272. CELLULAR BIOCHEMISTRY
The Escherichia coli Heat Shock Response:
Six-Week Introduction to Proteomics
Manual Prepared By
Eric S. Eberhardt and Elisa M. Woolridge
From the Department of Chemistry of Vassar College
and the Department of Chemistry & Physics at Marist College
TABLE OF CONTENTS
I. INTRODUCTION
A. PROTEOMICS
B. MOLECULAR CHAPERONES AND THE HEAT SHOCK RESPONSE
C. 2-D ELECTROPHORESIS
D. OVERVIEW OF THE EXPERIMENT
II. EXPERIMENTAL PROCEDURE
A. LAB WEEK 1
i. PRE-LAB TESTING
ii. SCIENTIFIC METHOD AND EXPERIMENTAL DESIGN ESSAY I
iii. PREPARATION OF STARTER CULTURES
iv. HEAT SHOCK CULTURE
v. CELL LYSIS & PROTEIN CONCENTRATION DETERMINATION
vi. FIRST DIMENSION PROTEIN SEPARATION: ISOELECTRIC
FOCUSING
B. LAB WEEK 2
i. SECOND DIMENSION PROTEIN SEPARATION: SDS-PAGE
ii. GEL STAINING
a. FLUORESCENT GEL STAINING WITH SYPRO® RUBY
C. LAB WEEK 3
i. GEL IMAGING
ii. SPOT EXCISION
iii. TRYPSIN DIGESTION
D. LAB WEEK 4
i. MALDI TOF DATA ACQUISTION
E. LAB WEEK 5
i. DATABASE MINING
ii. PROTEIN IDENIFICATION
F. LAB WEEK 6
i. PRESENTATIONS
ii. POST LAB TESTING
iii. LABORATORY WRITE-UP
iv. SCIENTIFIC METHOD AND EXPERIMENTAL DESIGN ESSAY II
III. LITERATURE CITED
IV. APPENDICES
A. APPENDIX 1: GEL IMAGING INSTRUCTIONS
B. APPENDIX 2: MATERIALS & REAGENT PREPARATION
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I. INTRODUCTION
PROTEOMICS
Proteomics is one of the many new important disciplines to emerge from the genome sequencing
projects and the Human Genome Project. Some of these new areas of study include genomics,
which seeks to determine the structure and organization of a genome as well as variations
between species; bioinformatics, which extracts or mines biological information from DNA
sequence information; functional and structural genomics, which shifts the emphasis from
mapping the genomes to determining the biological function of open reading frames or
determination of three-dimensional structures determination of protein. Proteomics builds upon
these other areas by striving to connect physiological processes to biological pathways,
regulatory mechanisms and signaling cascades not only through the identification and
quantification of proteins expressed by a cell, but also by determining the location of proteins,
specific protein-protein interactions and post-translational modifications. Proteomics is derived
from the term proteome and generally describes the systematic study of the PROTEin
complement expressed by a genOME of a cell, or tissue [1-2]. A typical approach for examining
the proteome involves four basic steps: i) separation of proteins by two-dimensional gel
electrophoresis, ii) isolation and enzymatic cleavage of particular proteins of interest, iii) peptide
analysis by either matrix-assisted laser desorption/ionization (MALDI) or electrospray (ESI)
mass spectrometry, and iv) submission of the peptide fingerprint results to DNA or protein
databases that allow for the identification of the isolated protein.
The genome projects have provided a tremendous amount of DNA sequence information
concerning the proteins potentially available to an organism, however many important questions
remain. For example, this sequence information does not tell us which proteins are expressed
and how much of each protein is expressed by the cell. This situation is further complicated by
the fact that protein structure and function cannot be predicted de novo. For a complicated
organism like humans with estimated 30,000-50,000 different genes, our ability to predict
protein structure and function is even more challenging since mechanisms like alternative
splicing of a single gene may afford a dozen different protein products. Further, the genomic
information does not provide insight into the diverse collection of post-translational
modifications (such as phosphorylation or glycosylation) that are involved in the regulatory
mechanisms that modulate cellular processes. In contrast to the information fixed in the genome,
the proteome of the cell is dynamic, with the protein expression profile changing over the matter
of minutes. Consequently, the proteome reflects the particular stage of development or the
current environmental condition the cell or organism finds itself experiencing. Ultimately, the
comparison of healthy and diseased tissues may afford the opportunity to identify the molecular
nature of a disease and provide potential new targets for drug development. The field of
proteomics seeks to answer many of these questions [2].
MOLECULAR CHAPERONES AND THE HEAT SHOCK RESPONSE
The heat shock response is a phenomenon that is common to all living cells and the
Escherichia coli (E.coli) system may be considered a simplified form of the response observed in
other organisms [3]. When bacteria are stressed with elevated temperatures (>42 °C), a set of
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proteins are synthesized in order to maintain the viability of the cell. These proteins are known
as heat shock proteins (Hsp)-many of which are molecular chaperones, but proteases and
regulatory factors are also included [3-7]. The E. coli chaperones are perhaps the best studied
proteins associated with the heat shock response; they include GroEL (Hsp70), GroES (Hsp10),
and DnaK (Hsp60). Their role in response to thermal stress is thought to be related to their
function as molecular chaperones.
Molecular chaperones are family of proteins that display a broad substrate specificity that usually
recognize non-native proteins involved in intermediate stages of folding, degradation, or
membrane translocation [6]. The structure of the GroEL/GroES complex has been solved in the
presence and absence ATP bound and has provided insight into the chaperonin folding cycle in
E. coli. GroEL forms a 14-mer of identical subunits arranged into stacked rings with
hydrophobic binding sites that line the cavity of the complex [8]. In the absence of ATP, the
misfolded or denatured proteins bind the GroEL cavity causing a conformational change that
promotes the binding of ATP and subsequent binding of the co-chaperone GroES. These
binding events cause a large conformational change that serves to seal the chaperone cavity.
While the precise mechanism of the ATP hydrolysis and substrate release remain unclear, a
single amino acid has been suggested in GroEL, Asp398, appears to play a key role in the
ATPase activity and GroES appears to prevent release of the polypeptide until folding is
complete [7]. Also unclear is the precise mechanism by which the protein folding reaction is
facilitated by the GroEL/GroES complex. One suggestion is that by preventing aggregation of
folding intermediates is sufficient to mediate protein folding. Alternatively, a more complicated
mechanism may be involved where protein folding is actively assisted in the cavity of the
GroEL/GroES complex [7].
