BIOB12H3S:
Cell
&
Molecular
Biology
Laboratory
Biochemistry
module
(PDF
#7)
Introduction
to
biochemical
techniques
The
techniques
of
biochemistry
are
used
to
purify
and
characterize
various
types
of
small
molecules
and
macromolecules,
and
to
put
the
properties
of
these
molecules
into
a
biological
context.
This
might
be
equated
with
how
the
structure
and
chemical
properties
of
a
molecule
determine
its
structure
and
function
in
a
living
cell.
The
macromolecules
include
proteins,
nucleic
acids,
sugars,
fats
and
other
cellular
components.
Because
of
the
differences
in
size,
composition,
solubility
in
water,
and
other
physical
and
chemical
parameters,
different
approaches
are
taken
to
study
these
groups,
and
different
techniques
and
instruments
are
used
in
experiments.
In
nearly
all
cases
involving
multicellular
organisms,
one
of
the
first
steps
is
to
secure
the
tissue
or
cell
type
of
interest.
If
one
was
interested
in
studying
the
basic
cellular
process
of
meiosis,
floral
buds
might
be
the
starting
material
from
plants,
while
the
testes
could
be
the
starting
material
from
animals.
Having
secured
enough
starting
material,
the
next
step
is
to
lyse
the
cells
under
conditions
that
will
preserve
the
desired
structures
and/or
the
activities
of
the
desired
enzymes
or
other
macromolecules.
Usually
this
is
performed
by
using
a
detergent,
but
this
may
also
be
coupled
with
enzymatic
lysis,
physical
methods
like
grinding
in
sand
or
a
blender,
sonication,
or
passing
the
cells
through
a
small
hole
under
high
pressure.
Once
the
cells
are
broken
open,
the
resulting
suspension
is
termed
a
homogenate.
Depending
on
the
component
that
you
are
interested
in
purifying,
you
may
include
a
variety
of
metabolites
or
reagents
in
your
lysis
buffer
to
preserve
structure
and
function.
If
you
are
only
interested
in
looking
at
the
total
protein
population
by
SDS-polyacrylamide
gel
electrophoresis
you
may
choose
to
lyse
the
cells
and
denature
the
proteins
in
one
step
(this
is
what
you
will
be
doing
in
this
laboratory).
Once
the
cells
are
broken
open,
in
many
cases
subcellular
fractionation
is
undertaken.
This
usually
centers
on
differential
centrifugation
to
sediment
subcellular
components
that
differ
in
their
size
and
density.
Centrifugation
generates
a
liquid
fraction
(the
supernatant),
and
a
solid
or
particulate
fraction
(the
pellet).
Subcellular
fractionation
is
always
coupled
with
either
a
marker
enzyme
assay
and/or
a
functional
assay
for
the
molecule
of
interest.
Marker
enzyme
assays
are
used
to
screen
for
the
presence
of
a
specific
component
in
the
various
fractions
and
is
based
upon
knowing
that
a
positive
test
for
that
marker
will
identify
those
fractions
that
contain
a
specific
subcellular
fraction
or
organelle
(for
example
a
positive
test
for
the
enzyme
cytochrome
oxidase
indicates
that
the
fraction
contains
mitochondria).
The
functional
assay
is
generally
devised
by
the
1 investigator
to
screen
for
the
presence
of
the
component
of
interest.
Obviously,
in
order
to
devise
a
test,
one
must
know
something
about
the
properties
and/or
function
of
the
molecule.
An
example
of
a
functional
assay
would
be
to
test
the
ability
of
an
inhibitor
to
alter
enzyme
activity.
The
β-‐galactosidase
study
you
carried
out
last
laboratory
is
another
example
of
a
functional
assay.
For
the
purification
of
proteins,
the
supernatant
and/or
pellet
fractions
of
interest
are
subjected
to
the
techniques
of
protein
chemistry
in
order
to
begin
to
fractionate
the
many
proteins
found
in
the
starting
material.
These
techniques
are
diverse,
but
most
rely
on
physiochemical
properties
of
the
proteins.
