex) were
produced as described in section 2.4.1 with the following modifications: E. coil RZ1032
was used as the host strain and uridine was included in the medium at 1 j.tg/mL.
Synthetic oligodeoxyribonucleotide primers used for mutagenesis were prepared
(Atkinson and Smith, 1984) by the UBC Nucleic acid and Protein synthesis unit (NAPS)
with an Applied Biosystems 380A DNA synthesizer. Oligonucleotides were purified
by precipitation with n-butanol as described by Sawadogo and Van Dyke (1991). The
oligonucleotides (25mers) employed in this study are described in Table 2.3.
23
2.4.3 Sequencing of DNA
Sequencing of plasmid DNA used the modified dideoxy chain termination
method described previously (Tabor and Richardson, 1987) with the following
modifications: in the primer extension reaction, T7 DNA polymerase (Sequenase)
was used, the reaction temperature was increased to 43°C and 7-deazaGTP was
substituted for dGTP.
Table 2.3. Oligonucleotides used in site-directed mutagenesis of cex
Oligonucleotide sequence*
Mutation in Cex
5’P-G CGC ATC ACC GAC CTC GAC ATC CGC-Y.
E233D
5’P-G CGC ATC ACC CAG CTC GAC ATC CGC-3’.
E233Q
5P-AC GTC GTC AAC GAC GCG TTC CCC GA-3’
E127D
5’P-AC GTC GTC AAC CCG GCG TTC GCC GA-3’
E127A
5’P-AC GTC GTC AAC GGC GCG TTC CCC GA-3
E127G
5’P-TC GCG TCG TGG GCC GTC GTC AAC GAG-3’
D123A
*P indicates the oligonucleotide is 5’ phosphorylated
24
2.5
Detection of cex gene expression
2.5.1 MUCase and PNPCase assays
E. coli or S. lividans transformants expressing exoglucanase activity were
detected as follows. Cells were plated onto LB agar containing the appropriate
antibiotic and 100 jiM methylumbelliferyl 13-D- cellobioside (MUC). Following growth
overnight at 30°C, colonies expressing exoglucanase activity fluoresced under UV light
(365nm). This assay was also used as an initial screen for Cex mutants. Following sitedirected mutagenesis, non-fluorescing colonies, indicating loss of exoglucanase activity,
were isolated and plasmid DNA was prepared for sequencing.
Cex activity in crude-cell extracts or culture supernatants could be estimated by
using the following assay (Gilkes et al., 1984b): Culture supernatant or diluted cell
extract (0.5 mL) was added to 0.5 mL of 12.5 mM PNPC to give a final PNPC
concentration of about 10 x Km as determined previously for Cex (Gilkes et a!., 1984b).
The mixture was incubated at 37°C for about 15 mm or until a yellow colour appeared.
The reaction was stopped by the addition of 1 M 3
CO (0.5 mL) and the absorbance
2
Na
at 400 nm was determined. One unit of activity was defined as that which released 1
jimol PNP per minute (Gilkes et a!., 1984b).
2.5.2 Binding to Avicel
When the activity of the enzyme was unknown (for example, with the Cex
mutants), the levels of gene expression could be approximated from crude-cell extracts
or culture supernatants as follows: Approximately 100 tL of cell extract (described in
section 2.7.2) or 0.5 mL of culture supernatant was added to 20 mg of Avicel
(microcrystalline cellulose) in a final volume of 1 mL 50 mM phosphate buffer, pH 7.0.
The protein was adsorbed to the Avicel by mixing for 1 h at 4 °C. The Avicel was
25
centrifuged for 30 s, washed once with 500 pL IM NaC1 in phosphate buffer, then once
with phosphate buffer. The Avicel was collected by centrifugation, then boiled for 2
mm in SDS loading buffer (Laemmli, 1970). The bound polypeptides were analyzed by
SDS-PAGE (section 2.6.1). An estimation of mutant protein yield could be made by
comparison with a known quantity of wild-type Cex.
2.6 Detection of protein
2.6.1 SDS-PAGE and Western blotting
Proteins were resolved by 0.1% sodium dodecyl sulfate-10% polyacrylamide gel
electrophoresis (SDS-PAGE) as described previously (Laemmli, 1970; Schägger and
Von Jagow, 1987) using a Bio-Rad MiniPROTEANTM apparatus. Molecular weight
standards (Sigma) were as follows (kDa): Myosin, 212; f3-galactosidase, 130;
phosphorylase B, 97.4; bovine serum albumin, 68; catalase, 57; glutamate
dehydrogenase, 53; alcohol dehydrogenase, 45; ovalbumin, 41; glyceraldehyde-3phosphate dehydrogenase, 36; carbonic anhydrase, 29; bovine trypsin inhibitor, 20;
cytochrome C, 12.4. Protein bands were visualized by staining with Coomassie blue
(Meril, 1990). Western blots were performed as described previously (Towbin et at.,
1979). Western blots were probed with rabbit anti-Cex serum (1/4000 dilution) using
goat anti-rabbit IgG-alkaline phosphatase conjugate (BRL) as the secondary antibody
(Gilkes et at. , 1988) at 1/7000 dilution. The detection reagents were 5-bromo-4-chloro3-indolyl-phosphate and nitroblue tetrazolium dye (Sigma). Prestained molecular
weight standards were from BRL or Bio-Rad.
26
2.6.2 Glycoprotein detection
Glycoproteins in SDS-PAGE gels were detected with the periodic acid-Schiff
reagent (Zacharius et a!., 1969). Glycoproteins were detected following western blotting
by probing the blots with concanavalin A-horseradish peroxidase (ConA-HRP;
Seikagaku). The blots were developed with HRP development reagent (Bio-Rad).
2.6.3 N-terminal amino acid sequencing
N-terminal amino acid sequences were determined at the University of Victoria’s
protein sequencing unit by automated Edman degradation, using an Applied
Biosystems 470A gas-phase sequenator.
2.6.4 Determination of protein concentration
Protein concentrations were determined by dye binding (Bradford, 1976) using
the Bio-Rad protein kit, with bovine serum albumin as the standard. The concentration
of purified Cex was determined spectrophotometrically from A280. The extinction
coefficient of Cex
1 mg/mL
2
(E
=
1.61) was determined experimentally by the method of
Scopes (1974).
2.7
Production and purification of recombinant Cex
2.7.1 Production of Cex in Streptomyces lividans TK64
S. lividans TK64 (250 .iL from a frozen stock culture) containing the cex gene
encoded on p1J702-cex or p1J680-cex was inoculated to 10 mL LB medium supplemented
with 5 jig /mL thiostrepton. Cultures were incubated for 3 days at 30°C, then added to
100 mL of the same medium. After growth overnight at 30°C and 225 rpm, the culture
was added to 10 L of the same medium in a Chemap FZ3000 fermenter. After growth
for 45 h at 300 C, 600 rpm and 5 L/min aeration, the mycelium was removed by
27
passage of the culture through a Sharples centrifuge. Cex was purified from the culture
supernatant as described in section 2.7.3.
2.7.2 Production of Cex and Cex mutants in E. coli
E. coil containing the plasmid pUC12-Llcex(PTIS) encoding wild-type Cex or
mutants of Cex were grown in 2L-cultures in shake flasks or in a 60 L Chemap FZ3000
fermenter. Cultures were grown in LB supplemented with ampicillin at 100 .ig/mL.
At A600
=
1.0, IPTG was added to 0.1 mM and growth was allowed to continue
overnight. The cells were collected by centrifugation at 10 000 x g for 10 mm at 4 °C.
The cell pellet was resuspended in approximately 1/100 of the original volume in 50
mM phosphate buffer pH 7.0 (phosphate buffer) at 4 °C. Cells were ruptured by
passage twice though a French pressure cell. PMSF was added to 0.5 mM, Pepstatin A
to 1 iM and EDTA to 1 mM. The crude-cell extract was clarified by centrifugation at 40
000 x g for 30 mm at 4°C. Streptomycin sulfate was added to 1.5% and the mixture
stirred at 4 °C overnight. The extract was centrifuged again at 40 000 x g for 30 mm at
4°C. Cex was purified from the clarified cell extract as described below.
2.7.3 Purification of Cex
Culture supernatant from Streptomyces or clarified cell extract from E. coil was
stirred with CF-I cellulose (Sigma) (about ig CF-1/mg Cex) in phosphate buffer for 3 h
at 4 °C. The cellulose was recovered by vacuum filtration on a glass fibre filter disk
(GF/C, Whatman), washed twice with 500 mL I M NaCl in phosphate buffer and
twice with 500 mL phosphate buffer. The cellulose was packed into a column (5 cm X
25 cm) and attached to an FPLC system (Pharmacia). Washing was continued by
passing a column volume of phosphate buffer through the column at a flow rate of 1
mL/min In the case of Cex produced in E. coli, adsorbed polypeptides were eluted
28
with distilled water at a flow rate of 1. mL/min. In the case of Cex produced in
Streptomyces, adsorbed polypeptides were eluted with a linear gradient of 0-8.0 M
guanidinium hydrochloride (Gdm HC1) in phosphate buffer, total volume 500 mL at a
flow rate of 1 mL/min. In both cases, the absorbance of the eluate was measured
continuously at 280 nm and peak fractions were assayed for Cex activity with PNPC
(section 2.5.1) The appropriate fractions were pooled and centrifuged at 40 000 x g for
30 mm at 4°C to remove any cellofines. The supernatant was concentrated to greater
than 1 mg protein/mL by diafiltration through an Amicon PM1O membrane. For Cex
from Streptomyces, the Gdm HC1 was replaced with phosphate buffer during the
diafiltration step such that the final concentration of Gdm HC1 was calculated to be less
than 0.5 jiM. The purified polypeptides were analyzed by SDS PAGE (section 2.6.1).
Final protein concentration was determined by A280 as described in section 2.6.4.
Purified protein solutions were passed through a 0.22 jim filter (Millipore) and then
stored at 4°C.
2.8
Proteolysis of Cex from S. lividans and E. coli
2.8.1 Substrate-bound Cex
C. fimi protease, generously provided by Shen Hua, was prepared as described
previously (Gilkes et a!., 1988). Purified Cex from S. lividans or from E. coli was
adsorbed to 5 mg Avicel as described in section 2.5.2 then resuspended in 25 jiL 20 mM
Tris.HC1, pH 7.5. After adding 0.5 units* of C. firni protease, the suspensions were
incubated at 37°C for 24 h. The Avicel was recovered by centrifugation and bound
polypeptides were analyzed by SDS-PAGE as in section 2.5.2. The supernatants,
containing polypeptides released from the Avicel by protease treatment, were also
analysed by SDS-PAGE.
*
(1 unit is defined as that which will release 1.0 0D
/h @
585
37°C, pH 7.5 from cowhide powder azure).
29
2.8.2 Cex in solution
Purified Cex from E. coli, S. lividans and C. fimi, was incubated with C. fimi
protease at 37°C for 24 h in 20 mM Tris.HC1, pH 7.5. The samples were analyzed by
SDS-PAGE as above.
2.9
Enzyme kinetics
2.9.1 Determination of steady-state kinetic parameters
Michaelis-Menten parameters for aryl cellobiosides and aryl glucosides were
determined by continuous spectrophotometric measurement of the release of p
nitrophenol, dinitrophenol or p-bromophenol using a Hitachi U 2000
spectrophotometer with a temperature-controlled cell holder. Reactions were carried
out in 50 mM phosphate buffer (pH 7.0), 37°C and contained BSA at I mg/mL where
indicated. Reaction mixtures were pre incubated within the water-circulated cell
holder at the appropriate temperature for a period of 10 mm prior to addition of
enzyme. Where indicated, sodium azide was included in reactions at the
concentrations shown. For each substrate to be tested, the appropriate concentrations
of substrate to be used was estimated by monitoring the initial rates of hydrolysis at
three different substrate concentrations and then estimating the Km value from a Hanes
plot. Initial rates of enzyme-catalysed hydrolysis for a particular substrate were
measured at 6 to 10 different substrate concentrations ranging from about 1/7 x Km to 7
x K where practical. Incubation times ranged from 5 minutes to 18 hours (in the case
of 4-BrPC). Values for Km and kcat were determined from the initial rate of hydrolysis
) vs. substrate concentration, by non-linear regression analysis using the computer
0
(V
program Grafit 2.0 (Erithacus Software Ltd., Staines, U.K.). An example calculation of
Km and kcat is presented in the Appendix.
30
The substrates used in this study, along with the wavelengths monitored and
extinction coefficients for the corresponding aglycon units, are shown in Table 2.4.
Table 2.4. Wavelengths monitored and extinction coefficients for hydrolysis of aryl
cellobiosides and aryl glucosides.
Substrate
Aglycon unit
Wavelength
monitored (nm)
Extinction coefficient (AE)
.cm at given
(M
)
1
wavelength, pH 7.0
PNPC
PNP
400
7280
PNPG
PNP
400
7280
2,4-DNPC
DNP
400
10900
2,4-DNPG
DNP
400
10900
4-BrPC
PBrP
288
1 120
2.9.2 Kinetics of Inactivation of Cex mutants with 2F-DNPC.
Inactivation of Cex mutants with 2F-DNPC was performed essentially as
described previously for inactivation of wild-type Cex by 2F-DNPG (Tull et a!., 1991).
The enzyme was incubated at 37°C in 50 mM phosphate buffer (pH 7.0) containing 2FDNPC at the concentrations indicated. At various times, 10 iiL aliquots of the
inactivation mixture (as described above) was transferred into a solution of 2,4-DNPC
(100 tM) and sodium azide (60 mM) in 50 mM phosphate buffer (pH 7.0; 37°C; final
volume 500 j.iL). Enzyme activity was determined by continuous spectrophotometric
measurement of the release of dinitrophenol using a Hitachi U 2000 spectrophotometer
31
with a temperature-controlled cell holder. For each inactivator concentration, residual
enzyme activity (v/v
) at various time points was calculated by comparison with the
0
activity of the control sample which contained no inactivator. From the slope of a plot
of in [V/Vol for each inactivator concentration, pseudo-first order rate constants (kobs)
were calculated. The inactivation rate constant (kj) and the equilibrium binding
constant (1(i) can be determined from a double-reciprocal plot (Lineweaver-Burk) of
kobs vs. inactivator concentration. This curve is presented in the results section for
display purposes and was not used for calculations. The inactivation rate constant (ki)
and the equilibrium binding constant (Ki) were determined from kobs vs. inactivator
concentration by non-linear regression analysis using the computer program Grafit.
2.10 Characterization of the products of enzymatic hydrolysis
Products were characterized from reactions generally containing 100 j.Lg/mL
mutant Cex or 1 j.tg/mL wild-type Cex and 6 mM substrate (as indicated) in phosphate
buffer, incubated overnight at 37°C. Sodium azide was included where indicated.