The heat shock response is characterized by changes at the transcriptional, translational, and
post-translational levels [9-11]. Within seconds of a shift in temperature, transcription of heat
shock genes is increased until the rate of protein synthesis of GroEL and DnaK is increased by
13-fold and 8-fold respectively [10]. Induction of the heat shock genes is modulated by a
positive regulator, the s32 subunit of RNA polymerase [3, 12-13]. The s32 subunit replaces the
s70 subunit during transcription for the core RNA polymerase and under heat-shock conditions
confers specificity to the heat shock promoters. The s32 subunit is observed to have a transient
increase in expression level upon heat shock, which subsequently declines to new steady state
level that is elevated from the initial 30 °C expression level [9]. At 42 °C, a two-fold increase is
observed for these major heat shock proteins; whereas, a 10-fold increase is observed at 46 °C
with the major Hsp’s such as GroEL and DnaK ultimately estimated to account for 20% total
cellular protein [10]. Under heat-shock conditions, a large population of cellular proteins begin
to denature, and then are recognized by the molecular chaperones. Therefore, the increase in
Hsp expression is thought to help the cell survive and manage the large number of denatured or
misfolded cellular proteins that accumulate as a consequence of thermal stress.
2-D ELECTROPHORESIS
Electrophoresis is the movement of charged molecules in an electric field. This technique is an
extraordinarily important method of separating and analyzing biological molecules. The
movement of a molecule in an electric field E is a function of net charge q and the frictional
4
coefficient f. The velocity v at which a molecule moves in an electric field is a function of the
field strength and net charge as defined by the relationship given below.
Eq
n=
f
A molecule’s mobility m is defined as the ratio of velocity to field strength. By extension,
mobility is also equal to the ratio of charge and frictional coefficient as illustrated below:
q
m=n =
E f
The frictional coefficient is a measure of the hydrodynamic size of the molecule wherein the
larger the molecular size of a molecule the larger the frictional coefficient. Molecular shape is
also an important component of the frictional coefficient. Biological molecules like proteins and
polynucleic acids have distinctive shapes. Therefore, the electrophoretic mobility of a biological
macromolecule is a function of its size, shape, and net charge.
In 2-D protein electrophoresis the first dimension separates proteins according to charge, while
the second dimension separates proteins according to size.
The first dimension of electrophoresis, where protein separation is based on charge, is known as
isoelectric focusing (IEF). Proteins are amphoteric molecules, meaning they contain both acidic
and basic functionalities. They carry a positive, negative, or zero net charge depending on the
pH of their environment. The pH at which a protein has a zero net charge is called its isoelectric
point (pI). At pH values above its pI the protein has a negative net charge, while at pH values
below its pI the protein has a positive net charge. IEF takes advantage of this characteristic of
proteins. When placed in an electric field, each protein will migrate within a pH gradient to the
point where the pH equals the protein pI, i.e., where protein net charge is zero. Positively
charged proteins will move toward the cathode while becoming progressively less charged until
they reach their pI. Conversely, negatively charged proteins will move toward the anode until
they reach their pI. If a protein drifts from its pI, then it gains a net charge and will migrate back
to its pI. In this way, proteins are separated in the first dimension.
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) is employed as the
second dimension in 2-D electrophoresis and is accomplished in a vertical gel format.
Polyacrylamide gels are formed by the polymerization of the monomer acrylamide into
polymeric polyacrylamide chains and cross linking with N,N'-methylene-bis-acrylamide. The
polymerization reaction is facilitated by the action of N,N,N',N'-tetramethylethylenediamine
(TEMED) which catalyzes the formation of free radicals from ammonium persulfate. The role
of SDS, an anionic surfactant, is to denature the proteins while coating them to provide a uniform
negative charge per unit length. The addition of a reducing agent such as b-mercaptoethanol
(b M E ) or dithiothreitol (DTT) completes the denaturation by eliminating intramolecular
disulfide bonds. The rate of protein migration, consequently, can be correlated to its mass, i.e., is
independent of the molecule’s native shape or charge.
MALDI-TOF MASS SPECTROSCOPY
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MALDI-TOF mass spectrometry is an important tool used to identify proteins through the
generation of peptide mass fingerprints from protease digestion of a target protein [17-19]. While
trypsin is the most commonly used protease, other enzymes such as chymotrypsin,
endopeptidases Asp-C or Lys C, or even a chemical reagent like cyanogen bromide (CNBr) can
be used to cleave the polypeptide backbone in a sequence specific fashion. When peptides of a
protein are generated by digestion a characteristic profile of masses, or peptide fingerprint, can
be observed when examined by mass spectrometry.
Scheme
This scheme provides a graphic illustration of the MALDI TOF instrumentation for both linear
and reflector operation with the ions fight path indicated by a dashed line. Briefly, samples of
digested protein sample are combined with a matrix and applied to the sample plate, or target,
and allowed to dry. The sample spot (matrix and digested protein co-crystals) are then hit with a
brief pulse of laser light that is absorbed by the matrix. Peptide ions are thought to form in the
gas phase where highly excited, protonated matrix molecules transfer protons to the neutral
peptide molecules. Through the use of a strong electrical field in the source region, ions are
accelerated into the field-free flight tube where the peptides separate on the basis of their
respective kinetic energies. The mass dependent flight times through the flight tube to the
detector are then used for mass determination.
To enhance mass resolution and accuracy, MALDI-TOF instruments typically use delayed
extraction techniques to minimize the distribution of initial kinetic energies after laser excitation
as well as ion mirroring when the instrument is used with a reflector. The mass resolving power
for a linear instrument is in the range of 4000 Da; whereas, a reflector instrument can have a
resolving power up to 10,000 Da.
OVERVIEW OF THE EXPERIMENT
In this laboratory you will examine the heat shock response of E. coli K-12 MG1655 cells. This
is the same strain used in the E. coli genome-sequencing project and is obtained from the E. coli
Genetic Stock Center at Yale University (http://cgsc.biology.yale.edu).
