Transcript Crystallization Laboratory - UCLA

Crystallization Laboratory
The agony and the ecstasy of
protein crystallization
M230D,Jan 2008
Goal: crystallize Proteinase K and its
complex with PMSF
• Non-specific serine
protease frequently
used as a tool in
molecular biology.
• PMSF is a suicide
inhibitor. Toxic!
• Number of amino
acids: 280
• Molecular weight:
29038.0
• Theoretical pI: 8.20
MAAQTNAPWGLARISSTSPGTSTYYYDESAGQGSCVYVIDTGIEASH
PEFEGRAQMVKTYYYSSRDGNGHGTHCAGTVGSRTYGVAKKTQLFGVKVLDDNGS
GQYSTIIAGMDFVASDKNNRNCPKGVVASLSLGGGYSSSVNSAAARLQSSGVMVA
VAAGNNNADARNYSPASEPSVCTVGASDRYDRRSSFSNYGSVLDIFGPGTSILST
WIGGSTRSISGTSMATPHVAGLAAYLMTLGKTTAASACRYIADTANKGDLSNIPF
GTVNLLAYNNYQA
Ala (A) 33
11.8%
Arg (R) 12
4.3%
Asn (N) 17
6.1%
Asp (D) 13
4.6%
Cys (C)
5
1.8%
Gln (Q)
7
2.5%
Glu (E)
5
1.8%
Gly (G) 33
11.8%
His (H)
4
1.4%
Ile (I) 11
3.9%
Leu (L) 14
5.0%
Lys (K)
8
2.9%
Met (M)
6
2.1%
Phe (F)
6
2.1%
Pro (P)
9
3.2%
Ser (S) 37
13.2%
Thr (T) 22
7.9%
Trp (W)
2
0.7%
Tyr (Y) 17
6.1%
Val (V) 19
6.8%
Why is it necessary to grow a
crystal to solve a protein
structure by X-ray diffraction ?
Protein crystals are ordered (periodic) arrays
of protein molecules.
protein in solution.
One
dimensional
order
Two
dimensional
order
Three
dimensional
order
Crystals are needed to amplify the diffraction
signal.
Diffraction
from a crystal
is strong.
Diffraction
from a single
molecule is
weak.
What is the most important
property of a crystal ?
It is the “order” of a crystal that ultimately
determines the quality of the structure.
Order- describes the degree of regularity (or periodicity) in the arrangement of
identical objects.
supersaturated
protein solution.
DISORDERED
One
dimensional
order
Two
dimensional
order
Three
dimensional
order
ORDERED
“Order” is perfect when the crystallized object is
regularly positioned and oriented in a lattice.
c
a
b
When a crystal is ordered, strong diffraction
results from constructive interference of photons.
Interference is constructive because path lengths differ
by some integral multiple of the wavelength (nl).
detector
5
crystal
4
3
2
6
1
5
4
3
Incident X-ray
2
6
1
7
5
4
3
2
1
This situation is possible only
because the diffracting objects
are periodic.
Nonregularity in orientation or position
limits the order and usefulness of a crystal.
Perfect order
Rotational disorder
Translational disorder
Disorder destroys the periodicity leading to
Streaky, weak, fuzzy, diffraction.
When a crystal is disordered, poor diffraction
results from destructive interference of photons.
Interference is destructive because path lengths differ
by non integral multiple of the wavelength (nl).
6
detector
7
crystal
2
9
Incident X-rays
.
.
Path lengths differences are
not nl because of disorder.
Crystal order (and resolution) improves with
increasing number of lattice contacts
• Potassium channel (1p7b)
• 3.7 Å resolution
• Solvent content=77.7%
• Trypsin (1gdn)
• 0.8 Å resolution
• Solvent content=36.6%
Lattice contacts can form only where
the protein surface is rigid.
By exposing rigid surface area, you enable new
crystal forms previously unachievable.
• Eliminate floppy, mobile
termini (cleave His tags)
• Express individual
domains separately and
crystallize separately, or…
• Add a ligand that bridges
the domains and locks
them together.
• Mutate high entropy
residues (Glu, Lys) to Ala.
or
Crystallization: The task of
coaxing protein molecules
into a crystal.
Is crystallization spontaneous under
biological conditions?