2.10.1 Thin layer chromatography
Thin layer chromatography was performed on 0.2 mm silica gel aluminum
plates (#60 F254; E. Merck). Approximately 2 !IL of each reaction mixture or standard
was applied to the base of the TLC plate. The plate was transferred to an Erlenmeyer
flask and dried under vacuum for about 5 mm. The chromatography was performed
with a mixture of ethyl acetate/methanol/water (7:2:1 v/v/v) and allowed to air dry
for 5 mm. The chromatographs were dipped in a solution of 10% 4
S0 in methanol
2
H
and heated with a Red Devil heat gun until the reaction products were visible. f3cellobiosyl-azide, f3-glucosyl-azide, cellobiose and glucose were the standards used.
32
2.10.2 1
H-NMR spectrometry of the products of enzymatic hydrolysis
Enzymatic reactions were carried out as described above. Prior to H-NMR, the
enzyme was removed by filtering the solution through a 10 kDa cutoff polysulfone
membrane (ultrafree-MC; Millipore). Samples were prepared for 1
H-NMR analysis by
Dr. Thisbe Lindhorst, Department of Chemistry, UBC, by repeatedly dissolving in 2
D
0
,
freeze drying several times, and finally dissolving in 2
D
0
. Spectra were recorded with
a Bruker 400 MHz spectrometer at the NMR Facility, Department of Chemistry, UBC,
with the assistance of Dr. Thisbe Lindhorst.
2.11 Protein Molecular Mass determination
Protein molecular mass was predicted from amino acid sequence data using the
MacProMass computer program (Beckman Research Institute, Duarte, CA). Actual
protein molecular mass was determined by Ion spray mass spectrometry using a
PESCIEX API III ion spray LC/MS system by Dr. Shichang Miao, Department of
Chemistry, UBC. Where indicated, the enzyme was incubated with substrate (2,4DNPC) for 1 mm at 37 °C prior to mass spectrometric analysis.
2.11 Amino acid sequence alignments and database searches
Amino acid sequences were aligned on an IBM-compatible computer using
PCGENETM (Intelligenetics, Mountainview, CA). Amino acid sequence information for
family-F cellulases and xylanases was obtained through electronic mail
([email protected]) from the GenBank and Swiss-Prot databases at the National
Center of Biotechnology Information (NCBI), National Library of Medicine, NIH, in
Bethesda, MD.
33
3. Results (part 1): Expression of the cex gene in S. lividans
3.1 Construction of p1J702-cex and p1J680-cex.
Plasmids for expression of cex in Streptomyces were constructed as outlined in
Figures 3.1 and 3.2, respectively. In p1J702-cex, the cex gene and its promoter are
situated downstream from the tyrosinase or “melanin” (mel) promoter of p1J7O2. In
p1J680-cex, the cex gene and its promoter are situated downstream from the S. fradiae
aminoglycoside phosphotransferase (a ph) promoter of p1J680. A construct was also
obtained in which the C. fimi DNA was inserted in the reverse orientation (p1J702-xec,
not shown). The constructs p1J702-cex and p1J680-cex, contained pUC12-1.lcex in its
entirety permitting replication in both Streptomyces and E. coil. E. coli JM1O1 was
transformed with the ligation mixture and clones were selected on LB medium
supplemented with ampicillin. The desired constructs were isolated from E. coil by
small-scale alkaline lysis (Sambrook et a!., 1989) and then used to transform S. lividans
TK64 protoplasts as described by Hopwood et a!. (1985). Streptomyces transformants
were selected on R2YE solid medium supplemented with thiostrepton and MUC.
Transformants hydrolysing MUC (expressing exoglucanase activity) fluoresced under
UV light. The presence of p1J702-cex or p1J680-cex was confirmed by analysing plasmid
DNA (isolated as described in section 2.4) by agarose gel electrophoresis.
3.2 Production of Cex by S. lividans TK64 [p1J702-cex]
As indicated in Figure 3.3, exoglucanase (PNPCase) activity was secreted into the
culture supernatant by S. lividans TK64 containing p1J702-cex. Exoglucanase activity in
the culture supernatant peaked at 0.008 units/mL culture following about 45 hours of
growth. This represented less than 1 mg of active Cex per litre of culture, based on the
34
Figure 3.1. Construction of p1J702-cex. p1J702 (Hopwood et a!., 1985) was isolated from
S. lividans TK24 by the method of Kendall and Cullum (1984). pUC12-1.lcex, containing
the exoglucanase (cex) gene from Cellulornonasfirni, was described previously (O’Neill et
a!., 1986a). Plasmid p1J702-cex (11.1 kb) was constructed by ligating Barn HI cut pUC121.lcex (5.3 kb) into the Bgl II site of p1J702 (5.8 kb). This resulted in the destruction of
both sites (*)
35
BamHl
89111
Barn Hi
BgI II
T4 DNA ligase
36
Figure 3.2. Construction of p1J680-cex. p1J680 (Hopwood et al., 1985) was isolated from
S. lividans TK24 by the method of Kendall and Cullum (1984). Plasmid p1J680-cex (10.6
kb) was constructed by ligating Barn HI cut pUC12-1.lcex(5.3 kb) into the Barn HI site of
p1J680 (5.3 kb).
37
BamHI
BamHI
BamHI
T4 DNA ligase
38
0.010
-J
E
D
>
4-,
>
4-,
C)
c
ci
U)
0.004
C)
D
x
w
0.002
0.000
20
40
60
80
100
120
Time (h)
Figure 3.3. Production of exoglucanase by S. lividans TK64[p1J702-Cexl. S. lividans TK64
containing plasmids p1J702-cex (0), p1J702-xec () or p1J702(D) was grown in 100 mL of
LB supplemented with thiostrepton (5 jig /mL) in a 1-L baffled shake flask at 30°C, 225
rpm. The initial cultures had an A600 = 0.01. Samples of 5 mL were withdrawn at
approximately 3-h intervals, and culture supernatants were collected by centrifugation
for 7 mm at 3000 x g. Exoglucanase activity was determined from culture supernatants
as described in section 2.5.1.
39
specific activity of the wild-type enzyme. Production of exoglucanase (PNPCase)
activity in S. lividans TK64 [p1J702-xec], in which the C. firni fragment (including the cex
gene and its promoter) was inserted in the reverse orientation, was only about 10% of
that containing p1J702-cex. Only trace amounts of exoglucanase activity could be
detected from culture supernatants of S. lividans TK64 containing p1J702 without an
insert.
3.3 Production of Cex by S. lividans TK64 [p1J680-cexl
Exoglucanase (PNPCase) activity was secreted into the culture medium by S.
lividans TK64 [p1J680-cexl as indicated in Figure 3.4. Maximum activity in the culture
supernatant (0.06 units/mL) was reached after about 40 h of growth. Based on activity,
this represents about 5.5 mg of Cex per litre of culture, an 8-fold increase over that
produced by S. lividans TK64 [p1J702-cex]. As a fraction of total protein present in the
culture medium, the activity peaked at 3.3 units/mg (about 14% of total secreted
protein) following 35 h of growth. This value subsequently declined because of
additional protein appearing in the culture medium during the late log and stationary
phases of growth.
As shown in Figure 3.5A, p1J680-cex encoded a 49 kDa polypeptide (p
) which
49
was the major polypeptide in the culture supernatant. The proportion of p49 in the
culture supernatant increased with time, consistent with an increase in exoglucanase
activity with time as indicated in Figure 3.4. The identification of p49 as Cex was
confirmed by its reaction with rabbit anti-Cex as indicated in Figure 3.5B. The 49 kDa
polypeptide could be purified from the culture supernatant by adsorption to Avicel
(Figure 3.5A, lanes 6 and 7) indicating the presence of a functional cellulose-binding
0.060
0.070
0.000
10
0.010
0.020
0.030
0.040
20
30
50
Time (h)
40
60
70
80
0
90
v
30
a)
—S
C
0.0
n
I-v
2.0
‘)ñ
J.u
4.0
_j
.
>
ca
I—
0
-I
-E
0
+
a)
—S
Figure 3.4. Production of exoglucanase by S. lividans TK64 [p1J680-cexJ. S. lividans TK64 [p1J680-cexl was grown
in
100 mL of LB supplemented with 5 .tg Th/mL in a 1-L baffled shake flask at 30°C, 225 rpm. The initial culture
had an A600 = 0.01. Samples of 5 mL were withdrawn at approximately 3-h intervals, and culture supernatants
were collected by centrifugation for 7 mm at 3000 X g. Exoglucanase activities were determined from culture
supernatants as described in section 2.5.1. Protein concentrations were determined by dye-binding as described in
section 2.6.4. Mycelium dry weight measurements were determined as described in section 2.3.
Cl)
a)
ct
C.)
>
•.. 0.050
0
C
41
Figure. 3.5. Identification of extracellular Cex in S. lividans [p1J680-cex] cultures.
S. lividans cultures were grown as described in Figure 3.4. Samples of culture were
removed at various time intervals, the mycelium removed by centrifugation and 500 jiL
of the supernatants made 10 % in trichioroacetic acid. The precipitated proteins were
collected by centrifugation and analysed by SDS-PAGE. A further 500 jiL sample of the
supernatant was shaken with 12.5 mg Avicel (microcrystalline cellulose) for 1 h at 4°C.
Polypeptides which adsorbed to the Avicel were analyzed by SDS-PAGE as described
in section 2.5.2. Panel A: Gel stained with Coomassie blue. Panel B: Western blot
(samples prepared as above) probed with rabbit anti-Cex serum (see section 2.6.1).
Lanes: 1, molecular weight standards as indicated (kDa); lanes 2 and 8, 2 jig and 0.5 p.g
Cex purified from E. coli (section 2.7.3); lanes 3 and 9, supernatant from S. lividans TK64
after 4 h growth; lanes 4 and 10, supernatant from S. lividans TK64 [p1J680-cexl after 20 h
growth; lanes 5 and 11, supernatant from S. lividans TK64 [p1J680-cexl after 40 h growth;
lanes 6 and 12, unbound polypeptides after treatment of the 40 h supernatant from S.
lividans TK64 [p1J680-cex] with Avicel; lanes 7 and 13, bound polypeptides after
treatment of the 40 h supernatant from S. lividans TK64 [p1J680-cex] with Avicel.
(‘4
CD
to
I
hUT
1J 1’IH
C4JO o LOC)LO,—CD
CDF0C)
CDLO
C)
0
0)
43
domain. The other major polypeptide (p42) present in the culture supernant (lanes 4
and 5), reacted with rabbit anti-Cex (lanes 10 and 11), but did not adsorb to Avicel (lane
6). Therefore, this polypeptide is probably a degradation product of Cex which is
lacking a functional cellulose-binding domain.
Cex produced from S. lividans had an apparent molecular mass 2 kDa more than
that of the polypeptide produced from E. coli. Determination of the N-terminal amino
acid sequence of the polypeptide from S. lividans gave two sequences in approximately
equal proportions (section 2.6.3), Ala-Thr-Thr-Leu-Lys (ATTLK) and Gln-Ala-Ala-Thr
Thr (QAATT), indicative of leader peptide processing at two adjacent sites, i.e., between
A(-1) and A(+1) and between A(-3) and Q(-2) respectively (Table 3.1). In E. coli and in C.
fimi, Cex is processed at a single site between A(-1) and A(+1) (O’Neill et a!. 1986c).
,
The additional amino acids were insufficient to account for the difference in the
apparent molecular masses.
Table 3.1. N-terminal processing of native and recombinant Cex
Source of Cex
Site(s) of processing
:
1
-3 -2 -1 ÷1 +2 +3 +4 ÷5
2
E.coliJMlOl
2
C.fimi
S. lividans TK64
A+A T T L K.
A+A T T L K.
A+A T T L K
A+QAATTLK
I
Determined by N-terminal (5-cycle) amino acid sequencing. A (+1) represents the N
terminal amino acid in Cex from C. firni. 2 The N-terminal sequence was determined
previously (O’Neill et a!., 1986c).
44
3.4. Glycosylation of Cex by Streptomyces lividans
It was reported previously that the difference in size between Cex from C. fimi
and that from E. coli is a consequence of glycosylation of the native enzyme (Langsford
et al., 1984; Gilkes et a!., 1988). The enzyme produced in E. coli is not glycosylated.
Although there were no reports of the glycosylation of a heterologous polypeptide by
Streptomyces prior to this study, Streptornyces was known to produce glycoproteins
(Mihoc and Kluepfel, 1990). It appeared reasonable that the difference in size between
the E. coli and S. lividans-produced enzymes were the result of glycosylation of the
latter.
Cex produced by S. lividans [p1J680-cexl and E. coli JMIOI [pUCI2-1.lcex] was
purified to apparent homogeneity by affinity chromatography on cellulose (Fig. 3.6) as
described in Materials and Methods (section 2.7.3). Purified Cex from C. firni was
generously provided by Emily Kwan. The specific activity of the purified enzyme from
Streptomyces was 11.2 U/mg. The specific activities of Cex purified from E. coli and C.
fimi were similar: 12.0 U/mg and 13.1 U/mg, respectively. As shown in Figure 3.6A,
the recombinant polypeptide from S. lividans appeared to be the same molecular mass
as the native, glycosylated polypeptide from C. fimi (49 kDa) as judged by SDS-PAGE.
It was slightly greater than that from E. coli (47 kDa). As revealed by Schiff’s staining
(Figure 3.6B) and by reaction with Concanavalin-HRP (Figure 3.6C), Cex produced by S.
lividans was also glycosylated.
It was determined previously that the glycosyl groups on Cex from C. fimi
protect it from a protease present in C. fimi culture supernatants if Cex is adsorbed to
cellulose, and to a lesser extent when the enzyme is in solution (Langsford et a!., 1987;
Gilkes, N.R., unpublished). In contrast, non-glycosylated Cex from E. coli is susceptible
to proteolysis whether or not the enzyme is bound to cellulose. This yields a major
2
3. 4
567
B
1
8
Figure 3.6. Glycosylation of Cex by S. lividans. Cex produced by S. lividans [p1J680-cex], E. coli JM1O1 [pUC121.lcex] and C. fimi were purified to apparent homogeneity by affinity chromatography on cellulose (section 2.7.3).
The purified polypeptides were analyzed by SDS-PAGE. Panel A: gel stained with Coomassie blue. Panel , gel
stained by the periodic acid-Schiff method to detect glycosylation. Panel , Western blot probed with
concanavalin A-horseradish peroxidase to detect glycosylation. Lane 1: size standards as indicated. Lanes 2, 5 and
8,2 j.tg Cex purified from E. coli; lanes 3, 6 and 9, 2 .tg Cex purified from C. fimi; lanes 4, 7 and 10, 2 p.g Cex
purified from S. lividans.
41’
36’
29’
97.4 S.
68.0%..