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Your instructor will begin the experiment by establishing 50 mL starter E. coli cultures. You
will be provided a starter culture that is in exponential growth phase. A small aliquot of this
culture will be used to inoculate a large culture (1000 mL) that will ultimately be subjected to the
heat shock. The cells in the large culture will be allowed to achieve exponential growth phase at
30 °C before subjecting them to the shock at 46 °C. You could evaluate the status of cell growth
by simple visual inspection, but for more reproducible results cell density is best determined
using the UV/VIS spectrophotometer. Previous experiments have determined that an E. coli
culture with an optical density between 0.2 and 0.8 at 595 nm is in exponential growth phase.
Culture aliquots will be collected at 0 and 45 min into the heat shock, and the cells harvested via
centrifugation.
To accomplish 2-D electrophoresis of the E. coli proteome, the proteins must first be liberated
from the cells. Further, the proteins must be fully solubilized, disaggregated, denatured, and
reduced. There are multiple methods by which proteins can be extracted from cells. In this
experiment, a solution known as the lysis buffer serves to both extract the proteins and prepare
them for electrophoresis. The lysis buffer contains urea, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS), and tris(hydroxymethyl)aminomethane (Tris).
Urea causes proteins to denature, leaving proteins in a linearized form, a requirement for
isoelectric focusing. CHAPS is a zwitterionic cholamidosulfobetaine detergent that causes
proteins to be solubilized and disaggregated. CHAPS is preferable to other detergents such as
SDS since it is relatively soluble in urea solutions. Tris buffers the solution.
Once the cells have been lysed, UV/VIS spectroscopy will be used to determine the
concentration of protein in each sample. Amino acids absorb light at a frequency of 280 nm,
while nucleic acids absorb light at 260 nm. Due to the abundance of different levels of electrical
energy within the quantized energy levels, the spectrophotometer does not return narrow peaks.
As a consequence, the spectrophotometer returns broad peaks such that peaks at 260 nm and
280 nm interfere. In order to obtain the concentration of protein in a sample, therefore, the
concentration of nucleic acids must be taken into account. This is done using the Warburg
Christian assay:
[Protein] = 1.55A280 - 0.757A260
where protein concentration has units of milligrams protein per milliliter of solution [18]. Thus,
the Warburg Christian assay accounts for error in estimations of protein concentrations due to the
interference of nucleic acids.
Separation of the proteins from the lysed cells will be accomplished via 2-D electrophoresis,
where the first dimension involves IEF and the second dimension SDS-PAGE. In the case of
IEF, the strips must be rehydrated in a denaturing buffer using ampholytes appropriate for the
selected IPG strip, namely 4.7-5.9 in the case of the heat shock analysis. Prior to resolution via
the second dimension, where separation is based on molar mass, the proteins must be treated
with DTT and iodoacetamide to irreversibly reduce and alkylate any protein disulfide bonds.
Following electrophoresis, the proteins will be visualized via staining. There are several possible
choices including SYPRO Ruby, colloidal blue, and silver stains. The choice of a stain is a
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tradeoff between cost, sensitivity, imaging capability, and the need for subsequent protein
excision and analysis. Silver stain provides the best gel images, but solution preparation for this
procedure is extensive, the reagents are expensive, and the proteins cannot be analyzed via mass
spectrometry. In silver staining, proteins are visible as smoky brown bands as a consequence of
the binding of silver atoms to protein sulfhydryl and carboxyl groups. Fluorescent stain using
SYPRO Ruby provides fine imaging, although a UV light is required for visualization. This
stain acts by binding to basic, hydrophobic, and aromatic regions of proteins. While SYPRO
Ruby is easy to use and is compatible with mass spectroscopic protein analysis, this stain is quite
expensive. Colloidal blue is a less expensive alternative. This stain, which acts by binding
indiscriminately to many parts of the protein, provides adequate imaging results and allows for
the extraction of proteins. This dye, however, is less sensitive than either the silver or SYPRO
Ruby stains.
Images of stained gels will be obtained and the results compared to results available at the
SWISS-2DPAGE site. This database is a component of the Expert Protein Analysis System
(ExPASy) proteomics server (http://us.expasy.org) of the Swiss Institute of Bioinformatics.
8
EXPERIMENTAL PROCEDURE
LAB WEEK 1
PREPARATION OF STARTER CULTURES
Three 50 mL starter cultures will be prepared by your instructor via serial dilution of a glycerol
stock of E. coli K-12 MG1655 cells. To accomplish this, a 50 mL media aliquot will first be
inoculated with 200 mL of thawed stock cells. A second 50 mL media aliquot will then be
inoculated with 500 mL of the first culture, amounting to a 100-fold dilution. A third 50 mL
media aliquot will finally be inoculated with 500 mL of the second culture, for an overall dilution
of 1x104 relative to the first culture. All three cultures are incubated overnight at 37 °C. The
purpose of creating serially diluted E. coli cultures is to ensure that at least one culture, most
likely the most dilute, will be in exponential growth phase for inoculation of the large (1 L)
culture for the heat shock experiment. Details of starter culture preparation are provided here:
1. Inoculate the first 50 mL starter culture with 200 mL of the recently thawed E. coli
glycerol stock. Swirl the flask to ensure proper distribution of the cells.
2. Inoculate the second 50 mL starter culture with 0.5 mL of culture taken from just
prepared first starter culture. Swirl the flask.
3. Inoculate the third 50 mL starter culture with 0.5 mL of culture taken from just prepared
second starter culture. Swirl the flask.
4. Allow all three starter cultures to grow overnight at 37 °C with shaking (300-400 rpm).
HEAT SHOCK CULTURE
5. Prepare the spectrophotometer to read at 595 nm. Blank the instrument at this
wavelength using uninoculated media. Disposable cuvettes may be used at this
wavelength.
6. Pipet 1 mL of the starter culture sample into a cuvette and place the cuvette into the
sample chamber of the spectrophotometer. Record the optical density at 595 nm.
Inoculation techniques are key in this portion of the experiment. Every time the
cap or foil covering is removed from a culture flask, the opening of the flask must
be flamed. Similarly, each time foil is replaced, the flask should again be flamed.