Crystallization
Solubilization
Solvated lysozyme
monomers
Random orientation and position
A lysozyme crystal
Orientation and position of
molecules are locked in a 3D array
High “order”
The barriers to crystallization
•
Unstable
nucleus
Energy penalty
–
–
Lose 3 degrees of freedom in
orientation of protein molecules
Lose 3 degrees of freedom in
translation of protein molecules
•Energy reward
–Some entropy gained by
freeing some surface bound
water molecules.
–Small enthalpic gain from
crystal packing interactions.
DG
1 crystal
(lysozyme)N
N soluble
lysozyme
molecules
•Also, nucleation imposes a
kinetic barrier.
•Unstable because too few
molecules are assembled to
form all lattice contacts.
nM→Mn
The barriers to crystallization
Unstable
nucleus
DG is decreased and the
nucleation barrier lowered by
increasing the monomer
concentration [M].
N soluble
lysozyme
molecules
nM→Mn
DG=DGo+RTln( [Mn]/[M]n )
1 crystal
(lysozyme)N
DG
Lesson: To crystallize a protein,
you need to increase its
concentration to exceed its
solubility (by 3x). Force the
monomer out of solution and
into the crystal. Supersaturate!
nM→Mn
Methods for achieving supersaturation.
1) Maximize concentration
of purified protein
•
•
•
•
•
•
•
Centricon-centrifugal
force
Amicon-pressure
Vacuum dialysis
Dialysis against high
molecular weight PEG
Ion exchange.
Slow! Avoid precipitation.
Co-solvent or low salt to
maintain native state.
We are going to dissolve
lyophilized protein in a
small volume of water.
Concentrate
protein
Methods for achieving supersaturation.
2) Add a precipitating
agent
•
Polyethylene glycol
•
•
•
High salt
concentration
•
•
•
PEG 8000
PEG 4000
(NH4)2SO4
NaH2PO4/Na2HPO4P
olyethylene glycol
PEG
Polymer of ethylene glycol
Small organics
•
•
ethanol
Methylpentanediol
(MPD)
Precipitating agents monopolize
water molecules, driving proteins
to neutralize their surface charges
by interacting with one another.
Systematic vs. Shotgun Screening
• Shotgun- for finding
initial conditions,
samples different
preciptating agents,
pHs, salts.
• Systematic-for
optimizing
crystallization
condtions.
First commercially
Available crystallization
Screening kit.
Hampton Crystal Screen 1
Methods for achieving supersaturation.
Drop =½ protein + ½ reservoir
3) Further dehydrate the
protein solution
•
•
•
•
Hanging drop vapor
diffusion
Sitting drop vapor diffusion
Dialysis
Liquid-liquid interface
diffusion
2M ammonium sulfate
Note: Ammonium sulfate concentration is
2M in reservoir and only 1M in the drop.
With time, water will vaporize from the drop
and condense in the reservoir in order to
balance the salt concentration.—
SUPERSATURATION is achieved!
The details of the method.
Practical Considerations
Linbro or
VDX plate
Begin with reservoirs:
1) pipet req’d amount of
ammonium sulfate to each
well.
2) Pipet req’d Tris buffers, to
each well
3) Same with water.
Then swirl tray gently to mix.
When reservoirs are ready, lay
6 coverslips on the tray lid,
then pipet protein drops on
slips and invert over
reservoir.
Only 6 at a time, or else dry
out.
Proper use of the pipetor.
Which pipetor would you use for
delivering 320 uL of liquid?
P1000
P200
P20
Each pipetor has a different range
of accuracy
P1000
200-1000uL
P200
P20
20-200uL
1-20uL
Which pipetor would you use for
delivering 170 uL of ammonium
sulfate?
P1000
P200
P20
How much volume will this pipetor
deliver?
P200
0
2
7
|
|| ||
How much volume will this pipetor
deliver?
P20
1
7
0
|
|| ||
How much volume will this pipetor
deliver?
P1000
0
2
7
|
|| ||
What is wrong with this picture?
P1000
0
2
7
|
|| ||
50 mL
-
What is wrong with this picture?
P1000
0
2
7
|
|| ||
50 mL
-
Dip tip in stock solution, just under the surface.
P1000
0
2
7
|
|| ||
50 mL
-
Withdrawing and Dispensing Liquid.
3 different positions
Start position
First stop
Second stop
P1000
P1000
P1000
0
2
7
|
|| ||
0
2
7
|
|| ||
0
2
7
|
|| ||
Withdrawing solution: set volume, then push
plunger to first stop to push air out of the tip.