57.5
53..-
1
kDa
212\
A
Ui
46
proteolysis product of 35 kDa corresponding to the catalytic domain of Cex (Langsford
et al., 1987; Gilkes et a!., 1991a).
In order to determine whether the glycosyl groups on the enzyme from
Streptomyces were functionally similar in this regard to those of the native enzyme, its
susceptibility to the C. fimi protease was analysed. As shown in Figure 3.7A and 3.7B,
the Streptomyces-produced enzyme behaved identically to the native enzyme. Unlike
the enzyme produced in E. coli, it was not cleaved when adsorbed to cellulose (Figure
3.7A). The Streptomyces-produced enzyme was hydrolyzed when in solution (Figure
3.7B), although more slowly than non-glycosylated Cex from E. coli. Cex from S.
lividans gave a major proteolysis product of 42 kDa, similar to that of Cex from C. fimi.
47
Figure 3.7. Sensitivity of Cex from S. lividans to the protease from C. fimi.
Panel A: Cex adsorbed to cellulose (Avicel). Purified Cex (20 tg) from S. lividans and
from E. coli was each adsorbed to 5 mg Avicel, then resuspended in 20 mM Tris.HC1,
pH 7.5. The suspension was incubated with 0.5 units of C. firni protease at 37°C for 24 h.
Polypeptides bound to the Avicel were analysed by SDS-PAGE. The supernatants,
containing polypeptides released from the Avicel by protease treatment, were also
analysed by SDS-PAGE. Lane 1, molecular weight standards; lanes 2-5, Cex from S.
lividans; lanes 6-9, Cex from E. coli; lanes 2 and 6, bound polypeptides left after protease
treatment; lanes 3 and 7, polypeptides released from the Avicel by protease treatment;
lanes 4 and 8, bound polypeptides without protease treatment; lanes 5 and 9,
unadsorbed polypeptides without protease treatment.
Panel : Cex treated in solution with C. fimi protease. Purified Cex (10 .tg) from E. coli,
S. lividans and C. fimi, was incubated with 0.5 units of C. fimi protease at 37°C for 24 h in
35 .tL 20 mM Tris.HC1, pH 7.5. 10 iL of the S. lividans and E. coli samples and all of the
C. fimi sample was analyzed by 0.1% SDS-10% PAGE. Lanes 10 and 17, molecular
weight standards; lanes 11 and 12, Cex from S. lividans; lanes 13 and 14, Cex from E.
coli; lanes 15 and 16, Cex from C.firni. Lanes 11, 13 and 15, with protease treatment;
lanes 12, 14 and 16 without protease treatment.
48
A
kDa
1
2
3
4
5
6
7
8
9
16
17
130
97.4
—
29’
kDa
B
130
97.4
10
12
13
14
—
Z
—
53
11
kDa
130
—
A.
=
— -53
.-
—
—
—
.—
36 /
29”
15
—
—97.4
68.O
—575
41
36
49
4. Results (part 2): Identification of catalytic residues in Cex
3-1,4-glycanases can be grouped into at least 12 families on the basis of similar
amino acid sequences (Henrissat et at., 1989; Beguin, 1990; Gilkes et at., 1991b; Henrissat,
1991, Shen et a!., 1994). Cex belongs to family F of 3-1,4-g1ycanases (Henrissat et a!.,
1989). Searches of the GenBank database revealed at least 20 enzymes that can now be
assigned to the family (see Introduction, Table 1.1) Some of these enzymes contain
more than one catalytic domain; some contain carbohydrate binding domains unrelated
to that of Cex. To date, three-dimensional information is not available for any enzymes
in family-F. In order to pinpoint potential catalytic residues, the enzymes were aligned
to reveal conserved amino acid residues, The alignment was made with family-F
catalytic domains related to the catalytic domain of Cex. As shown in Figure 4.1, there
are several conserved acidic residues. Three aspartates and three glutamates are
conserved in all members of the family, corresponding to Glu 43, Asp 123, Glu 127, Asp
170, Glu 233, and Asp 277 in Cex. In addition, Asp 235 is conserved in all but one
member of the family.
4.1 Wild-type Cex: kinetics
Cex was purified from cultures of E. coli JM1OI LpUC12-1.1cex] as described in
sections 2.7.2 and 2.7.3. The purified protein obtained by affinity chromatography on
cellulose was >95% homogeneous when analyzed by SDS-PAGE (not shown).
Approximately 30 mg of purified Cex was obtained from a 2-L culture.
The kinetic parameters were determined for hydrolysis of various cellobiosides
and glucosides (Table 4.1). The kcat values for p-nitrophenyl 13-D- cellobioside (PNPC),
and 2”,4”-dinitrophenyl -D-ce1lobioside (2,4-DNPC) and t
,2
4’-dinitrophenyl 13-D-
“---“
“_“
Figure 4.1. Alignment of Family F catalytic domains. Only a portion of each enzyme is indicated. The putative
catalytic nucleophile in Cex is indicated by an “n”. The putative acid/base catalyst is indicated by an “a/b”.
Conserved acidic residues are indicated in bold-face.
means the amino acid sequence is contiguous on either
side of the gap.
means the end of the sequence or that sequence information is incomplete. Accession
numbers (Genbank or SWISS-PROT) are indicated in parenthesis. a) C. fimi Cex (L11080); b) S. lividans XynA
(M64551); c) C. thermocellum XynZ (M22624, P10478); d) A. kawachii XynA (D14847); e) T. aurantiacus XYN [partial
sequence only] (P23360); f) P. chrysogenum Xyn [partial sequence only] (S31307); g) R. flavefaciens XynA (P29126);
h) B. fibrisolvens XynA (P23551); i) B. fibrisolvens XynB (X61495, S55274); j) C. saccharolyticum ORF4 (M34459); k)
Thermophilic bacterial sp rt8.84 XynA (L18965); 1) C. saccharolyticum Ce1B (A43802, X13602); m) C. thermocellum
XynX (M67438); n) Thermoanaerobacter saccharolyticum (strain B6A-RI) Xy1A (M97882); o) Bacillus sp strain C-125
XynA (D00087, P07528); p) C. stercorarium strain F9 (D12504); q) C. saccharolyticum XynA (M34459); r) P. fluorescens
subsp-cellulosa XynB (P23030); s) C. albidus XynA (JS0734); t) P. fluorescens subsp-cellulosa XynA
(X15429).
t
r
s
p
q
c
m
n
o
k
1
j
b
d
e
f
g
h
i
a
t
r
p
q
m
n
o
k
1
j
f
g
h
i
e
d
c
b
a
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
+ +
-
-
-
181
VASWDVVNEAFADG_DGPP
QDSAFQQKLGNG
YIETAFRAARAA_DPTAXLCINDYNVEGINAKSN
IVQWDVVNEAFADGSSGAR
RDSNLQNVIGQD
YLDYAFRYAREA_DPDALLFYNDYNIEDLG_PKSN
IYAWDWNEIF
NEDGSLRDSVFYKVIGDD
YVRIAFETARAA_DPNAKLYINDYNLDSASYPKLA
AFAWDVVNEAF
NITGRL
EVAAASRTDPNAKLYINDYNLDSARYPKTQ
VMKNHITTVMKQYKGK
LYAWDWNEIF
EEDGTLRDSVFSRVLGED
FVRIAFETAREA_DPEAKLYINDYNLDSATSAKLQ
RLESMIKNTFAALKSQYPNLD_VYSYDVCNELFLNNGGGMRGAD
NSNWVKIYGDDSFVINAFKYARQY
APAGCKLYLNDYNEYI P_AKTN
RLEFYVKSVNGHFYSGKTGST_LVYWDVCNETL
HAp
NSGWEAVYGSNKTNAVYVKKAFNYAYQVLEQYKLTNSVKLFYNDYNTYMEVN
RLESYIHGVLDFVQTNYPGI_IYAWDVVNE_IVDEGA_FRK
SIWTETVGED
FFIKAFEFARKY
AAPEVSLFYNDYETAQP_WKRD
RLESYIKQVIEFCQKNYPGV_VYCWDVVNEAILDDGS_WREI
NNNWYTINKEK
YVEKAFYYARKY
AKKDVALFYNDYNVFL P_AKRE
RLKKHIQTVVGRYKGK
VYAWDWNEAIDE
NQPDGYRRSDWYNILGPE
YIEKAFIWAHEAD_PKAKLFYNDYSTEDP_Y_KRE
RLKQYIYDVVGRYKGK
VYAWDWNEAIDE
NQPDSYRRSTWYEICRSGND_WIEVAFTRARAAD_PSAELCYNDYNVENWTWAKTQ
VMKNHITTVMTHYKGK
IVEWDVANECMDDSGNG
LRSSIWRGPE
YIEKAFIWAHEAD_PNAICLFYNDYNTEIS_K_KRD
RLKTHITTVLDHFKTKYGAQNPIIQWDWNEVLD
DNGSLRNSKWLQI IGPD
YIEKAFEYAHEAD_PSMKLFINDYNIENNGV_KTQ
RLKTHITTVLDHFKTKYGSQNPIIGWDVVNEVLD
DNGNLRNSKWLQIIGPD
YIEKAFEYAHEAD_PSNKLFINDYNIENNGV_KTQ
RMENHIKTVVERYK
DDVTSWDVVNEVID
DGGGLRESEWYQITGTD
YIKVAFETARKYG_GEEAELYINDYNTEVP_SJCRD
RLENYIRAVVLRYK
NDPGGMRNS PWYQITGTE
DDIKSWDVVNEVIEP
YIEVAFRATREAG_GSDIKLYINDYNTDDP_V_KRD
RLREHIKTLCERYK
DVVYAWDVVNEAVED
KTEKLLRESNWRKIIGDD
YIKIAFEIAREYAG_DAXLFYNDYNNEMP_Y_KLE
-QWIRDYCARYPDT
NIDVVNEAVPGHQPAGYAQRAFGNNWIQRV
FQLARQYC-- PNSILILNDYNNIR- -WQHN
VLKNHIDNVIGRYKDD
LAYFDIVNE PL
YIETALRYAHE_VAPKIvIKLCINDYNIETVN_AKSQ
NENGTYIKSNVWYNVGLES
DFARHIDTVAAJ-{FAGO
VKSWDWNEALFDSADDPDGRGSANGYRQSVFYRQFGGP
EYDEAFRRAPRA_DPTAELYYNDFNTEE_NGAKTT
103
AMVNHVTKVADHFEGK
4IDHINGVMàHYKGK
VJxIKNHITTVMQHYKGK
123 127 (a/b)
32
102
IADSEFNLVVAENANKWD
ATEPSQN_SFS
FGAGDRVASYAADTGI<ELYGHTLVWHS_QLPDWA
KNLN_G_SAFES
IAGREFNMVTAENEMKID
ATEPQRG_QFN
FSSADRVYNWAVQNGIKQVRGHTLAWIIS_QQPGWM
QSLS_G_RPLQ_
ILQREFSMVVCENEMKFD
ALQPRQN_VFD
FSKGDQLLAFAERNGMQMRGHTLIWHN_QNPSWL
TNGNWNRDSLLA
VIKADFGALTPENSMKWD
ATEPSRG_QFS
FSGSDYLVNFAQSNNKLIRGHTLVWFIS_QLPSWV
QAIT_DKNTLIE
IIQADFGQVTPENRMKWD
ATE PSQG_NFN
FAGADYLVNWAQQNGKLIYGHTLVWWS_QLPPWV
VSIT_DI<_____
IIKANFGQLSPENSMKWD
ATEPSQG_QFS
FAGSDYFVEFAETNGKLIRGHTLVWHS_QLPSWV
SSIT_D1CTTLTD
FLKI{HYNSITPENELKPES ILDQGACQQKGNNVNTQ
ISLSRAAQTLKFCEQNGIALRGHTFVWYS_QTPDWFFRENFSQNG_AYVS
KDIMNQ
DEAKRLGYYIPSNYIKERWPK
IDFRTVDEAVKICYENGLKMRGHTLVWHS_QTPTWLFRENYSGNG_RFVN
TATMDA
LLAEQFNSFTCENDMKPMYYLDREANK_KDPEKYNLS PALTFENAIPYLEFAKDNKIANRGHTLVWHN_QTPKWFFCERYNENF_PMAD
RETILA
MKQQYLLDYEATh_ASK_NGMPVCKFDSCIPALQFCKENGIKMRGI-IVLVWHN_QTPEWFFHKDYDVSK_PLVD
AATNAR
VIKRHFNSITPENEMKPESL
QPYEG_GFS
FS IADEYVDFCKKDNISLRGHTLVWHQQTPSWFFTN_PETGEKL
TNSEKDKEILLD
MVLK}-IFNSITAENEMKPESLL_AGQTSTGLSYR
FSTADAFVDFASTNKIGIRGHTLVWHN_QTPDWFFKD_S_NGQRL
S
KDALLA
LTAKHFNNLVAENANKPESL
QPTEG_NFT
FDNADRIVDYAIAHNNKMRGHTLLWHN_QVPDWFFQD_P
SDPTK_PASRDLLLQ
LTAKHFNNLVAENANKPESL
QPTEG_NFT
FDNADKIVDYAIAHNMKMRGHTLLWHN_QVPDWFFQD_P
SDPSK_SASRDLLLQ
ILKHHYNSLVAENAMKPESL
QPREG_EWN
WEGADKIVEFARKHNMELRFHTLVWHS_QVPEWFFID_EDGNRMVDETDPDKREANKQLLLE
LYKKHVNMLVAENAMKPASL
QPTEG_NFQ
WADADRIVQF(ENGMELRFRTLVWHN_QTPTGFSLQKEGKPMVEETDPQKREENRKLLLQ
ILLKHFNSLTPENNKFENI
HPEEQ_RYN
FEEVARIKEFAIKNDMKLRGHTFVWHN_QTPGWVFLDKNGE
EASKELVIE
---RYWNQITPBNESKWGSV
EGTRNVYNWAPLDRIYAYARQNNIPVKAHTFVW_GAQSPSWL
ILESQFDAITPENEMKWE
VVEPTEGNFD
FTGTDKIVAEAKKTGSLLRGHNICWDS_QLTPAYV
TSIT_DPTKLKK
IVRAEFNQITAENIMKM
SYMYSGS_NFS
FTNSDRLVSWAAQNGQTVHGHALVWH PSYQLPNWA
SD_S_N_ANFRQ
s
t
m
n
o
p
q
r
k
1
j
I
h
g
f
e
d
c
b
a
t
m
n
o
p
q
r
s
k
1
j
h
1
g
f
e
d
c
b
a
-
-
-
-
-
-
-
-
-
-
-
-
-________
QVDDYYT
QKARYKE
SVSACLG_NDLCPGVSIWQFADPTSW
IVQAYLEVVPPGRRGGITVWGIADPDSWjYTHQNLPDWPLLFFNDNLQPKPAYQGVVEALSG
-
-
-
--
-
-
-
--
-
-
314
VVQACMQVT_RCQGVTVWGITDKYSWVPDVFPGEGAALVWDASThJ(KPAYAAVMEAFGA
VTNVCLAVSRCLGITVWGVRDSDSW
RSEQTPLLFNNDGSKKAAYTAVLDALNG
LMKICLA_NPNCNTFVMWGFTDKYTWIPGTFPGYGNPLIYDSNYNPKPAYNAIKEALMG
VVEACLQQPKCIGITVWGVADPDSW
TSTDYVD
VVNACLQQPKCVGITVWGVADPDSW
QADLYEKIF
LANQNSAQIPAVTIWGTQDTVSWRSS
K
QNP
LLFSAGYQPKPAY
LNNYAYRLF
KNI_NAAKKNGGNISCITWWGPSDAETWIRN
EKP
LIWSNIGVAKPAY
LATRYQEFF
QTYL_DAKKSGKANITSVTFWNLLDENSWLSG_FRRETSYP
LVFKGKCEAKEAYYAVLKA-QADRYYEMM