Avoid laying the flask cap on any surface as this increases the likelihood of
contamination.
7. Identify a starter culture that possesses an optical density at 595 nm (OD595 ) between 0.2
and 0.8. Asceptically inoculate 1 L of media in a 3 or 4 L Fernbalch flask with 5 mL of
the chosen starter culture.
8. Grow the new culture at 30 °C with shaking at 300-400 rpm until the OD595 reaches
0.5-0.8. Harvest a 50 mL aliquot for centrifugation and distribute the aliquot between
two 50 mL centrifugation tubes. Return the culture to the incubator.
9
9. Increase the temperature of the incubator to 46 °C. Once the temperature hits target,
begin timing the heat shock. After 45 minutes, remove a second 50 mL aliquot for
centrifugation.
10. Centrifuge counter-balanced harvested cell samples at 16,000xg (10,000 rpm in a GSA
rotor or 12,000 rpm in an SS-34 rotor) for 10 min at 4 °C. Decant the supernatant. The
E. coli cells are contained in the pellets. If the cells will not be lysed on the day of
harvest, store the pellets at -80 °C.
Destroy any remaining E. coli cultures with bleach before disposal.
CELL LYSIS & PROTEIN CONCENTRATION DETERMINATION
11. Carefully wash the pellets three times with 500 mL of 40 mM Tris-Cl, pH 8.8. Discard
the washes.
When performing the washes, be sure not to disturb the pellet. The easiest way to
do this is to tilt the centrifuge tube so that the pellet is positioned upside-down.
Pipet the wash into the tube then turn the tube so that the wash runs over the
pellet removing the residual salts. The removal of these salts is important for the
isoelectric focusing process. If the pellet becomes dislodged simply recentrifuge.
12. Resuspend the cell pellets in 250 mL of cell lysis buffer (8 M urea, 4% w/v CHAPS, and
40 mM Tris). This is accomplished by repeated flushing of the same 250 mL lysis buffer
over the pellet. Be sure to fully resuspend the pellets. Vortex mixers may be used to aid
the resuspension.
Avoid resuspending the cell pellets in warm lysis buffer as this can cause
hydrolysis of urea to cyanate, and cyanate can covalently modify protein. This
has the undesired consequence of modifying protein pI values.
13. Place the cell suspensions on ice.
14. Dilute 10 mL of the cell suspension to 1000 mL with 990 mL lysis buffer. Measure the
absorbance of the diluted, lysed cells at 260 nm and 280 nm, using lysis buffer as a blank.
In order to ensure accurate results, the absorbance values at both 260 nm and
280 nm must fall between 0.1 and 1.0. If this is not the case for your measured
sample, adjust the dilution accordingly.
15. Use the following equation to estimate protein concentration, expressed as mg/mL. Be
sure to account for the dilution, e.g., 100-fold in the case of dilution of 10 mL lysed cells
to 1000 mL.
[Protein] = 1.55(A280) - 0.757(A260)
16. Determine the volume of sample required to obtain the mass of protein appropriate for
the desired stain. Gels treated with silver, fluorescent, and colloidal blue stains
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respectively require total protein amounts of approximately 0.25 mg, 0.50 mg, and
1.25 mg.
FIRST DIMENSION PROTEIN SEPARATION: ISOELECTRIC FOCUSING (IEF)
After the sample protein concentration has been determined, the volume of sample to be used
per well on the IEF must be calculated. This volume will depend on three things: the sample
concentration, the strip pH gradient range, and the stain type. Different stains require
different amounts of protein. The following tables give approximations for various stains
and strips:
Type of Stain
Amount of Protein
Silver Stain
0.25 mg
Fluorescent
0.50 mg
Colloidal Blue
1.25 mg
pH Gradient Range
3-10
4-7
3-6
5-8
7-10
Protein for 11cm Strip
20-200 mg
40-200 mg
40-200 mg
40-200 mg
50-300 mg
pH Gradient Range
3.9-5.1
4.7-5.9
5.5-6.7
6.3-8.3
Protein for 11cm Strip
50-250 mg
50-125 mg
50-250 mg
125-250 mg
Refer to your instructor for the exact amount of protein to use.
17. Prepare the sample for IEF by mixing with rehydration buffer (8 M urea, 2% w/v
CHAPS, 0.2% BioLytes*, 0.001% w/v bromophenol blue, and 50 mM DTT added just
before use). Each IEF well can hold 185 mL solution, and at least 60 mL of this volume
should be rehydration buffer. Subtract the calculated sample volume from 185 mL. The
difference is the volume of rehydration buffer that should be used. If the calculated
sample volume exceeds 125 mL, consult your instructor. Mix the sample and rehydration
buffer in sterile 1.5 mL centrifuge tubes.
Gloves should be used at all times while handling ReadyStrip IPG strips, the IEF
apparatus, and any solutions used in the IEF process to prevent protein
contamination, notably skin keratin.
*The following table highlights which carrier ampholyte buffer should be used with each
type of strip:
ReadyStrip
3-10
4-7
3-6
5-8
Buffer
Bio-Lyte 3-10
Bio-Lyte 3-10
Bio-Lyte 3-10
Bio-Lyte 3-10
ReadyStrip
3.9-5.1
4.7-5.9
5.5-6.7
6.3-8.3
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Buffer
ReadyStrip 3.9-5.1 Buffer
ReadyStrip 4.7-5.9 Buffer
ReadyStrip 5.5-6.7 Buffer
ReadyStrip 6.3-8.3 Buffer
7-10
ReadyStrip 7-10 Buffer
18. Obtain an 11 cm IEF focusing tray. Pipet 185 mL of each sample as a line along the back
edge of separate wells (i.e., one sample per well). The line of sample should extend the
whole length of the well except for about 1 cm at each end. Be careful not to introduce
any bubbles which could interfere with even distribution of the sample within the strip.
19. Carefully remove the cover sheet from the ReadyStrip IPG strip using two sets of
tweezers.
20. Gently place the strip gel side down onto the sample. The “+” and the pH range should
be legible, and the “+” should be aligned with the embossed “+” on the focusing tray.
Any sample on the plastic backing of the gel will not be absorbed, so be careful to keep
the sample under the strip. Also, take care not to trap air bubbles beneath the strip. If
this happens, use forceps to lift the strip up and down until the air bubbles move to the
end and out from under the strip.