These
include
the
solubility
of
proteins
in
organic
solvents
and
concentrated
salt
solutions,
and
adsorption
to
particular
materials.
Chromatography
methods
are
nearly
always
required.
These
include
ion
exchange,
gel
filtration,
and
affinity
chromatography.
Electrophoresis
is
employed
quite
often,
and
certainly
at
the
end
of
the
purification
to
determine
the
number
of
polypeptides
in
the
active
fractions.
Specifically,
Sodium
dodecyl
sulfate
(SDS)-‐polyacrylamide
gel
electrophoresis
(PAGE)
can
be
used
to
determine
the
molecular
weight
of
protein
subunits.
In
this
Biochemistry
module
you
will
use
the
same
three
strains
used
in
Lab
6A.
Each
pair
will
be
assigned
one
of
three
E.coli
strains,
wild
type,
lacI
mutant
or
LacZ
mutant.
You
will
not
be
told
which
strain
you
have
been
given.
Each
pair
will
induce
one
sample
with
IPTG
and
leave
the
other
uninduced.
You
will
lyse
the
cells,
prepare
the
proteins
for
SDS-‐PAGE.
The
proteins
from
the
uninduced
and
induced
cells
will
be
separated
on
a
10%
SDS-‐polyacrylamide
gel
and
you
will
compare
the
protein
expression
profiles.
In
addition
you
will
use
an
in-vivo
β-galactosidase
assay
using
IPTG
and
X-‐gal
to
look
for
enzyme
activity
in
your
cells.
2 LAB
8B:
Growth
&
Induction
of
cells
with
IPTG
and
preparation
of
a
homogenate
(per
pair
of
students)
Materials
needed
per
pair
E.coli
cells
at
an
OD 600
of
0.8-‐0.9
in
5
mls
of
M9-‐glycerol
medium.
M9
medium
IPTG
Microfuge
tubes
Microcentrifuge
SDS
sample
buffer
containing
SDS,
B-‐mercaptoethanol,
glycerol
and
Brompohenol
blue
(tracking
dye)
in
a
Tris
buffer
at
pH
6.8
Heating
block
1. Take
two
3
ml
samples
of
the
unknown
E.coli
strain
assigned
(strain
1,
2
or
3).
Add
one
ml
of
M9
medium.
Label
one
tube
uninduced
and
the
other
induced.
Make
sure
you
name
is
on
the
tubes
2. Place
the
two
samples
in
the
37°C
water
bath
and
allow
the
cells
to
grow
while
shaking
for
40
minutes.
3. Remove
the
tube
labeled
induced
from
the
water
bath
and
add
500μl
of
IPTG
to
this
tube.
4. Return
the
tube
to
the
37°C
water
bath
5.
Incubate
the
cells
at
37°C
for
60-‐70
minutes
(allows
cells
to
continue
to
grow
and
the
one
culture
to
be
induced
to
express
the
Lac
operon)
6.
Remove
both
tubes
of
cells
from
the
water
bath
7. Label
two
1.5
ml
microfuge
tubes
with
your
initials
and
the
label
“uninduced”
or
“induced”
8. Gently
mix
the
culture
tubes
taken
from
the
water
bath
and
remove
1.5ml
of
culture
into
the
appropriately
labeled
microfuge
tubes
9. Place
the
two
microfuge
tubes
containing
the
cells
into
the
microcentrifuge
with
other
members
of
your
laboratory.
Make
sure
the
centrifuge
is
balanced
(have
the
TA
check!)
10.Set
the
centrifuge
to
5000
rpm
and
the
time
for
5
min.
3 11.
When
centrifugation
is
complete,
pour
off
the
supernatant
into
the
appropriate
discard
container
for
bacterial
waste.
Make
sure
you
do
not
disturb
the
pellet
when
decanting
the
supernatant.
12.Add
another
1.5
ml
of
the
appropriate
bacterial
culture
to
the
pellet
(after
this
step
you
will
have
spun
down
a
total
of
3
ml
of
culture)
and
repeat
steps
10
and
11
13.To
each
of
the
two
pellets
(induced
and
uninduced)
add
60
μl
of
SDS
sample
Buffer
containing
bromophenol
blue.