YLWDKNCNPKPCFYSFLQAKLLLKEDTDNGGPCNITCVTVFGICDDYPLYKN_FK2CM
QAQKLKAIFDVLKKYRNVV
TSVTFWGLKDDYSWLRG
DMPLLSDKDYQPKFAFWSLID--QSQKYKEIFTMLKKYKNVV
KSVTFWGLKDDYSWLRS_FYGKN
DWPLL_FFEDYSAKPAYWAVIEQARLYKQLFDLFKAEKQYI
TAVVFWGVSDDVTWLS
KPNAPLL_FDSKLQAKPAYWAIAD--QARLYEQLFDLFKAEKQYI
TAVVFWGVSDDVTWLS
KPNAPLL_FDSKLQAKPAFWAVVD--QADRYDQLFELYEELAADI
SSVTFWGIADNHTWLDGRAREYNNGVGIDAPFV_FDHNYRVKPAYWRIIDQAKRYQELFDALKENKDIV
SAVVFWGISDKYSWLNG_FPVKRTN
APLL_FDRNFMPKPAFWAIVDQAKVYEDVFAVFREYKDVI
TSVTLWGISDRHTWKDN$PVKQRKDWPLL_FDVNGKPKEALYRI
250
QAADYKK
PASTYAN
QANNYKE
SSTDYVE
182
249
SLY_DLVKDFKARGVPLDCVGF
QSHLIVG_QVPGDFRQNLQRFADL_GVDVRITELDIRMRTPSDATK
LAT
-AMY_NMVRDFI{QRGVPIDCVGF
QSHFNSG_SP_YNSNFRTTLQNFAAL_GVDVAITELDIQ
GA
-AVFNMIKSMI<ERGVPIDGVGF
QCHFINGMSPEYLASIDQNIKRYAEI_GVIVSFTEIDIRI PQSEN
PATAFQV
-GMVSHVKKWIEAGIPIDGIG
SQTH_LSGGAG
ISGALNALAGA_GTKEIAVTELDI
AGA
-AIVNRVKQWCAAGVPIIGIG
NQTARAA
-GMV_SHVKKWIAAGVPIDGIG
SQTH_LGAG
AGAAASGALNALASA_GTEEVAVTELDI
AGA
-DIY_NMANKLI{QL_GYIDGIGM
QSH_LATNYP
DANTYETALKKFLST_GLEVQITELDITC
TNS
AE
-DV
IKLV
NYINQGKKVCAGVGNQSH_LGTGFP
SVDYYTNALNSFLRA_GFEVQITELDITN
KGD
YD
-FILEKVLGPLIDK_KLIDGMGM
QSJ-1_LLMDHP
DISEYRTALEMYGST_GLQIHITELDMH
NADPSEESMHA
-AIY_NLAQKLKEK_GLIDGLGL
QPT_VGLNYPELDSDDIDSFKTTLETFAKL_GLQIHITELNFEI
KGDESNRTP_ENLKK
-FIY_KLIKNLKAKGVPVHGVGL
QCI-I_ISLDWPD
VSEIEETVKLFSRI PGLEIHFTEIDISIAKNMTDDDAYN
RYLLVQ
-FIY_NMVKNLKSKGIPIHGIGM
QCHINVNWPS
VSEIENSI_LFSSIP_GIEIHITELDMSL
YNYGSSENYSTPPQDLLQK
-AMY_DLVKKLKSEGVPISGIGM
QMH_INI_NSN
IDNIKASIEKLASL_GVEIQVTELDMNN_flG
NVSNEALLK
-AMY_DLVKI<LKSEGVPIDGIGM
QMH_INI_NSN
IDNIKASIEKLASL_GVEIQVTELDMNM_NG
NISNEALLIK
-DLY_NLVKDLLEQGVPIDGVGH
QSH.JQIGWPS
IEDTRASFEKFTSL_GLDNQVTELDMSL_YGWPPTGAYT
SYDDIPAELLQA
-ILYELVKNLLEKGVPIDGVGH
QTH_IDIYNPP
VERIIESIKKFAGL_GLDNIITELDMSIYSWNDRSDYG
DSIPDYILTL
-IKTY_KVLKELLERGTPIDGIGI
QAH_WNIWDIKNLVSNLKKAIEVYASL_GLEIHITELDISV_FEFEDKRT_D
LFEPTP_EMLEL
-EFIALAKAQGNYIDAVGL
QAHELIKGMTAAQVXTAIDNIWNQVGKPIYISEYDIGDTNQVQLQNFQAHFPVF-AMA_IKVAAGLLAKGAPLHCIGMFKNAKRRSSGLLIRTASSGLESHFIGGSTPKDIPAAMNLFSD_QGLEVPMTELDVRIP
VNGNDM_PANATVAKE
-ALV_NLVQRLLNNGVPIDGVG
FQMHVMND_YPS
IANIRQANQKIVALSPTLKIKITELDVRLNNPYDGNSSNDYTNRNDC_AVSCAGLDR
+
233 (n)
01
53
Table 4.1. Kinetic parameters for hydrolysis of cellobiosides and glucosides by
wild-type Cex
1
Enzyme
Substrate
)
1
kcat (miir
Km (mM)
kcat/Km
)
1
1 .mM
(min
Wild-type
DNPG
DNPC
PNPC
PNPG
860
419
677
1.4
1.87
0.06
0.53
8.33
460
6983
1278
0.17
Parameters were determined at 37°C in 50 mM phosphate buffer, pH 7.0.
1
54
glucoside (2,4-DNPG) were within the same order of magnitude at 677 min
, 419 miiv
1
1
and 860 mm
4 respectively. The ‘<cat value for p-nitrophenyl 13-D-glucoside (PNPG), 1.4
, was much lower. The Km values were lowest for the cellobiosides, 0.06 mM for
4
mm
2,4-DNPC and 0.53 mM for PNPC, whereas those for the glucosides were much higher:
1.87 mM for DNPG and 8.33 mM for PNPG. The kcat/Km values were 3 to 4 orders of
magnitude greater for the cellobiosides than they were for the glucosides.
The hydrolysis of 13-1,4-glycans by a retaining enzyme involves a two-step
process: the formation (glycosylation) and hydrolysis (deglycosylation) of an x-D
glycopyranosyl-enzyme intermediate (see Introduction, Figure 1.4). The rate of the
deglycosylation step in the reaction will be identical for substrates containing the same
sugar residue(s) since the aglycon unit is no longer present. It was previously
determined (Tull and Withers, 1994) that the rate-limiting step is glycosylation for the
glucosides and deglycosylation for the cellobiosides. The results of this study are
consistent with the findings of Tull and Withers (1994). The cellobiosides, with aglycon
units of quite different leaving group ability, were hydrolysed at essentially the same
rates suggesting that deglycosylation is rate-limiting. In contrast, the leaving group
ability of the aglycon unit on the glucosides had a dramatic impact on the rate of
hydrolysis, suggesting that glycosylation is rate-limiting for the glucosides.
4.2 The putative catalytic nucleophile in Cex: Glu 233
The nucleophile in a retaining 3-1,4-glycanase can be identified using an
activated 2-deoxy-2-fluoro glycoside mechanism-based inactivator which functions by
forming a highly stabilized intermediate in which the sugar is covalently bonded to the
catalytic nucleophile. Using the inactivator 2-deoxy-2-fluoro J3-D glucoside (2F-DNPG),
G1u358 was identified as the catalytic nucleophile in an Agrobcicterium 13-glucosidase
55
(Abg) (Withers and Street, 1988; Withers et al, 1990) a retaining f3-glucanase from
another family of enzymes. When the active site nucleophile was targeted by sitedirected in vitro mutagenesis, Abg retained measurable activity upon mutation of
G1u358 to Asp, but virtually no activity upon mutation to Asn or Gin. Glu358 is part of
a conserved ITE motif found in many retaining glycosidases. Similarly, in Cex, Glu 233
is part of a highly conserved ITE motif within the F-family of f3-1,4-glycanases (Figure
4.1).
4.2.1 Generation of mutants at position 233
Glu 233 was targeted by site-directed mutagenesis to determine whether it had
properties consistent with those of a catalytic nucleophile. Site-directed in vitro
mutagenesis was performed in order to substitute Glu 233 with Asp and Gin using the
oligonucleotides as listed in Materials and Methods (Table 2.3). The phagemid pTZ18Rcex was constructed as outlined in Figure 4.2. Following mutagenesis, pTZ18R-cex was
transformed to E. coliJMlOl. Initial screening for Cex mutants was on LB agar
supplemented with 100 ig/mL ampicillin and 100 j.iM MUC. Approximately 50% of
the colonies were non-fluorescing, indicating loss of exoglucanase activity. Mutation
anywhere within the codon for Glu 233 resulted in the destruction of a Sca I restriction
site. Restriction analysis of plasmid DNA isolated by small-scale alkaline lysis revealed
approximately 80% of the non-fluorescing colonies had lost the ScaT restriction site.
DNA sequencing of an 800 base pair Barn H1-Pstl fragment of several of these clones
revealed they contained only the mutation of interest. The 800 bp sequenced mutant
cassettes (the Barn H1-Pstl fragments) were subcloned from pTZI8R-cex into pUC121.lcex(PTIS), replacing the equivalent wild-type fragment of DNA as shown in Figure
4.3. pUC12-1.lcex(PTIS) was transformed to E. coli JM1O1 for expression of the mutant
proteins.
56
Figure 4.2. Construction of pTZ18R-cex. Plasmids pTZI8R (Mead et al., 1986) and
pUC12-1.lcex(PTIS) (ONei11 et a!., 1986a) were digested with BamHI and HindIII.
The 1.8 kb BamHI-Hin dill fragment containing the cex gene was isolated and
ligated with the 2.9 kb BamHI-HindIII fragment of pTZ18R to give plasmid pTZ18Rcex (4.7 kb).
57
tli.fidll1 ,Pstl
BamHl
BamHl
Iaczpo
IacZpo
on
Fl on
cex
pTZ18R
U
Ap
Pstl
V
pUC12-1.lCex
(PTIS)
R
4.5kb
R
IacZ
Hindill
Pstl
Sail
Digest each with BamHI/HindIlI
Isolate 1 .8 kb fragment
by GeneCleanTM
ligate DNA
BamHl
IacZpo
oil
Pstl
cex
pTZ18R-Cex
4.7kb
lion
IacZ
Sail
Pstl
Hindlil
R
Ap
58
Figure 4.3. Generation of pUC12-I.Icex(PTIS) encoding Cex mutants. In vitro
mutagenesis was performed as described Materials and Methods (section 2.4.2). For
each mutant, an 800 bp mutant cassette in pTZI8R-cex was sequenced to confirm
that only the desired mutation was present. The 800 bp fragment was isolated
following digestion with Barn HI and Pstl. pUC12-I.lcex(PTIS) was also digested
with Barn Hi and Psti and the 3.7 kb fragment was isolated. The 800 bp mutant
cassette was ligated with the 3.7 kb fragment to replace the equivalent fragment in
pUC12-i.lcex(PTIS). The positions of the mutation(s) generated in this study are
indicated.
in vitro mutagenesis
59
BamHl
BamHl
IacZpo
E127
IacZpo
on
oil
E233’
cex
cex
R
pTZ18R-Cex
4.7 kb
pUC12-tiCex
Pstl
Ap
Ap
4.5 kb
ii on
IacZ
IacZ
Sail
Pstl
Hindlll
Hindlil
Sail
Digest each construct
with BamHI/Pstl
Isolate 0.8 kb sequenced
mutant cassette by GeneCleanTM
Isolate 3.7 kb fragment
by GeneCleanTM
*
ligate DNA
BaniHi
Dl24
El27
!acZpo
on
E233
Pstl
cex
pUC12-1.lCex
(PTIS)
4.5kb
R
Ap
IacZ
Hindu
Sail
60
The two mutants obtained, encoding Cex E233D and E233Q, were expressed at
approximately the same levels as the wild type gene. The mutant proteins were
purified by affinity chromatography on cellulose. When analysed by SDS-PAGE
(Figure 4.4), the mutant polypeptides had been purified to electrophoretic homogeneity
and had the same apparent molecular mass as wild-type Cex. No differences in
behaviour from the wild-type were observed during the purification procedure.
4.2.2 Determination of kinetic parameters for Glu 233 mutants
The kinetic parameters for hydrolysis of PNPC by mutants at position 233 are
presented in Table 4.2. In comparison with the wild-type enzyme, E233D had only
1/4000 the activity. The Km value for this substrate rose only slightly upon mutation to
Asp, from 0.53 mM to 1.2 mM suggesting that mutation of this residue likely effects both
the glycosylation and deglycosylation steps of the reaction. If the mutation
preferentially effected one step of the reaction, the concentration of the glycosyl-enzyme
intermediate would change, which would likely be reflected in the Km value. No
activity could be detected upon mutation of Glu to Gin.
The kinetic parameters for the Glu 233 mutants in Cex were consistent with those
found for mutants of the catalytic nucleophile in Abg (Withers and Street, 1988; Withers
et al, 1990). Simultaneously, it was shown that Cex, like Abg, could be inactivated with
2F-DNPG leading to the accumulation of the covalent enzyme intermediate in which
the sugar was esterified to Glu 233 (Tull et al., 1991). Labeling of this same residue was
also been observed recently with 2’ ‘,4’ ‘-dinitrophenyl 2-deoxy-2-fluoro-13-cellobioside
(2F-DNPC) (Tull and Withers, 1994) providing further evidence that Glu 233 is the
catalytic nucleophile in Cex.