21. Cover each strip with 1.5 to 2.0 mL mineral oil to prevent sample evaporation during the
rehydration process. Add the mineral oil slowly so as not to disturb the sample.
22. Cover the tray with the plastic lid, and rehydrate under active conditions. The IEF
rehydration protocol follows:
•
•
•
•
•
•
•
•
•
Select “Preset Method”
Select “Linear”
Program window:
Rehydration
Yes
Gel length
11 cm
Focus Temperature 20 C
Program window:
Rehydration step
Active @ 50V
Enter Temperature
20 C
Program window:
Enter Time
12:00
Insert pause after rehydration Yes
Program window:
S1 250 V
00:15
S2 Select: hrs:min
Vhours
S2 8000
3:30
Program window:
S3 hrs:min
Select: vhours
S3 8000 V
35000
S4 500V/hold
Select: yes
Program window:
Method:
Linear dV
Limit/Gel
50 Ua
Enter # of Gel
???
Press the green man to start
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23. After rehydration is complete, saturate two wicks per well used with deionized distilled
water. Blot excess water on filter paper. Carefully lift the ends of each strip with forceps
and insert a wet wick between the strip and the electrodes. These wicks act to remove
any residual salts in the sample prior to the electrophoresis.
24. Press the green man to start the electrophoresis run.
25. At the end of the run, remove the mineral oil from the IPG strips. Using forceps, transfer
the strips to a disposable 17 cm rehydration/equilibration tray. Place one strip per
channel, gel side up.
26. Store the strips in the covered tray at –20 °C until the next lab period.
LAB WEEK 2
SECOND DIMENSION PROTEIN SEPARATION: SDS-PAGE
The following directions are written for use with 11 cm IPG Strips and BIO-RAD’s Criterion
Precast Gels. Mini format gels may be easily substituted for the mid-size Criterion gels, but then
7 cm IPG strips must be used for the first dimension of electrophoresis.
27. Bring the strips that had been stored at –20 °C to room temperature.
28. Add 6 mL Equilibration Buffer I to each channel. Place the covered tray on an orbital
shaker and gently shake for 10 min.
Equilibration Buffer I contains 6 M urea, 20% glycerol, 0.375 M Tris-HCl, pH 7.8,
2% w/v SDS, and 130 mM DTT. Equilibration Buffer II contains 6 M urea,
20% w/v glycerol, 1.5 M Tris-HCl, 2% w/v SDS, and 135 mM iodoacetamide. Both
buffers should be prepared just before use (e.g., make Equilibration Buffer II while the
strips are incubating with Equilibration Buffer I).
29. At the end of the 10 min incubation, decant the buffer from the tray.
30. Add 6 mL Equilibration Buffer II to each channel. Place the covered tray on an orbital
shaker and gently shake for 10 min.
31. At the end of the 10 min incubtion, decant the buffer from the tray.
32. For each strip to be analyzed, remove the appropriate Criterion Precast Gel (e.g., 4-12%
Bis-Tris, 1 mm) from its container. Pull out and discard the green comb. Peel off and
discard the sticker from along the bottom of the gel. Insert the gel into the
electrophoresis cell so that the large well area faces the inside of the box.
33. Transfer the strips to the vertical gel so that the plus (+) sign on the strip is next to the
molecular weight standards lane outlined on the outside of the gel. Recall that during an
IEF run, the gel strips are oriented with the plastic backing facing up, i.e., gel side facing
down. For the second dimension of electrophoresis, the gel side of the strip should be
oriented toward the outside of the electrophoresis unit. The strip will fit in the slot
created by the two plastic plates containing the gel. Gently press the strip down to
remove bubbles, and so that there is good contact between the strip and the gel.
Using forceps, handle only the ends of the strips where no gel is present.
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34. Add 10 mL of boiled broad range molecular weight standards to the single lane adjacent
to the IPG strip.
While not vital to the experiment, the use of molecular weight standards will
allow for visual conformation that the gels are running properly.
35. Prepare 500 mL per strip of 1X XT MOPS running buffer from the 20X XT MOPS
concentrate. A Criterion electrophoresis unit, running two gels, uses approximately 800900 mL of running buffer.
36. Add 1X running buffer to the fill line found on the side of the electrophoresis cell and to
the well on the gel so that the slot with the strip is completely covered. Firmly place the
cover onto the box so that the red wire on the cover connects to the red electrode on the
box. Plug the wire into the power supply so that red is again aligned. Turn on the power
and run the gels at 200 V for 55 minutes.
37. At the completion of the run, turn off the power and remove the gel from the
electrophoresis unit. To separate the gel from the plates for staining, turn the gel
sandwich upside down and press the sandwich down over the protruding part of the
electrophoresis box top until a cracking sound is heard. Carefully pull apart the two
plastic plates and discard the side without the gel. Orient the remaining plate gel-side
down over a container that will be used for staining and storing. Using the blunt end of
the forceps, tease the edge of the gel gently off the plate into the container. Allow gravity
to do most of the work. Each container should contain only one gel.
FLUORESCENT GEL STAINING WITH SYPRO® RUBY
38. Pour enough of a fixative solution containing 7% v/v glacial acetic acid and
10% v/v methanol over each gel so that the gels are covered and moves freely [19].
Agitate the gels on an orbital shaker at 50 rpm for 30 minutes. Decant the fixative into a
waste container for methanol disposal.
39. Add enough SYPRO® Ruby Protein Gel Stain to allow the gels to float freely. Stain the
gels on an orbital shaker at 50 rpm for a minimum of 90 minutes, or preferably overnight.
Do not use glass containers for this stain, and avoid touching the gels, even with
gloved hands.
40. Transfer the gels into the fixative solution for storage until visualization. Dispose of the
decanted SYPRO® Ruby stain as hazardous waste.
If the gels, when imaged under UV light, have high background intensity or
appear speckled, decant the staining solution and replace it with enough fixative
solution to approximately equal the amount of staining solution removed. Place
this on the orbital shaker for 30 minutes at 50 rpm.