Mix
to
resuspend
the
cells.
The
SDS,
which
is
an
anionic
detergent,
will
lyse
the
cells
and
coat
the
proteins.
The
solution
should
clear.
The
role
of
the
SDS
sample
buffer
is
discussed
further
in
Lab
9A.
14.Place
your
two
samples
into
the
heating
block
set
at
100°C
and
incubate
for
5
minutes.
This
will
aid
in
the
lysis
of
the
cells
and
denature
the
proteins.
15.At
this
point,
your
proteins
are
ready
for
loading
onto
the
SDS-‐
polyacrylamide
gels.
Store
the
samples
in
the
rack
provided.
The
samples
will
be
place
at
-‐20°C
until
the
next
lab.
In
Lab
9A,
you
will
perform
SDS-‐
PAGE.
4 LAB
9A:
SDS-PAGE
(Sodium
dodecyl
sulfate-Polyacrylamide
Gel
Electrophoresis)
&
plating
on
X-gal
plates
In
most
research
labs,
SDS
polyacrylamide
gel
electrophoresis
is
a
technique
central
to
many
experiments
in
cell
and
molecular
biology,
and
is
employed
to
examine
protein
profiles.
This
can
be
accomplished
by
running
the
gel
and
staining
it
with
a
non-‐specific
stain
like
Coomassie
Brilliant
Blue.
Alternatively,
the
samples
may
be
radiolabeled
and
after
electrophoresis,
autoradiography
can
be
employed
to
examine
the
radiolabeled
components.
Finally,
in
many
instances
where
complex
mixtures
of
proteins
exist
(nearly
always
the
case!),
after
electrophoresis
the
proteins
are
transferred
to
a
solid
support
and
immunoblotting
with
a
specific
antibody
is
undertaken
to
screen
for
the
presence
of
a
specific
molecule
or
family
of
molecules.
Most
research
labs
make
their
own
gels.
This
has
three
disadvantages
in
an
undergraduate
lab.
First,
the
gel
mold
must
be
precisely
made
and
the
gel
must
be
cast
correctly
or
it
will
leak
out
of
the
mold.
Second,
the
acrylamide
monomer
is
a
potent
neurotoxin
and
great
care
must
be
exercised
in
handling
it.
Third,
it
takes
several
hours
to
prepare
a
gel,
time
which
we
don’t
have.
Thus
we
will
use
commercially
available,
precast
gels
from
a
company
called
Novex.
Procedure :
We
will
use
the
Novex
electrophoresis
modules
and
Novex
precast
gels.
The
SDS-‐
gels
were
made
to
contain
a
final
concentration
of
10%
acrylamide.
Below
are
some
diagrams
of
how
to
assemble
the
gel
in
the
apparatus,
though
your
TA
will
likely
guide
you
through
it.
There
are
a
few
things
to
keep
in
mind.
1. Be
sure
to
wear
gloves
when
handling
the
gel.
2. Hold
the
gel
by
the
sides
only
(i.e.,
along
the
edges,
not
on
the
flat
surface).
3. You
must
take
the
gel
out
of
its
plastic
pouch,
peel
the
tape
off
the
bottom,
and
in
one
quick
motion,
take
the
comb
out
of
the
cassette.
4. Now
you
must
work
fairly
quickly
to
assemble
the
apparatus,
and
flood
the
wells
with
buffer
(so
the
gel
doesn’t
dry
around
the
wells
leading
to
cracking
and
possibly
the
mixing
of
adjacent
samples).
5. Have
about
1
liter
of
1X
running
buffer
ready
(see
below).
Running
buffer:
We
have
prepared
the
running
buffer
for
you
as
a
10X
stock.
It
is
located
on
the
side
bench
and
is
labeled
10X
SDS
PAGE
buffer.
The
formula
for
1X
is
25mM
Tris,
192mM
glycine,
1%
SDS.
To
make
1
liter,
simply
put
100ml
of
the
10X
stock
into
a
1
liter
graduate
cylinder
and
introdu
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