61
kDa
1
2
3
4
5
6
78
130
97.4
68.0
57.5—
53 -,
45—
41 /
36 /
29
E233D
E233Q
Figure 4.4. Purification of Cex E233D and E233Q. Cex P233D and E233Q were
purified from 2-L cultures of E. coli containing pUC12-1.lcex E233D (PTIS) and
pUC12-1.Icex E233Q (PTIS) as described in Materials and Methods (sections 2.7.2
and 2.7.3). The purification was followed by SDS-PAGE: Lane 1, molecular weight
standards with sizes as indicated; lane 2, 3 tL (9.4 mg/mL) crude extract; lane 3,
same volume extract following streptomycin sulfate precipitation; lane 4, same
volume extract following 40,000 x g centrifugation; lanes 5 and 6: following cellulose
affinity chromatography, E233D 2 ig and 10 .tg; lanes 7 and 8: E233Q 2 g and 10
pg respectively.
62
Table 4.2. Kinetic parameters for hydrolysis of p-nitrophenyl-I3-D cellobioside
(PNPC) by Cex and Cex E233 mutants.
1
Enzyme
2
V
max
1 .mg
(.tmo1.min
)
1
Cex wild-type
14.4
Km
(mM)
kcat
)
4
(mm
677
3
o-
E233D
3.6 x
E233Q
no activity detected
0.53
.168
1.2
—
kcat/Km
4 .mM
min
)
4
1278
1.97
—
Parameters were determined at 37°C in 50 mM phosphate buffer, pH 7.0. 2
1
j.imol
PNP released per minute per rng of purified enzyme.
63
4.3 Identification of the acid/base catalyst
In addition to Glu 233, there are six other conserved acidic residues within the Ffamily of 3-glycanases which could be candidates for the acid/base catalyst in Cex
(Figure 4.1). Of these, Glu 127 in Cex is part of a highly conserved region, WDVVNEA.
Within this region, the short consensus sequence NEX (where X is a small hydrophobic
residue) occurs in a large number of retaining glycanases in other sequence-related
families. This type of motif has been suggested previously (Baird et al., 1990) to play an
important role based upon alignments of sequences of a number of glycanases. It
appeared therefore that Glu 127 might be a suitable candidate for the acid/base catalyst
in Cex.
4.3.1 Generation of mutants at position 127
In vitro mutagenesis was performed in order to substitute Glu 127 with Asp, Ala
and Gly using the oligonucleotides listed in Materials and Methods (Table 2.3).
Approximately 50% of the colonies were non-fluorescent, indicating loss of
exoglucanase activity. DNA sequencing of the 800 bp Barn H1-Pst I fragment of several
of these clones revealed that about 80% of them contained only the mutation(s) of
interest. Subcloning was performed as described for the Glu 233mutants. The three
mutants obtained, E127A, E127G and E127D were expressed at approximately the same
levels as the wild type. The purified proteins obtained by affinity chromatography on
cellulose had the same apparent molecular mass as wild-type Cex and were
electrophoretically homogeneous as judged by SDS-PAGE (Figure 4.5.) No differences
in behaviour were observed during the purification procedure.
Purified wild-type Cex and Cex E127A and E127G were subjected to mass
spectrometry to verify that the actual molecular mass agreed with that predicted from
64
kDa
1
2
3
4
5
6
7
8
130
97.4
68.0
57.5
—I-.
41
36
29
E127A
E127D
E127G
Figure 4.5. Purification of Cex E127A, E127G and P127D. Cex E127A was purified
from a 20-L culture of E. coli containing pUC12-1.lcex E127A (PTIS). Cex E127D and
E127G were purified from 2-L cultures of E. coil containing pUCI2-1.lcex E127D
(PTIS) and pUCI2-1.lcex E127G (PTIS) respectively as described in Materials and
Methods (sections 2.7.2 and 2.7.3). Purified protein was analyzed by SDS-PAGE:
Lane 1, molecular weight standards with sizes as indicated; lane 2, 1 iL crude
extract (0.lmLs of original culture volume); lanes 3 and 4, following cellulose
affinity chromatography, 2 ig and 20 tg (by A280); lanes 5 and 6: E127D following
cellulose affinity chromatography, 2 tg and 10 ‘g; lanes 7 and 8, E127G following
cellulose affinity chromatography, 2 ig and 20 pg respectively.
65
the amino acid sequence. As shown in Table 4.3, the difference in mass between the
mutants and the wild type were consistent with that predicted from the amino acid
composition, within experimental error. When the mutants were reacted with 2,4DNPC prior to mass spectrometry, the masses increased by an amount equivalent to the
mass of cellobiose (less an OH) indicating the presence of the cellobiosyl-enzyme
intermediate as expected based on the mechanism proposed for the wild-type enzyme
(see Introduction, Figure 1.4)
4.3.2 Kinetic characterization of mutants of Glu 127
The catalytic mechanism of retaining glycosidases such as Cex involves a two
step process; formation and hydrolysis of a glycosyl-enzyme intermediate (see
Introduction, Figure 1.4). It is likely that a single amino acid residue (the acid/base
catalyst) is responsible for both proton transfer steps. The first step is acid catalysis, in
which a proton is transferred from the acid/base catalyst to the glycosidic oxygen,
facilitating bond cleavage through stabilization of the leaving group. In the second step,
the same residue functions as a general base catalyst, removing a proton from water in a
concerted process as the water attacks the anomeric center of the glycosyl-enzyme
intermediate. It is clear, therefore, that mutation of the acid/base catalyst will affect the
rates of both steps. The extent to which each step is affected however, is not necessarily
equivalent. The effect of modifying the acid/base catalyst on the rate of the second
step, deglycosylation, will necessarily be identical for all substrates containing the same
sugar residues. However, the effects on the first step (glycosylation) will depend upon
the leaving group ability of the aglycon (refer to Figtires 1.4 and 1.6). Those of high pKa,
therefore poor leaving group ability, should be affected most following mutation of the
acid/base catalyst, while those of low pKa, which need little or no protonic assistance
for departure, should be affected very little.
316.1 ± 13.2
320.7 ± 11.4
47 370.2 ± 6.6
47 363.7 ± 6.0
47 064.43
47 050.40
47 054.8 ± 6.6
47 043.1 ± 5.4
E127G
The Mr of 2,4 DNPC is 508.4. The Mr of cellobiose (less OH) is 325.3
2 Substrate was added to the enzyme sample prior to mass spec analysis. n.d. means not determined
1 Predicted Mr is based on the aa sequence and is calculated as the average isotopic (MH+) mass
n.d.
E127A
n.d.
47 122.47
47 119.5 ± 3.8
Difference
(b-a)
Wild-type Cex
Reacted with
2 (b)
’
1
2,4-DNPC
Predicted Mr 1
Determined Mr
(a)
Mass spectrometry of purified Cex and E127 mutants
in the absence and presence of 2,4-DNPC
Protein
Table 4.3.
67
For each mutant at position 127, kinetic parameters were determined for various
cellobiosides and glucosides with differing requirements for acid catalysis (Table 4.4).
Three different cellobioside substrates were studied: PNPC (pKa of phenolic leaving
group
=
7.18), 2,4-DNPC (pKa of phenolic leaving group
=
3.96) and 4-BrPC (pKa of
phenolic leaving group
=
9.34). Kinetics were also determined for 2,4-DNPG (pKa of
phenolic leaving group
=
3.96). The effect of mutation of Glu 127 to Ala, Glu or Asp on
kcat values for PNPC, 2,4-DNPC and 2,4-DNPG was to decrease them approximately
200 to 400-fold. There were also dramatic decreases in Km for these substrates, ranging
from 20-fold for PNPC to 600-fold for DNPG. With the mutants, the Km values for 2,4DNPC were less than 1 p.M, a drop of about 200-fold relative to the wild-type enzyme.
The kcat/Km values however, for PNPC and 2,4-DNPC remained relatively unchanged
relative to the wild-type.
Based on the data in Table 4.4 , it seems likely that deglycosylation is the ratedetermining step for hydrolysis of PNPC and 2,4-DNPC for the Glu 127 mutants, as it is
for the wild-type enzyme. First, essentially the same kcat values were seen for the two
substrates with quite different leaving-group ability. Second, the Km values dropped
dramatically. The drop in Km suggests an accumulation of the glycosyl-enzyme
intermediate as would be expected if the deglycosylation step were rate-limiting.
Similarly, the rate-determining step for DNPG appears to be deglycosylation. For this
substrate, the rate-determining step seems to have switched from glycosylation in the
case of the wild-type enzyme, to deglycosylation with the mutant. It is apparent,
therefore, that since deglycosylation is rate-limiting in the presence of a good leaving
group (i.e. pNP or DNP), the decrease in kcat observed following substution of Glu 127
is due to slowing of the deglycosylation (base catalysis) step.
68
Table 4.4.
Kinetic parameters for hydrolysis of various substrates
by Cex and E127 mutants’
Enzyme
Substrate
)
4
kcat (mm
Km (mM)
kcat/Km
mM
(min
)
1
Native Cex
DNPG2
2
DNPC
PNPC2
4-BrPC
860
419
677
255
1.87
0.06
0.53
2.0
460
6983
1278
128
E127A
DNPG
DNPC
PNPC
4-BrPC
2.7
2.4
2.3
0.04
0.0032
0.0003
0.025
1.9
843
7742
92
0.02
E127G
DNPG
DNPC
PNPC
2.2
2.1
2.0
0.0035
0.0004
0.029
628
5526
69
E127D
DNPC
PNPC
1.6
3.3
0.0017
1.74
941
2
1
P
arameters were determined at 37°C in 50 mM phosphate buffer, pH 7.0.
Data from Table 4.1.
2
69
Further evidence of the slowing of the deglycosylation step and the resulting
accumulation of the glycosyl-enzyme intermediate can be seen from examination of the
mass spectrometry data (Table 4.3). As mentioned previously, reaction of the 127A or
127G mutants with 2,4-DNPC prior to mass spectrometry resulted in an increase in
molecular mass corresponding to an accumulation of the glycosyl-enzyme intermediate.
Such an increase in mass can not be observed with the wild-type enzyme presumably
because the enzyme-glycosyl intermediate is rapidly hydrolysed (S. Withers, personal
communication). Again, these results suggest that with substrates with a good leaving
group (which presumably do not require acid-catalytic assistance for hydrolysis), the
reductions in activity observed are likely due to a slowing of the deglycosylation step,
where base catalysis occurs, and not due to a slowing of the rate of glycosylation.
One measure of the effect of this mutation on the glycosylation step or acid
catalysis step for these substrates (i.e. the formation of the glycosyl-enzyme
intermediate) can be obtained by examination of kcat/Km values, as mentioned in the
introduction. Essentially no reduction in kcat/Km was observed with 2,4-DNPC, while
a 10-20 fold reduction was observed for PNPC. The effect of the mutation upon the
glycosylation step can also be estimated by studying the kinetics of inactivation with 2FDNPC. As discussed earlier, the inactivation of Cex with 2F-DNPC involves simply the
formation of the glycosyl-enzyme intermediate. The results of inactivation of E127A
with 2F-DNPC are shown in Figure 4.6. It is apparent that this mutant was inactivated
very quickly. The rate of inactivation of E127A by 2F-DNPC (k/K
was at least as fast as that of the wild type enzyme (k/Kt
=
=
1.6 1
min
m
M’)
)
1
0.61 minmM
determined previously (Tull and Withers, 1994). Again, these results suggest that
mutation of Glu 127 to Ala did not slow down the rate of the glycosylation (acid
catalysis) step in the reaction when a substrate with a good leaving group was
hydrolysed.
70
Figure 4.6. Inactivation of Cex E127A by 2F.-DNPC. Panel A: Residual DNPCase
activity vs. time for E127A incubated with 2F-DNPC at the following concentrations:
0.031 mM (V), 0.045 mM (), 0.090 mM (EJ) and 0.136 mM (0).
0.015 mM
Residual enzyme activity was monitored as described in section 2.9.2. Panel B:
double reciprocal plot of k
b (derived from panel A) vs. 2F-DNPC concentration.
0
Panel C: The inactivation rate constant (k) and the inactivation binding constant
) were derived from the data in panel B.
1
(K
(a),
71
A
0.0
1.0
0
> -2.0
>
-3.0
-4.0
10
0
30
20
40
Time (mm)
B
60
50
C
E
40
Co
30
-o
0
20
10
0
0
10
20
30
40
1/[2F-DNPC]
50
60
70
(1/mM)
C Inactivation rate and binding constants for E127A with 2F-DNPC
1 (min
k
)
1
1 (mM)
K
kj/Kj (minl.mm1)
0.44 ± 0.18
0.28 ± 0.16
1.6
±
0.34
72
E127A was also assayed with 4-BrPC (pKa of phenolic leaving group
=
9.34), a
substrate with a greater requirement for acid-catalytic assistance. As shown in Table
4.4, the kcat value was lowered 6400-fold in comparison with the wild-type enzyme,
whereas the Km value remained relatively unchanged. The kcat/Km value was also
reduced 6400-fold relative to the wild-type enzyme with this substrate. The ratedetermining step for this substrate with wild type enzyme is thought to be formation of
the glycosyl-enzyme. The fact that the Km value for the mutant was similar to that of the
wild type would suggest that the glycosylation step remained rate-limiting with the
mutant. The 6400-fold rate reduction therefore represents the effect upon the
glycosylation step (acid catalysis) for this substrate, a much greater rate-reduction than
that seen on the glycosylation step for 2,4-DNPC or PNPC. Hydrolysis of 4-BrPC would
be expected to be more dependent on the acid catalyst than hydrolysis of PNPC, 2,4DNPC and 2,4-DNPG. The marked reduction in kcat for hydrolysis of 4-BrPC and the
much smaller reductions for the hydrolysis of PNPC, 2,4-DNPC and 2,4-DNPG by
E127A are consistent with Glu 127 being the acid/base catalyst in Cex.
4.3.3 Effects of sodium azide on reaction rates
With mutants of Glu 127, the presence of a good leaving group on substrates
such as 2,4-DNPC or PNPC reduced the need for acid catalytic assistance in the
formation of the glycosyl-enzyme intermediate. In effect, the presence of a good leaving
group on substrates such as 2,4-DNPC or PNPC compensated for the missing acid
catalyst function of the mutants. The drop in activity seen with these substrates reflects
primarily the effect of the mutation on the deglycosylation step, which would normally
be facilitated by base catalysis. An attempt was made to compensate for the missing
base catalyst function. With the wild-type enzyme, hydrolysis of the glycosyl-enzyme
intermediate involves the transfer of a proton from water to the acid/base catalyst. The
73
resultant hydroxyl (-OH) species then acts as a nucleophile to displace the catalytic
nucleophile Glu 233, resulting in a product showing retention of configuration at the
anomeric carbon. The mutation of Glu 127 to Ala or Gly presumably generates a cavity
in the active site. Anions of high nucleophilicity might bind at this site and react in
place of water without any need for general base catalysis. Various competitive
nucleophiles including acetate, azide, cyanide, formate, imidazole, and thiocyanate
were tested for their ability to enhance the rate of hydrolysis of PNPC by the Glu 127
mutants. Of the nucleophiles tested, only sodium azide was found to effect
substantially the rate of hydrolysis of PNPC. The rate-enhancement was maximal with
about 100 mM azide. As indicated in Figure 4.7, the effect was specific to the alanine
and glycine mutants. No rate-enhancement was observed with either E127D or with the
wild-type enzyme. This was probably because the glutamate or aspartate carboxylate
prohibited the access of the azide nucleophile, either by steric hindrance, or more likely,
through electrostatic repulsion.