14
ALTERNATIVE GEL STAIN: COLLOIDAL BLUE
A. Pour enough of a fixative solution containing 12% w/v trichloroacetic acid (TCA) and
3.5% w/v sulfosalicylic acid over each gel so that the gels are covered and move freely.
Agitate the gels on an orbital shaker at 50 rpm for 60 minutes. Decant the fixative and
dispose of as hazardous waste.
B. Add enough Colloidal Blue Stain to allow the gels to float freely. Stain the gels on an
orbital shaker at 50 rpm for 30 minutes.
Do not use glass containers for this stain, and avoid touching the gels, even with
gloved hands.
C. Decant the colloidal blue solution and replace it with 200 mL of ddH2O per gel and shake
for 7 hours to 3 days. Dispose of the decanted stain as methanol waste. If the gels must
be stored for longer then 3 days, then place them in 20% w/v ammonium sulfate at 4 °C.
LAB WEEK 3
GEL IMAGING
41. Once the protein gels are created and stained, use an imager to digitize the gel images.
The PDQuest software can be used to measure protein spot intensity, as well as to
analyze differences in protein expression levels between two or more gels. MALDI-TOF
mass spectrometry analysis can aid identification and calibration based on molecular
weights and pI values. Alternatively, although less accurate, you can attempt to identify
proteins based on comparison to gels in the SWISS-2DPAGE database
(http://us.expasy.org/ch2d). Detailed instructions for use of the PDQuest software are
provided in Appendix 1.
LAB WEEK 4
PROTEIN EXCISION & TRYPTIC DIGESTION
For all remaining steps, you must be extremely careful to avoid inadvertent
contamination. Always wear powder-free gloves and use sterile microfuge tubes
and pipet tips when handling samples.
1. Using a razor blade or glass pipette, and working over a UV light box for fluorescent
SYPRO® RUBY stained gels, make small incisions into the gel to remove the section(s)
15
containing the suspected heat shock protein(s). Place each gel slice into a separate,
labeled 500 _L sterile Eppendorf tube.
To minimize UV exposure, wear UV protective safety glasses, a lab coat or shirt
with long sleeves, and, for extended work over the box when the gel box shield
must be removed, a UV protective face shield.
2. Soak the excised gel pieces in 400 mL in 50% v/v acetonitrile in 12.5 mM NH4HCO3, pH
= 7.8, for 15 min. Remove the wash, being careful not to remove the gel piece. Repeat
this wash two more times. The purpose of this is to remove the stain that had been used
for imaging.
3. Soak gels in 100% acetonitrile for 5 minutes. The gels will turn opaque white. Remove
the acetonitrile.
4. Dry the gels in a centrifugal concentrator for 20-30 minutes. The gel pieces will reduce
in size and become transparent.
Note: Speed Vaccuum Operation See instructor before use.
As appropriate for your system, either turn on the refrigerated coil or surround a
cold finger with a dry ice and isopropanol bath. While the vacuum pump is still
off, make sure that the vacuum line is open to the atmosphere. Place the gelcontaining microfuge tubes in the rotor with the tube caps open and facing out;
this will allow for easier location of the precipitate at the end of the run. Next,
close the concentrator lid, turn the rotor on, and, only once the rotor is at full
speed, turn on the vacuum. Close off the atmosphere to the system and turn on
the heat to the rotor. After the designated run time, shut down the concentrator
by first opening the system to the atmosphere, followed by turning off the vacuum
pump. Finally, turn off the heat and stop the rotor. These steps must be
performed in the sequence described in order to avoid loss of sample. If the tubes
were open and vacuum applied in the absence of a centrifugal force, the sample
could be easily lost to the vacuum.
16
5. Rehydrate dried gels with approximately 10-15 mL of trypsin solution. The gels will
swell and turn clear. Check the reactions after 30 min to make sure there is sufficient
volume to completely wet the entire gel; add additional trypsin if needed. Incubate the
tightly capped reactions at 37 °C for 16-24 hr.
6. Soak the trypsin-treated gels in 25-50 mL of a solution containing 50% v/v acetonitrile
and 5% v/v trifluoroacetic acid (TFA) for 30-60 min with gentle agitation on an orbital
shaker (do not vortex). Transfer the supernatant to a clean, labeled tube and extract the
gels with a second 25-50 mL aliquot of the acetonitrile/TFA solution for 30-60 min.
Combine the two extracts and then take the extracted peptides to complete dryness in a
centrifugal concentrator; this will take approximately 1 hr.
If the MALDI-TOF plate will not be spotted in the next few hours, store these
samples indefinitely at -20 °C.
MATRIX AND CALIBRANT PREPARATION
1. To prepare the matrix, measure 15 mg of a-cyano-4-hydroxycinnamic acid (CHA) into a
tared microfuge tube. Add 499 mL ddH2O, 1 mL TFA, and 500 mL acetonitrile to the
CHA. Be sure to add TFA to the ddH2O to avoid loss of the volatile TFA. Vortex the
saturated matrix solution well and then touch spin to settle excess CHA.
2. For each calibrant to be used, add 1 mL calibrant standard to 24 mL of the matrix solution
in a 0.5 mL microfuge tube. Vortex well and then touch spin.
The matrix and calibrants should be prepared just before spotting the MALDI
plate.
17
SAMPLE CLEAN-UP AND MALDI PLATE SPOTTING
1. For each trypsin-digested protein sample, wash one ZipTipC18 (hereafter referred to as
ZipTip) with 10 mL 100% acetonitrile followed by 2 to 3 washes of 10 mL 0.1% TFA;
discard these washes.
2. Reconstitute the peptide samples with 10 mL 0.1% TFA (containing no acetonitrile).
Draw the sample into the ZipTip and pipette up and down 5 to 6 times to ensure peptide
binding; finally expel and discard the liquid. The peptides should be bound to the
ZipTip.
3. Wash each ZipTip 3 times with 10 mL aliquots of 0.1% TFA.
Note that to achieve a volume of 10 mL for the ZipTip the pipetman must be set to
13 mL. Similarly, a volume of 3 mL requires a setting of 7.5 mL. Also, allow
sufficient time for the liquid to be completely drawn into the ZipTip.
4. Clean a MALDI spot plate with soap and water using a soft toothbrush. Complete a log
sheet to guide sample placement on the MALDI plate.