Start position
First stop
Second stop
P1000
0
2
7
|
|| ||
50 mL
-
Dip tip below surface of solution. Then release
plunger gently to withdraw solution
Start position
First stop
Second stop
P1000
0
2
7
|
|| ||
To expel solution, push to second stop.
Start position
First stop
Second stop
P1000
0
2
7
|
|| ||
When dispensing protein, just push to first stop.
Bubbles mean troubles.
Start position
First stop
Second stop
P1000
0
2
7
|
|| ||
Hanging drop vapor diffusion
step two
Pipet 2.5 uL of concentrated
protein (50 mg/mL) onto a
siliconized glass coverslip.
Pipet 2.5 uL of the reservoir
solution onto the protein drop
2M ammonium sulfate
0.1M buffer
BUBBLES MEAN
TROUBLES
Expel to 1st stop,
not 2nd stop!
Hanging drop vapor diffusion
step three
•Invert cover slip over reservoir quickly
& deliberately.
•Don’t hesitate when coverslip on its side or
else drop will roll off cover slip.
•Don’t get fingerprints on coverslip –they
obscure your view of the crystal under the
microscope.
Dissolving Proteinase K powder
• Mix gently
– Pipet up and down
5 times
– Stir with pipet tip
gently
– Excessive mixing
leads to xtal
showers
5.25 mg ProK powder
100 uL water
4 uL of 0.1M
PMSF
• No bubbles
50 mg/mL
ProK
Dissolving Proteinase K powder
• Mix gently
– Pipet up and down
5 times
– Stir with pipet tip
gently
– Excessive mixing
leads to xtal
showers
Remove
50 uL
Add to 5 uL
of 100 mM
PMSF
• No bubbles
50 mg/mL
ProK
55 uL of
50 mg/mL
ProK+PMSF
complex
Proteinase K time lapse
photography
• Covers first 5
hours of
crystal
growth in 20
minute
increments
500 mm
Heavy Atom Gel Shift Assay.
Why?
Why are heavy atoms used to
solve the phase problem?
•
•
•
•
•
Phase problem was first solved in
1960. Kendrew & Perutz soaked
heavy atoms into a hemoglobin
crystal, just as we are doing today.
(isomorphous replacement).
Heavy atoms are useful because
they are electron dense. Bottom of
periodic table.
High electron density is useful
because X-rays are diffracted from
electrons.
When the heavy atom is bound to
discrete sites in a protein crystal (a
derivative), it alters the X-ray
diffraction pattern slightly.
Comparing diffraction patterns from
native and derivative data sets
gives phase information.
Why do heavy atoms have to be
screened?
• To affect the diffraction pattern,
heavy atom binding must be
specific
– Must bind the same site (e.g. Cys
134) on every protein molecule
throughout the crystal.
– Non specific binding does not
help.
• Specific binding often requires
specific side chains (e.g. Cys,
His, Asp, Glu) and geometry.
– It is not possible to determine
whether a heavy atom will bind to
a protein given only its amino acid
composition.
Before 2000, trial & error was the
primary method of heavy atom
screening
• Pick a heavy atom compound
– hundreds to chose from
• Soak a crystal
– Most of the time the heavy atom will
crack the crystal.
– If crystal cracks, try lower
concentration or soak for less time.
– Surviving crystal are sent for data
collection.
• Collect a data set
• Compare diffraction intensities
between native and potential
derivative.
• Enormously wasteful of time and
resources. Crystals are expensive to
make.
How many crystallization plates
does it take to find a decent heavy
atom derivative?
Heavy Atom Gel Shift Assay
• Specific binding affects
mobility in native gel.
• Compare mobility of
protein in presence and
absence of heavy atom.
• Heavy atoms which
produce a gel shift are
good candidates for
crystal soaking
• Collect data on soaked
crystals and compare with
native.
• Assay performed on
soluble protein, not
crystal.
None Hg Au Pt Pb Sm
Procedures
• Just incubate protein with
heavy atom for a minute.
– Pipet 3 uL of protein on
parafilm covered plate.
– Pipet 1 uL of heavy atom
(100 mM) as specified.
– Give plate to me to load on
gel.
• Run on a native gel
• We use PhastSystem
• Reverse Polarity
electrode
• Room BH269 (Yeates
Lab)