The kinetics of the rate enhancement were studied in detail with the E127A
mutant and several different substrates. The kinetic parameters for hydrolysis of PNPC,
2,4-DNPC and 2,4-DNPG were analyzed at various concentrations of sodium azide.
Maximal increases in kcat were about 8-fold for PNPC to 10-fold for DNPG in the
presence of 60 mM azide as shown in Figures 4.8 and 4.9, respectively. A dramatic
increase in kcat of over 200-fold was seen with 2,4-DNPC at up to 2 M azide (Figure
4.10), surpassing the rate of the wild-type enzyme. The most likely explanation for this
is that as the azide concentration increased, the deglycosylation rate continued to
increase until it surpassed the glycosylation rate. The limiting rate observed in each case
presumably reflected the rate of the glycosylation step for each substrate. With 2,4DNPC, glycosylation is facilitated by the good leaving-group ability of 2,4-DNPC, as
discussed earlier. In contrast, azide did not increase the keat for PNPG (not shown).
Q
74
E127A
E127G
0.25
0.4’
0.20E
E
0.15
0)
a
D
.S
0.3
0.2
0.10
>
>
U
C°
0.05
0.00-
0.1-
0
0
i.i.i.i-
50
0.0
I
150 250 350 450
Azide (mM)
i-i.i.i.
50
E127D
Wild—type
0.20-
I
I
150 250 350 450
Azide (mM)
20
0.15-
15
E
0,
0
0.10
10
>‘
>‘
II
>
>
005
n nn.
0
1111’
50
150
250
Azide (mM)
350
450
5,
0’
50
150 250 350 450
Azide (mM)
Figure 4.7. PNPCase activity of Cex and E127 mutants in the presence of various
concentrations of sodium azide. Reactions were carried out at 37°C in 50 mM
phosphate buffer, pH 7.0 with 6 mM (>10 x Km) PNPC for about 15 mm. The
reaction was stopped by the addition of 3
CO and the absorbance at 400nm was
2
Na
read. Activity is expressed in units 1mg enzyme where 1 Unit =1 tmo1 PNP
released/mm.
75
20
0
0.30
600
15
0.20
C
E
400
0
C
10
.4-’
(U
0
0.10
2
E
E
200
5
0
0
10
20
30
40
50
60
0.00
0
Azide concentration (mM)
Figure 4.8. Kinetic parameters for hydrolysis of PNPC by Cex El. 27A in the
presence of various concentrations of sodium azide. Parameters were determined
at 37°C in 50 mM phosphate buffer, pH 7.0.
2
76
30
30
20
o
15
20
C
-
-—
2
o
10
5
0
A
0
-
0
10
-
20
.
30
I
40
.
I
50
.
•
60
0
0
Azide concentration (mM)
Figure 4.9. Kinetic parameters for hydrolysis of 2,4-DNPG by Cex E127A in the
presence of various concentrations of sodium azide. Parameters were determined
at 37°C in 50 mM phosphate buffer, pH 7.0.
77
600
0.06
400
0.04
600
0
‘I
LI
400
E
(U
0
200
0.02
E
E
E
200
E
(U
0
0
0.00
0
500
1000
1500
0
2000
Azide concentration (mM)
Figure 4.10. Kinetics of hydrolysis of 2,4-DNPC by Cex E127A in the presence of
various concentrations of sodium azide. Parameters were determined at 37°C in 50 mM
phosphate buffer, pH 7.0. The dotted line represents the activity on this substrate (kcat)
for the wild-type enzyme.
78
This finding is entirely reasonable since the rate-determining step for this substrate was
shown previously to be glycosylation; thus, increasing the rate of the deglycosylation
step should not affect the steady state rate.
Dramatic increases in Km values with increasing azide concentration were
observed for each substrate, essentially paralleling the effects on kcat (Figures 4.8, 4.9
and 4.10). The net result of this was that values of kcat/Km remained essentially
constant at differing azide concentrations. The increases in Km were likely a
consequence of the decreased extent of accumulation of the glycosyl-enzyme
intermediate as the rate of the deglycosylation step increased.
This finding is
consistent with azide acting primarily on the deglycosylation step, that is, on the base
catalysis step and not on glycosylation.
4.3.4 Effects of sodium azide on products of hydrolysis
If Glu 127 is the acid/base catalyst, mutation to Ala or Gly should generate a
cavity close to the f3-face of the substrate. If azide activates the mutants by the
mechanism proposed above (see Figure 4.11), the reaction product should be the
corresponding 3-glycosy1-azide.
In order to test this hypothesis, the products of hydrolysis in the absence and
presence of azide were analysed by thin layer chromatography (TLC). Analysis of
reaction mixtures containing PNPC revealed that wild-type Cex produced only the
expected nitrophenol (visible under UV light) plus cellobiose, both in the absence and
presence of sodium azide as shown in Figure 4.12. With both E127A and E127G the
reaction products differed in the absence and presence of azide (Figure 4.13). In the
absence of azide, only cellobiose plus nitrophenol was observed in each case. About
50% of the starting material remained due to the reduced rate of hydrolysis by the
79
I
E127A
E127A
OH
OH
HO’°
HO0
OR
HO
Qo
HO
33
E233
-
Glycoson
ROH
E127A
OH
HO0
HO
gIycosy1-enzyn
I
Q0
intemdiate
33
De1Ycoo,,/)
+
N
...E127A
I
E127A
I
I
OH
HO-°
I
I
I
I
I
OH
HO-L..->’
HOào
33
I
33
I
Figure 4.11. Proposed nchanism for hydrolysis of glycosides by Cex E127A
in the presere of azide. Only 1 glucose residue is shown for simplicity.
80
Wild-type
Cex
pNPC
cellobiosyl-azide
cellobiose
s-
+
Figure 4.12. Hydrolysis of PNPC by wild type Cex in the presence and absence of
azide. Reactions ran for 2 h at 37°C in 50 mM phosphate buffer, pH 7.0 with 1
p.g/mL Cex, 6 mM PNPC and 60 mM azide where indicated. Thin layer
chromatography was performed as described in section 2.10.1. Standards lane (5): 2
i.LL each of 6 mM PNPC, cellobiose and cellobiosyl-azide. Reaction lanes (-) without
azide, (+) with azide, 2 jiL each.
81
E127A
El 27G
pNPC
cellobiosyl-a zide
cellobiose
S
-
+
S-
+
Figure 4.13. Hydrolysis of PNPC by Cex E127A and E127G in the presence and
absence of azide. Reactions ran overnight at 37°C in 50 mM phosphate buffer, pH
7.0 with 100 .tg/mL mutant Cex, 6 mM PNPC and 60 mlvi azide where indicated.
Thin layer chromatography was performed as described in section 2.10.1.
Standards lane (S): 2 jiL each of 6 mM PNPC, cellobiose and cellobiosyl-azide.
Reaction lanes: (-) without azide, (+) with azide, 2 tL each.
82
mutants. In the presence
of azide however, a different sugar product was formed. The
new compound co-migrated on TLC with 3-cellobiosyl-azide. The kinetics of
hydrolysis with the alanine mutant indicated that cellobiosyl-azide was in fact the
only
product produced, as shown in Figure 4.14. With reduced concentrations of azide, both
cellobiosyl-azide and cellobiose could be detected as products (not shown).
The stereochemistry of the new product was determined by H
1
-NMR
spectrometry. The analysis revealed an identical spectrum to that obtained from the
chemically synthesized f3-cellobiosyl-azide (Table 4.5), providing confirmation of the
f3-
stereochemistry. With 2,4-DNPG, the new product formed was 3-glucosyl-azide as
judged by TLC (not shown) and 1
H-NMR (Table 4.5).
Table 4.5. H
1
-NMR spectraa for -cellobiosyl-azide and f3-glucosyl-azide:
-cellobiosyl-azide, 1
H-NMR (400 MHz, D
0): 8 4.47 (d, Ji,
2
2
Hz, H-i); 3.89 (m, 3 H); 3.65 (m, 4 H); 3.35 (m, 6 H).
=
7.8
3-glucosyl-azide, 1
H-NMR (400 MHz, D
0): 8 —4.7 (d, Ji,
2
2 = 9.0 Hz,
H-i); 3.89 (dd, 1 H, J
12.4,
6 12.4, J
65 2.2 Hz, H-6); 3.71 (dd, 1 H,
5.6
H-6’);
Hz,
(m,
3.50
2
H,
H-3,
3.40
H,
5);
1
9.2,
9.8
(t,
’,
6
J
5
413
J
45
J
Hz, H-4); 3.24 (t, 1 H, J
21 9.0, J
3 8.9 Hz, H-2).
,
2
aprovided by T. Lindhorst.
Identical results were observed with both the alanine and glycine mutants. The
results are summarized in Table 4.6. The fact that only f3-cellobiosyl-azide or 3glucosyl-azide was detected confirmed azide attacked only from the top face of the
substrate. This is consistent with a location of the acid/base catalyst on this same face
and confirms that azide activates the mutants by the mechanism proposed.
0
0.75
1.5
2.25
3.0
3.75
4.5
5.25
Std
-
—cellobiose
—cello bios y I a z ide
—pNPC
Figure 4.14. Time course of hydrolysis of PNPC by Cex E127A in the presence of azide. Reactions were carried out
at 37°C in 50 mM phosphate buffer, pH 7.0 with 200 .tg.mL
1 mutant Cex, 6 mM PNPC and 60 mlvi azide. Thin
layer chromatography was performed as described in section 2.10.1. Standards lane (Std): 2 jiL each of 6 mM
PNPC, cellobiose and cellobiosyl-azide. Reaction lanes: 2 iL samples were withdrawn at various intervals as
indicated.
Time (h):
Cex El 27A
00
84
Table 4.6.
Products of hydrolysis with wild-type and E127 mutants in the
presence or absence of sodium azide
Enzyme
Substrate
Products
1
(no azide)
(with azide)
Wild-type
PNPC
cellobiose
cellobiose
E127D
PNPC
cellobiose
cellobiose
E127A
PNPC
DNPG
PNPG
cellobiose
glucose
glucose
j3-cellobiosyl-azide
13-glucosyl-azide
glucosyl-azide
E127G
PNPC
DNPG
cellobiose
glucose
f3-cellobiosyl-azide
3-glucosy1-azide
Products were determined by thin layer chromatography. Reaction mixtures were
incubated overnight at 37°C in 50 mM phosphate buffer, pH 7.0 with 100 j.ig/mL
mutant Cex or 1 Lg/mL Cex, 6 mM substrate and 60 mM azide where indicated.
Stereochemistry was determined by 1
H-NMR. Products also include p-nitrophenol
or dinitrophenol.
85
4.4 Asp 123: another conserved acidic residue
As a control, similar experiments involving sodium azide were performed
with mutants of another residue conserved within the F-family, Asp 123. Asp 123 is
also part of the WDVVNEA motif and is in proximity to Glu 127. Site-directed in
vitro mutagenesis was carried out in order to substitue Asp 123 with Ala as
described for mutants at position 233 and 127. The protein was purified by affinity
chromatography on cellulose as described for the previous mutants. The purified
protein is shown in Figure 4.15.
The rate of hydrolysis of PNPC by D123A was found to be affected both by
azide and by thiocyanate as shown in Figure 4.16. The rate-enhancement observed
with thiocyanate was about half that observed with azide. For either anion, the
rate-enhancement was maximal at about 500 mM. The kinetic parameters for
hydrolysis of PNPC and 2,4-DNPC by D123A are presented in Table 4.7. The kcat
value for 2,4-DNPC was similar to that of the wild-type enzyme, whereas that of
PNPC was reduced by about 1500-fold. Initially, the small rate-reduction for 2,4DNPC and the much larger rate reduction for PNPC appeared to be consistent with
a role of Asp 123 as acid/base catalyst. The
values for both substrates,
however, increased 10-fold and 33-fold for 2,4-DNPC and PNPC respectively. The
increase in Km is inconsistent with mutation at Asp 123 slowing down preferentially
the deglycosylation step of the reaction. This is in contrast to what was observed
with mutants at position 127, where Km values for these substrates were drastically
reduced. The kinetic parameters for hydrolysis of PNPC were determined also in
the presence and absence of azide. In the presence of 500 mM azide, the rate of
hydrolysis of PNPC rose about 10-fold, whereas the Km value dropped slightly.
The drop in Km is inconsistent with azide acting to increase the rate of the
86
1
kDa
2
3
200
116
97.4—
66
—
—
45
31
21.5—
D123A
Figure 4.15. Purification of Cex D123A. Cex D123A was purified from a 2-L culture
of E. coli pUC12-1.lcexDl23A (PTIS) as described in Materials and Methods
(sections 2.7.2 and 2.7.3). The purification was analysed by SDS-PAGE: Lane 1,
molecular weight standards with sizes as indicated; lanes 2 and 3, following
cellulose affinity chromatography, 2 ig and 10 jig respectively.
87
8.0
o
7.0
6.0
x
C
0
0
0
0
D
5.0
E
0
0
0
azide
thiocyanate
D
4.0
D
D
3.0
>
0
2.0
0
1.0
I
0.0
0
500
1000
—
I
1500
—
—
I
2000
Anion concentration (mM)
Figure 4.16. PNPCase activity of Cex D123A in the presence of azide or thiocyanate.
Reactions were carried out with 30 .ig/mL enzyme at 37°C in 50 mM phosphate
buffer, pH 7.0 with 5 mM PNPC and anion concentrations, NaN
3 (0) and KSCN (
D) as indicated. Hydrolysis of PNPC was monitored continuously at 400 nm over a
10 mm. period. Activity is expressed as initial rates of hydrolysis (Vo).
88
Table 4.7.
Kinetic parameters for hydrolysis of cellobiosides by D123A
1
Enzyme
Substrate
)
1
kcat (miir
Km (mM)
kcat/Km
1 mM
(min
)
1
Wild-type
DNPC
PNPC
419
677
0.06
0.53
6983
1278
D123A
DNPC
PNPC
PNPC
(O.5Mazide)
325
0.43
0.53
18
613
0.02
5.0
11
0.47
Parameters were determined at 37°C in 50 mM phosphate buffer, pH 7.0.
1
89
deglycosylation step. Azide did not enhance the rate of hydrolysis of 2,4DNPC (not
shown).