5. Set aside 3 cm squares of Parafilm, paper contact side up. Draw 3 mL of the CHA matrix
solution into the ZipTip and deliver this solution to a clean Parafilm square. Using a
2 mL pipetman, transfer 1 mL of the sample to the predetermined location on the MALDI
plate.
6. Once all protein samples have been spotted, place 1 mL of a diluted calibrant in the upper
left hand corner of each grid that contains sample.
7. Allow the plate to dry in the open air for at least 1.5 hours and then store under vacuum
in the MALDI-TOF mass spectrometer until analysis.
Once dry, spotted sample plates must be transferred to the spectrometer. Samples
are stable for one week under the instrument vacuum but only for a few hours in
the open atmosphere.
18
MALDI-TOF MASS SPECTROMETRY
1. Instrument Set Up
a. Turn on the Voyager-DE Pro monitor. Open the Voyager Control Panel on the
desktop and click the ‘hand’ followed by ‘eject plate’. Slide the plate in from the
side ensuring it is aligned correctly and is properly secured with the loading arm.
Click the ‘hand’ sign again, scroll down to plate one and load. Click the
lightening bolt to turn on the high voltage and allow the instrument to warm up
for 30 minutes. Open the instrument settings and select 500 - 5 K reflector mode.
2. Calibration
a. Ensure the selected calibrant matrix is _-cyano-4-hydroxycinnamic acid. Toggle
laser joystick to aim at calibrant 2 spot below the sample being analyzed.
Increase the shots per spectrum to 100. Maintain laser intensity between 1800 to
about 2500 intensity units. Scroll along the sample to find good crystal sites.
Toggle the joystick slightly after about 50 shots to avoid adding noise without
more ions. Save spectrum in the proper drive with an appropriate name and click
on the burning disk. Open ‘data explorer’, select ‘open file’, and click on the
spectrum. Select ‘add’, then ‘finish’.
b. To process the spectrum click ‘Process’ on the tool bar followed by ‘Advanced
Baseline Correction.’ Again click ‘Process’ followed by ‘Noise Filter/Smooth’
and ‘OK.’ Click ‘Tools,’ ‘Resolution Calculator,’ right click, hold and drag the
box across a C12 peak of a well-resolved ion near the middle of the spectrum. A
number will appear at the bottom of the screen. Open ‘Peaks’ and ‘Peak
Detection.’ Write the previous number in the box at the bottom and click
‘Apply.’ Next, go back to ‘Process’ and ‘Noise Filter/Smooth.’ Open the box
and click on ‘Default’ and then ‘OK.’ Finally, open ‘Process,’ ‘Mass
Calibration,’ and ‘Manual Calibration.’ Right click and drag across the Carbon
12 peak for each signal. If all peaks are correct and resolved click ‘Plot,’ ‘Apply
Calibration,’ and ‘Export.’ Minimize ‘data explorer’ and at the bottom right of
the screen click the ‘External Calibration’ button and select the recently save
19
calibrant. If all peaks are not correctly resolved it may be necessary to take
another spectrum of the calibrant, or try a different calibrant.
3. Sample
a. To process a spectrum for a sample follow the above instructions up until the
‘Manual Calibration’ step. Instead, click ‘Peaks,’ ‘Peak Detection’ and change
‘% Base Peak Intensity’ to a level above the highest noise peak and below the
lowest identifiable signal peak, and click ‘Apply.’ Next, open ‘Peaks’ and click
‘Peak Deisotoping.’ This will provide only those peaks that are identified as a
signal and will be used to determine which proteins were present from the
database. To export this spectrum open the ‘File’ menu, click ‘Export’ and
‘ASCII Spectrum.’ Save the spectrum to a 3_ floppy disk.
LAB WEEK 5
Database Mining
I.
Database Search
1. Open up Internet Explorer and go to the protein prospector site (http://prospector/).
Select M-S Fit program and choose SwissProt.r36. Set the maximum number of
cleavages missed to 1 and the mass tolerance to +/- 100 ppm. Next set the minimum
number of matches to 4, or 3 at the least. Last, copy the m/z values from the
WordPad file of the ASCII spectrum previously saved. Delete the second list of
numbers, which represent peak resolution, and only paste the first list. Also, it may
be necessary to go through the list of data and remove any values below 800 or any
that are 1-2 Da away from each other. Finally, click ‘Start Search’ and wait for the
list of proteins to appear for analysis and interpretation.
2. Removing values, which are 1-2 Da away from each other, optimizes the MOWSE
(Molecular Weight Search) score obtained when analyzes of the data has finished.
The MOWSE score is based on the likelihood of a peptide to be contained in a certain
group of molecular weights (http://prospector.ucsf.edu). MOWSE scores of
approximately 105 are considered a “hit” when identifying peptides.
20
LAB WEEK 6
Post-Laboratory Testing
Laboratory Presentations-10-15 min in class
ACKNOWLEDGMENTS
We thank the NSF CCLI Program (Award # 0230794), the NSF MRI Program
(Award # 0079393), and the Howard Hughes Medical Institute University Award Program for
funding that supported this project.
LITERATURE CITED
[1]
[2]
[3]
[4]
[5]
[6]
[7]
[8]
[9]
[10]
[11]
[12]
[13]
[14]
[15]
M. R. Wilkins (1997) Proteome Research: New Frontiers in Functional Genomics,
Springer-Verlag, New York.
R. M. Kemp, D. Kyriakidis, T. Choli-Papadopoulou, Eds. (1999) Proteome and Protein
Analysis, Springer, New York.
B. Bukau (1993) Regulation of the Escherichia coli Heat-Shock Respose, Mol.
Microbiol. 9, 671-680.
G. Segal, E. Z. Ron (1998) Regulation of heat-shock response in bacteria, Ann. N.Y.
Acad. Sci. 147-150.
F. U. Hartl (1996) Molecular chaperones in cellular protein folding, Nature 381, 571-580.
A. Richardson, S. J. Landry, C. Georgopoulos (1998) The ins and outs of a molecular
chaperone machine, Trends in Biochem. Sci., 23, 138-143.
H. R. Saibil, N. A. Ranson. (2002) The chaperonin folding machine, Trends in Biochem.
Sci., 27, 627-632.