TLC analysis of the products of hydrolysis of PNPC in the presence and absence
of azide revealed that only cellobiose was produced (Figure 4.17). Furthermore,
cellobiosyl-azide was found not to be a substrate. The lack of cellobiosyl-azide as a
product clearly distinguishes the behaviour of Asp 123 from that of Glu 127. Possible
roles for Asp 123 and the mechanism of azide and thiocyanate activation will be
discussed.
90
D123A
pNPC
cellobiosyl-azide
cellobiose
S
-
+
Figure 4.17. Hydrolysis of PNPC by Cex D123A in the presence and absence of
azide. Reactions mixtures were incubated overnight at 37°C in 50 mM phosphate
buffer, pH 7.0 with 500 ig/mL Cex D123A, 6 mM PNPC and 500 mM azide where
indicated. Thin layer chromatography was performed as described in section 2.10.1.
Standards lane (S): 2 p.L each of 6 mM PNPC, cellobiose and cellobiosyl-azide. with
azide, 2 jiL each. Reaction lanes (-) without azide, (+) with azide.
91
5. Discussion
5.1 Expression of the cex gene in Streptomyces lividans
The cex gene was subcloned and successfully expressed in S. lividans TK64 from
both the S. antibioticus melanin (mel) promoter of p1J702 and from the S. fradiae
aminoglycoside phosphotransferase (aph) promoter of p1J680. Cex was produced as a
soluble polypeptide, and retained full catalytic activity. Intracellular activity could not
be detected and Cex was efficiently secreted into the culture supernatant. Processing of
the leader peptide of Cex by Streptornyces resulted in equal proportions of the
polypeptide having either the natural amino-terminus or an amino-terminus with an
additional 2 amino acids. Studies of 40 secreted Streptomyces polypeptides have shown
that the -1 and +1 residues, relative to the processing site, are usually small neutral
amino acids, specifically alanines and serines (Brawner et al., 1991). Furthermore,
alanine is very often found at the -3 position. The normal (C. firni) processing site
(AQA t ATTLK) appeared to be well suited, therefore, for correct processing in
Streptomyces.
However, amino-terminal heterogeneity resulting from signal peptide
processing is not uncommon in foreign proteins expressed in Streptomyces (Brawner et
al., 1991). There was no indication, however, that the additional two amino-terminal
residues in Cex affected the activity of the enzyme.
Maximum exoglucanase activity in the culture supernatant of S. lividans TK64
[p1J680-cexl was reached following 40 h of growth, representing about 5.5 mg of Cex per
liter, based on the specific activity of the purified protein. The fragment of C. fimi DNA
in p1J680-cex and p1J702-cex contained the cex gene and its promoter. When the
fragment was inserted in the reverse orientation in p11702, the production of Cex was
reduced by about 90%. One interpretation of this result is that the cex promoter was not
92
efficiently recognized by S. lividans. It is of note that although S. lividans will recognize
many eubacterial promoters (Gusek and Kinsella, 1992; M. Bibb, personal
communication) Streptomyces promoters usually lack the typical -35 and -10 regions
found upstream of the cex gene and other eubacterial genes. Removal of the upstream
(740 bp) non-coding region of the cex gene and its promoter, and fusion of the sequence
encoding the leader peptide of Cex and the Cex cellulose binding domain to the aph
structural gene in p1J680 increased expression slightly (Ong et al., 1994). The increase in
expression might be a reflection of increased translational efficiency. In this case,
translation was initiated from the aph ribosome-binding site, and not the cex ribosome
binding site. The ribosome-binding site in cex (AGGAGG) is consistent with that of the
consensus ribosome-binding site deduced from 40 Streptomyces genes ((A/G)GGAGG)
(Anne and Van Mellaert, 1993) although it may less suitably located (4 nucleotides in
cex vs. 5 to 12 in Streptomyces genes relative to the initiation codon) for efficient initiation
of translation.
The level of expression from the aph promoter in p1J680-cex was about 6-fold
higher than that from the mel promoter of p1J702-cex. Other investigators have observed
the aph promoter to be superior to the mel promoter when used to express the same
fragment of foreign DNA. (L. Carlson, personal communication). The S. fradiae aph
promoter was the first promoter employed to direct the expression of a heterologous
gene in Streptomyces on the basis that the promoter reportedly directs the production of
its gene product to 10% of the total soluble cell protein (Brawner et at., 1991). It is clear,
however, that more study must be done in order to understand how the signals for
transcription initiation, translation and protein localization in Streptomyces effect the
efficiency of heterologous gene expression. Streptomyces is clearly a potentially very
useful host for the high level expression of heterologous genes. The level of expression
93
of the cex gene in Streptomyces , however, was not sufficiently high to offer an advantage
over E. coli as a host for the expression of this gene.
Native Cex from C. fimi is a glycoprotein. Interestingly, S. lividans also produced
a glycosylated Cex. Based on the apparent molecular mass of the recombinant protein,
the extent of glycosylation was very similar to that of Cex from C. firni. In addition, the
glycosyl groups on the Streptomyces-produced Cex were functionally similar to those of
C. fimi. The glycosylation afforded protection against proteolysis by C. fimi protease
particularly when the enzyme was bound to cellulose, and to a lesser extent when in
solution. This might be the result of greater steric hindrance of the protease when the
enzyme was immobilized on the cellulose surface. Previous work has shown that in the
case of non-glycosylated Cex from E. coli, proteolysis by either the C. fimi protease or
papain, occurs within and at the ends the PT linker region effectively separating the
catalytic domain and the cellulose-binding domain (Gilkes et al., 1991a). The linker
region may be susceptible to the C. fimi protease or papain because it presumably
adopts an extended conformation. It has subsequently been shown that in S. lividans
produced Cex, the PT linker region is in fact the site of glycosylation (Ong et a!., 1994).
S. lividans is known to produce native glycoproteins, for example, a
glucosidase (Mihoc and Kluepfel, 1990) and a xylanase (Kluepfel et a!., 1990). The
glycosylation of Cex observed in this study is significant in that it was the first report of
a heterologous glycoprotein produced by this organism (MacLeod et al., 1992).
Recently, a cellobiohydrolase from Microbispora bispora has also been shown to be
glycosylated by Streptomyces (Hu et al., 1993). Hosts which secrete and glycosylate
heterologous polypeptides expressed from cloned genes are potentially very useful both
for basic science and industrial applications. Although E. coli, is a commonly used host
for the expression of heterologous genes, it does not glycosylate polypeptides. C. fimi is
a potentially useful host for the expression of genes encoding glycoproteins, but suitable
94
vectors have yet to be developed. Sacchnrornyces cerevisiae, another commonly used host,
does produce glycoproteins. However, S. cerevisine may hyperglycosylate them, giving
them undesirable characteristics (Curry et al., 1988). The use of mammalian cells for the
expression of cloned genes often yields polypeptides with the correct, or at least
acceptable, patterns and levels of glycosylation (Goochee et a!., 1991). However, animal
cell culture processes are complex and expensive. It is clear, therefore, that the ability of
Streptomyces to glycosylate heterologous polypeptides is a very useful feature of this
bacterium.
5.2
Catalysis
5.2.1 The catalytic nucleophile
Glutamic acid 233 in Cex, the putative catalytic nucleophile, was targeted by sitedirected mutagenesis on the basis of amino acid sequence alignments. Glu 233 in Cex is
part of a conserved ITE motif within family F cellulases and xylanases. Glu 358 in Abg
from A robacterium, also part of a conserved ITE motif within its family, was identified
as the catalytic nucleophile in this retaining enzyme by inactivation with the
mechanism-based inactivator 2F-DNPG (Withers and Street, 1988; Withers et a!, 1990).
The kinetic parameters found in this study for mutants of Glu 233 in Cex were
consistent with those found for similar mutants of the catalytic nucleophile in Abg
(Withers et al., 1992). It is clear from these studies that the shortening of the carboxylate
side chain of the catalytic nucleophile by as little as lÀ (a Glu
-->
Asp mutation) has a
dramatic impact on the activity of the enzyme. Shortly after initiation of this study, the
labeling of Glu 233 with the inactivator 2F-DNPG (Tull et a!., 1991), and more recently
with 2F-DNPC (Tull and Withers, 1994) provided unequivocal evidence that Glu 233
was the catalytic nucleophile in Cex.
95
5.2.2 The acid/base catalyst
Glu 127 was proposed as the acid/base catalyst in Cex on the basis of amino acid
sequence alignments. The kinetic parameters determined for mutants at position 127
provide firm support for the assignment of Glu 127 as the acid/base catalyst in Cex.
The kcat/Km values for substrates with various leaving group abilities provide an
estimation of the effect of the mutation on the glycosylation step (where acid catalysis
occurs). With DNPC, which presumably does not require acid-catalytic assistance for
initial bond cleavage, the Glu 127 mutants exhibited kcat/Km values similar to that of
the wild-type enzyme. With PNPC, the leaving group of which has an intermediate
pKa, the mutants exhibited somewhat reduced kcat/Km values relative to the wild-type
enzyme. When 4-BrPC was tested with the E127A mutant, a substrate which
presumably requires acid-catalytic assistance for initial bond cleavage, the kcat/Km
value was dramatically reduced relative to the wild-type enzyme. Therefore, the rate
constants for glycosylation were affected relatively little for substrates not requiring
acid catalysis, and dramatically for substrates requiring acid catalysis. An accumulation
of the glycosyl-enzyme intermediate would be expected if the rate of the
deglycosylation step was reduced to a greater extent than that of the glycosylation step,
and should also result in decreased Km values. Relative to the wild-type enzyme, the
decreases in Km for the substrates (about 200-fold for DNPC, 20-fold for PNPC and
unchanged with 4-BrPC) are consistent with this interpretation. In other words, with a
good leaving group (i.e. DNP) the rate of the glycosylation step remained unchanged
relative to the wild-type. Consistent with this was the finding that E127A was in fact
inactivated with 2F-DNPC at a rate at least as fast as that of the wild-type enzyme. The
reductions in kcat values observed for DNPC and PNPC with the Glu 127 mutants (200400 fold) therefore reflect primarily the effect upon the deglycosylation step of the
reaction, due to the removal of general base catalytic assistance.
96
Sodium azide significantly enhanced the rate of hydrolysis of PNPC, DNPC and
DNPG by uncharged Glu 127 mutants. With increasing azide concentration, both kcat
and Km values rose concurrently, the net result of which was that kcat/Km values
remained essentially constant across the range. Therefore, the kinetic parameters for the
mutants in the presence of sodium azide were consistent with azide acting to increase
the rate of the deglycosylation (base-catalysis) step. The proposal that azide occupied a
vacant anionic site created by removal of the acid/base catalyst, and acted instead of
water (OH-) as a nucleophile in the hydrolysis of the glycosyl-enzyme intermediate was
confirmed by the detection of the 13-linked cellobiosyl-azide (13-glucosyl-azide with
DNPG as substrate). The fact that only f3-linked adducts were detected confirms that
azide attacked only from the
13 face of the substrate, a result consistent with a location of
the acid/base catalyst (Glu 127) on this same face.
Interestingly, the rate-enhancement with azide was specific to the Ala and Gly
mutants. A rate enhancement was not seen with the wild-type enzyme or with the
E127D mutant. Presumably the glutamate or aspartate carboxylates prohibited the
access of azide through electrostatic repulsion. The lack of a rate enhancement with
E127D was unlikely due to a steric effect in light of the finding of Cupples
et al. (1990)
that azide could in fact enhance the rate of an E461Q mutant in 13-galactosidase, an
uncharged, but equally bulky substitution. It is likely, therefore, that an E127Q or
E127N mutation in Cex would have a similar effect.
Although several nucleophiles other than azide were tested for their effects on
the E127A and E127G mutants in the current study, only azide was found to
significantly increase rates of hydrolysis and form adducts detectable by TLC. It’s
likely, however, that other nucleophiles should be able to function in this manner.
Huber and Chivers (1993) recently reported that similar 13-galactosyl adducts were
produced by uncharged E461 mutants of f3-galactosidase in the presence of nucleophiles
97
including azide, acetate and imidazole to name a few. It was further suggested that
with certain nucleophiles, f3-galactosyl adducts such as 13-galactosyl-acetate were
formed but are unstable.
The techniques employed in this study may be applicable to other glycosidases
belonging to sequence-related families (Henrissat, 1991). The approach would involve
generation of alanine or glycine mutants of conserved glutamic and aspartic acids. The
mutants would then be screened for the generation of new products of hydrolysis in the
presence of azide and other anionic nucleophiles. Positive mutants should then be
subjected to the following kinetic analysis. kcat and Km values should be measured for
a pair of substrates, one of which requires acid catalysis and one of which does not.
Values of kcat/Km for these two substrates should differ greatly. For the activated
substrate, the kcatlKm value should be much higher than that for the other. Values of
kcat and Km for highly activated substrates with these mutants should increase
dramatically with increasing azide concentration, and level off at a value dependent on
the leaving group ability of the aglycon. This sort of approach could facilitate
identification of the acid/base catalyst in other glycosidases. The acid/base catalyst in a
family G xylanase from Bacillus circulans (Glu 172) was deduced from examination of
the crystal structure (Wakarchuk et al., 1994). Uncharged mutants at this position tested
for similar azide effects in light of this investigation resulted in the generation of 13xylobiosyl-azide from p-nitrophenyl 13-xylobioside (PNPXX). In addition, preliminary
studies with acid/base catalyst mutants of Agrobacterium f3-glucosidase have yielded
similar results (S. Withers, personal communication).
The generation of the new f3-linked product in this study is clearly a significant
result. A rate-enhancement in the presence of the nucleophile in which a conserved
acidic residue has been mutated does not constitute a positive screen for an acid/base
catalyst. First, in the case of a retaining enzyme, the substrate used must be rate-
98
limiting for the deglycosylation step with the mutant in question. For example,
enhancement of the rate of hydrolysis with PNPG was not observed with the mutants in
the presence of azide, although glucosyl-azide could be detected by TLC. This result is
not surprising given that the rate-limiting step for this substrate with the wild-type
enzyme is glycosylation (Tull and Withers, 1994). Activation of the deglycosylation step
by azide, therefore, would not be expected to affect the steady-state rate of hydrolysis of
PNPG. Second, anions clearly will enhance the steady-state rates of mutants at other
positions. In this study, both azide and thiocyanate enhanced the rate of mutants of
Asp 123, a conserved acidic residue in close proximity to Glu 127. In this case however,
cellobiose was the only product in the absence or presence of azide. No concomitant
production of cellobiosyl-azide or cellobiosyl-thiocyanate was observed. In this case
azide or thiocyanate might bind at the site of the missing side chain restoring local
charge requirements. Asp 123, however, clearly plays a role in facilitating catalysis.
Conceivably, this residue might be involved in binding the substrate, or play a role in
maintaining the correct environment (i.e. pKa’s) at the acid/base catalyst or the catalytic
nucleophile possibly through a hydrogen bonding network. It is of note that in B.
circulans xylanase, there is reportedly a conserved acidic residue found within the active
site, in addition to the acid/base catalyst and the catalytic nucleophile (Wakarchuk et al.,
1994).
An inverting glycosidase, such as CenA from C. fimi, also hydrolyses f3-1,4glycosidic linkages. In this type of enzyme, however, the acid catalyst and the base
catalyst are assumed to be different amino acid residues (Sinnott 1990). Conceivably, a
similar approach to that employed in this study could be used to aid in the
identification of the base-catalyst in an inverting glycosidase. Since azide was shown to
increase the rate of the deglycosylation step in Cex (where base-catalysis occurs), anions
such as azide should also enhance the rate of an uncharged base-catalyst mutant in an
99
inverting enzyme.
In
contrast to a retaining enzyme, the new product formed would be
linked in an c’. configuration.
The E127A mutant of Cex is potentially useful for several reasons. Firstly, the
enzymatic production of 13-cellobiosyl-azide provides an alternative to the synthetic
route (T. Lindhorst, personal communication); the mutant, as discussed, may allow the
transfer of other anions to the sugar. Secondly, this mutant may be particularly suited
for co-crystallization with a substrate such as DNPC since it tends to accumulate the
glycosyl-en.zyme intermediate. The catalytic domain of Cex was crystallized in the
absence of substrate (Bedarkar et a!., 1992). Examination of the enzyme/substrate
complex should provide valuable information about the topology of the active site
relative to its substrate and should confirm the presence of the covalently bound
glycosyl-enzyme intermediate. Although the wild-type enzyme could be crystallized in
the presence of the inactivator 2F-DNPC, DNPC presumably represents a more natural
substrate. Lastly, enzymes which will transglycosylate to form oligosaccharides are
potentially commercially valuable. Cex has the ability to carry out a weak
transglycosylation reaction which essentially competes with the hydrolysis of the
glycosyl-enzyme intermediate (Tull and Withers, 1994). Since the deglycosylation step
of Glu 127 mutants of Cex with the appropriate substrate is slowed down, this mutant
could serve as a starting point for the engineering of an enzyme capable of synthesizing
oligosaccharides.
In summary,
based on amino acid sequence alignments and on knowledge of the
types of residues likely to be involved in catalysis by a retaining glycanase, mutants
have helped to identify the two key catalytic residues in Cex
in the absence of three
dimensional structural information. Preliminary X-ray crystallographic data analyzed
following the preparation of this thesis in fact confirm the presence of Glu 233 and Glu
127 within the active site cleft of the enzyme (White et a!,, 1994). The two residues are
100
oriented such that their side-chains face each other with an appropriate separation.
Based on the known structures of other retaining glycanases (Campbell et a!., 1993;
Wakarchuk et a!., 1994) these residues are suitably positioned to act as the catalytic
nudeophile and acid/base catalyst respectively. The Asp 123 side chain is not similarly
exposed to the active site (White et a!., 1994).
The approach used in identifying the acid/base catalyst in Cex in the absence of
three dimensional information may in fact be applicable to other glycosidases, both
retaining and inverting. This study will clearly serve to facilitate an understanding of
the roles of active site residues in Cex, pending the resolution of the crystal structure.
101
6. Appendix
6.1 Determination of kcat and Km: an example:
The example below shows the data obtained (as outlined in section 2.9) for the
hydrolysis of 2,4-DNPG catalysed by Cex E127A (at 4 tg/mL) in the presence of 3 mM
sodium azide: The initial rate of production of DNP 0
(v
)
, measured by AA400/min,
was converted to nmol DNP released/mm according to the following relationship:
=A
cL
where:
For DNP, at 2.
0
v
=
=
E
A
c
L
400 nm, pH 7.0,
=
nmol DNP released/mm
extinction coefficient
absorbance
concentration
(M)
cell path length
(cm)
=
=
=
=
10 900 1
.cm therefore (for a 1 cm path cell):
M
,
AA400/min
0.01 09
=
The following data were obtained [curve 11:
1.4
1.2
E
1
0.8
S
[curve 1]
0.6
0
>
0.4
0.2
0
0
20
40
DNPG (M)
60
102
Assuming a steady state, the velocity of an enzyme-catalysed reaction can be expressed
by the Michaelis-Menten equation:
0
V
Vpx [S]
Km + IS]
=
[3]
where v
0 is the velocity (measured as the initial rate of production of DNP in this case),
Vmax is the maximal velocity, Km is the Michaelis constant and [SI is the concentration
of free substrate (DNPG). The rate constants
and Km can be extracted graphically
from a linearly-transformed plot of the data shown above. A common linearized form
of the Michaelis-Menten equation is its reciprocal (the Lineweaver-Burk equation).
When both sides of the Lineweaver-Burk equation are multiplied by [SI this yields:
[5]
[S]
Vmax
+
V
0
V
[4]
Plotting [SI/v
0 vs. [SI results in a linear curve (a Hanes plot; curve 2), the slope
of which is
1 /Vmax, the y-intercept
=
Km/Vmax and the x-intercept = -Km.
50
.E
E
¶1
40
30
20
[curve 2]
10
-20-10
=
0
10
20
30
40
S (jiM DNPG)
50
0.9992
60
103
Therefore, from curve [2]:
Km
and
=
9.57.i.M
=
1.57 1
nmol.miir (for 4 pg enzyme)
=
0.4
1 (for 1 mg enzyme)
jimol.min
The maximal velocity Vm is directly proportional to the total enzyme concentration
[ET] such that Vm
therefore
kcat
=
=
=
=
kcat [ET]
Vmax
[ET]
0.4 j.tmol.m1n
1
0.02122 i.tmol
(where 1 mg enzyme
= 2.122 x 10 2 iimol)
18.85 min
1
Values of kcat and Km reported in this thesis were calculated directly by non
linear regression by fitting the data to the Michaelis-Menten equation (as shown in
curve [11) using the computer program Grafit 2.0 (Erithacus Software Ltd., Staines,
U.K.). In this case the values of keat and Km were determined to be 18.73 m1n
1 and 9.35
.tM respectively, in close agreement with the values obtained above.
104
6.2 Interpretation of kcat, Km and kcat/Km:
glycosylation
1
k
E+S
deglycosylation
2
k
ES
3
k
E-G
E+P
1
k-ROH
+H20
Equation [1] is representative of the mechanism of Cex as shown in Figure 4.1.
kcat, Km and kcat/Km can be expressed in terms of the rate constants of the individual
steps as indicated in equation [1] (Fersht, 1985; Schowen, 1978; Sinnot, 1990).
kcat
=
Km
=
kcat
Km
=
3
2
k
k +k
2
3
[2]
1
(
3
k
k- +k
)
2
(
1
k
2
)
3
i-k
k
[3]
2
k
1
+k
1
k2
[4]
When there is a rapid, reversible association step (formation of the E.S complex)
and hydrolysis of the glycosyl-enzyme intermediate is rate limiting, as in the case of
hydrolysis of cellobiosides by Cex (Tull and Withers, 1994), the kinetic relationships
would be k-p>>k
2 and 3
>>k When these conditions are applied to equations [2],
2
k
.
and [4] above, the kinetic constants can be reduced to:
kcat
3
k
[51
13]
105
Km
=
=
kcpt
Km
=
=
1
k
3
k
2
1
k
where k-i
1
k
=
Ks
jKs)
2
k
[6]
1
k
2
1
k
k2
Ks
[7]
When there is a rapid, reversible association step and formation of the glycosyl
enzyme intermediate is rate-limiting, as in the case of hydrolysis of glucosides by Cex
(Tull and Withers, 1994) the kinetic relationships would be k1
this case, equations [2],
>>
k and k
3
>>
. In
2
k
[31 and [41 can be reduced to:
kcat
=
Km
=
kcpt
Km
=
=
2
k
[81
k
1
1
k
Ks
[9]
1
k
2
k-i
2
k
Ks
[101
It can be seen from the above examples that kcat is a reflection of the rate of the
rate determining step. It can also be seen from equations [7] and [101 that kcat/Km
reflects the rate of the formation of the glycosyl-enzyme intermediate from the free
enzyme and substrate. This applies whether glycosylation or deglycosylation is rate-
106
limiting. From equation [61, the Km value is proportional to k
2
/
3
k when
deglycosylation is rate-limiting. This is consistent with the interpretation of Km as
outlined in the introduction.
6.3 Nucleotide and amino acid sequence of mature (processed) Cex
1
1/1
GCG ACC
ala thr
61/21
CCC AAC
pro asn
121/41
CTC CCC
val ala
181/61
CCC CCC
gly ala
241/81
ACG CTC
thr leu
301/101
GAG ACC
glu ser
361/121
TCC TCC
ser trp
421/141
TTC CAC
phe gin
481/161
CAC CCC
asp pro
541/181
AAC TCC
asn ser
601/201
CCC TTC
gly phe
661/221
CCC TTC
arg phe
721/241
CCC TCC
pro ser
781/261
TCC ATC
cys met
841/281
TCC CTC
trp vai
31/11
ACC CTC AAG GAG CCC GCC GAC CCC GCC CCC
thr leu lys glu ala ala asp gly ala gly
91/31
CCC CTC TCC GAG GCG CAG TAC AAC GCG ATC
arg leu ser glu ala gin tyr lys ala lie
151/51
GAG AAC CCC ATC AAC TCG GAC CCC ACC GAG
glu asn ala met lys trp asp ala thr glu
211/71
CGC CAC CGC CTC CCC ACC TAC CCC CCC CAC
gly asp arg val ala ser tyr ala ala asp
271/91
GTA TGC CAC TCG CAG CTC CCC CAC TCC CCC
val trp his ser gln leu pro asp trp ala
331/111
CCC ATG CTC AAC CAC CTC ACC AAG CTC CCC
ala met val asn his val thr lys val ala
391/131
CAC CTC CTC AAC GAG CCC TTC CCC CAC CCC
asp val val asn glu ala phe ala asp gly
451/151
CAC AAC CTC CCC AAC CCC TAC ATC GAG ACC
gin lys leu gly asn gly tyr ile glu thr
511/171
ACC CCC AAC CTC TCC ATC AAC CAC TAC AAC
thr ala lys leu cys ile asn asp tyr asn
571/191
CTC TAC CAC CTC CTC AAC CAC TTC AAC CCC
leu tyr asp leu val lys asp phe lys ala
631/211
CAG TCC CAC CTC ATC CTC CCC CAC CTC CCC
gin ser his leu ile val gly gin val pro
691/231
CCC CAC CTC CCC CTC CAC CTC CCC ATC ACC
ala asp leu gly val asp val arg ile thr
751/251
CAC CCC ACC AAC CTC CCC ACC CAC CCC CCC
asp ala thr lys leu ala thr gin ala ala
811/271
CAC CTC ACC CCC TCC CAC CCC CTC ACC CTC
gin val thr arg cys gin gly val thr val
871/291
CCC CAC CTC TTC CCC CCC GAG CCC CCC CCC
pro asp val phe pro gly glu gly ala ala
CCC GAC TTC GGC TTC CCC CTC GAC
arg asp phe gly phe ala leu asp
GCC GAC ACC GAG TTC AAC CTC GTC
ala asp ser glu phe asn leu val
CCC TCG CAC AAC AGC TTC TCC TTC
pro ser gin asn ser phe ser phe
ACC GCC AAC GAG CTG TAC GGC CAC
thr gly lys glu leu tyr gly his
AAC AAC CTC AAC CCC TCC CCC TTC
lys asn leu asn gly ser ala phe
GAC CAC TTC CAC CCC AAC CTC CCC
asp his phe glu gly lys vai ala
CCC CCC CCC CCC CAC GAC TCC CCC
gly gly arg arg gln asp ser ala
CCC TTC CCC CCC CCA CCT CCC CCC
ala phe arg ala ala arg ala ala
CTC CAC CCC ATC AAC CCC AAC ACC
val glu gly ile asn ala lys ser
CCC CCC CTC CCC CTC GAC TCC CTC
arg gly val pro leu asp cys val
CCC CAC TTC CCC CAC AAC CTC CAC
gly asp phe arg gin asn ieu gin
GAG CTC CAC ATC CCC ATC CCC ACC
glu leu asp ile arg met arg thr
CAC TAC AAC AAC GTC CTC CAC CCC
asp tyr lys lys val vai gin ala
TCC CCC ATC ACC CAC AAC TAC TCC
trp gly ile thr asp lys tyr ser
CTC CTG TGG CAC CCC ACC TAC CCC
leu val trp asp ala ser tyr ala
107
901/301
AAG AAG CCC
lys lys pro
961/321
ACC ACG CCC
thr thr pro
1021/341
TGC CAG GTG
cys gin val
1081/361
AAG AAC ACG
iys asn thr
1141/381
CAG CAG GTC
gin gin vai
1201/401
CCC AAC CCC
arg asn ala
1261/421
GGC TCC CAC
gly ser his
1321/441
ACG GTC CCC
thr vai giy
931/311
CCC TAC CCC CCC GTG ATC GAG CCC TTC CCC
ala tyr ala ala val met glu ala phe gly
991/331
ACC CCG ACC CCC ACC ACG CCC ACG CCC ACC
thr pro thr pro thr thr pro thr pro thr
1051/351
CTC TGG CCC CTC AAC CAG TGG AAC ACC CCC
ieu trp gly val asn gin trp asn thr gly
1111/371
TCC TCC GCT CCC CTC GAC CCC TGC ACC CTC
ser ser ala pro val asp gly trp thr leu
1171/391
ACC CAC CCC TGC AGC TCC ACC GTC ACC CAC
thr gin ala trp ser ser thr val thr gin
1231/411
CCC TGG AAC CCC TCC ATC CCC CCC CCC CCC
pro trp asn gly ser iie pro ala gly gly
1291/431
ACC CCC ACC AAC CCC CCC CCC ACC CCC TTC
thr gly thr asn ala ala pro thr ala phe
TCA
STOP
GCG AGC CCC ACG CCC ACG CCC
ala ser pro thr pro thr pro
CCG ACG TCC CCT CCC CCC CCC
pro thr ser gly pro ala gly
TTC ACC CCC AAC GTC ACC CTC
phe thr ala asn val thr val
ACC TTC ACC TTC CCC TCC CCC
thr phe ser phe pro ser gly
TCC CCC TCC CCC CTC ACC GTC
ser gly ser ala val thr val
ACC CCC CAG TTC CCC TTC AAC
thr ala gin phe gly phe asn
TCC CTC AAC CCC ACC CCC TCC
ser leu asn gly thr pro cys
(enuf is enuf!)
‘Adapted from O
Neill et al. (1986b). Corrections to the original gene sequence have
t
been deposited to GenBank (New accession # L11080). Numbering refers to the
nucleotide/amino acid number from the mature N-terminus. Glu 127 and Glu 233 are
indicated in bold type.
108
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