Z. H. Xu, A. L. Horwich, P. B. Sigler The crystal structure of the asymmetric GroELGroES-(ADP)(7) chaperonin complex, Nature, 388, 741-750.
P. G. Lemaux, S. L. Herendeen, P. L. Bloch, F. C. Neidhardt (1978) Transient rates of
synthesis of individual polypeptides in E. coli following temperature shifts, Cell, 13,
427-434.
S. L. Herendeen, R. A. VanBogelen, F. C. Neidhardt (1979) Levels of major proteins of
Escherichia coli during growth at different temperatures, J. Bacteriol. 139, 185-194.
M. Y. Sherman, A. L. Goldberg (1992) Heat shock in Escherichia coli alters the
protein-binding properties of the chaperonin groEL by inducing its phosphorylation,
Nature, 357, 167-169.
D. B. Straus, W. A.Walter, C. A. Gross (1987) The heat shock response of E. coli is
regulated by changes in the concentration of sigma 32, Nature, 329, 348-351.
D. B. Straus, W. A.Walter, C. A. Gross (1990) DnaK, DnaJ, and GrpE heat shock
proteins negatively regulate heat shock gene expression by controlling the synthesis and
stability of sigma 32, Genes Dev. 4, 2202-2209.
Methods Enzymol., 1957
R. J. Simpson (2003) Proteins and Proteomics: A Laboratory Manual. Cold Spring
Harbor Laboratory Press, Cold Spring, New York.
21
APPENDIX 1: GEL IMAGING INSTRUCTIONS
Double click the PDQuest icon to open the software. The program’s layout has a menu bar on
top with the standard “File”, “Edit” and other commands. Below this is the toolbar. The buttons
on it are used throughout in manipulation of the gel and its data. To the right of the tool bar are
two small windows. The first displays which tool is currently selected for use. To unselect a
tool, click its name in this window or select a different tool. Attached to this window is a quick
help window. When the cursor is held over a tool, a quick description of that tool’s function is
displayed in this window. The names of the buttons referred to in these directions originate from
holding the cursor over the icon and reading the small pop-up label.
To start, open the door to the MultiImager and place the gel face up on the clear plate inside. It
is a good idea to clean it with some ddH2O and kimwipes before and after use to avoid
interference from residual matter. If the gel is to be imaged is in the visible region of the
electromagnetic spectrum, press the position button, which will create a live image of the gel and
a crosshair center to aid in positioning the gel. If the gel requires UV light, the preview button
may be tried, but it will likely require using the acquire button and transforming the image to get
the gel centered. Once the gel is centered, close the imager’s door, press the Acquire button,
and give the gel a descriptive title (i.e. heatshock30min) in the pop-up window. If the image
comes out bleached white or too dark, adjust the exposure time and reacquire the image. In
general, an exposure time of two seconds works well for gels imaged in the visible light region
while a time of five seconds is better for UV imaging. Also, make sure that 2D Scan is selected
under Type in the pop-up. Images may be retrieved later using File->Open and selecting the gel
from the appropriate folder.
The next step is transforms the image to maximize spot resolution. This process does not
permanently change the file and is therefore reversible. Under Image in the menus bar, select
Transform. Use the Autoscale button to quickly optimize the gel, and further fine tune this by
moving the sliders to maximize spot resolution and minimize background noise. Next, open the
Image Tools bar by clicking its icon in the tools bar. Select the crop tool and drag a box around
the area to keep. It is recommended that the very outside of the gel be cut of as the usually bright
edges often show up as spots. Move the cursor inside the box when satisfied (the cursor will turn
into a pair of scissors) and click. Other options such as flipping and rotating can be found in the
Image Tools bar. Repeat this process on all gels. The last gel should not be left on the plate for
a long period of time as this might cause permanent stains. Wash the plate with ddH2O or
ethanol when finished.
With the proper settings, the computer will attempt to find all of the spots on the gel image.
Under Spots in the menus bar, select Spot Detection Wizard. A window will pop up and
prompt you to select three types of spots. With these spots will determine what areas of the gel
contain spots and what is just background. Select the Zoom tool from the tool bar and zoom in
on a region of the gel containing a faint spot. Click the Select Faint Spot button, located on the
right side of the window in under “Step 1,” and click the center of the faint spot on the zoomed
image of the gel. Use the Full Image button to restore the image. The next step asks for
selecting a small spot. If the faint spot is the smallest spot then move on to the large spot
selection, else repeat the above procedure using the Select Small Spot button. For selecting a
22
large spot, click the corresponding button and drag a box around the largest spot on the gel.
Next, using the Select Background button, drag a box around a large area of characteristic
background that has no spots. If the gel has a large amount of streaking, select the appropriate
options, on the bottom right under “step 3,” for reducing the desired type of streaks. The streak
radius setting should not be changed. Click Find Crosshair Centers under “step 2.” Yellow
crosshairs will appear over any areas the computer considers a spot. If spots were missed, try
resetting the faint, small, and large spots. When satisfied, click Process All Gels and provide a
descriptive name for saving. A pop-up box appears after this in which all of the gels that have
been stained, run, and scanned under the same conditions should be selected by clicking their
names. The parameters just saved should be selected, so press the OK button. This will process
all of the gels for spots. Another pop-up window prompts for creating a match set, which can be
done immediately or done later using the Create Match Set command. Again, enter a
descriptive title and use select or load to add gels to be compared. Click on the gels name in the
sub-window to remove it. Click Create and select a gel for making the match set Standard. The
gel selected should be of high quality and be characteristic of the experiment (i.e. a control gel).
This creates a match set with the Standard in the top, left most window. The Standard is an
image made from the Gaussian representation of spots from other gels in the matchset. The rest
of the images are either Matchset Members or Member Images. Matchset Members are the
Gaussian representations of the Member Images, which contain the original image. “Shift +
F11” can be used to switch between the two types of images. The Matchset Members are
smoother and easier to work with, but one should refer back to the Member Images to make sure
the Gaussian models are accurate. Since the Standard is an artificial image, it does not have a
corresponding Member Image. Use the Configure Subwindows button on the tools bar to view
one image at a time by clicking the picture of a single window. To change the image displayed,
click the Assign button in the tools bar, click the image, and select the desired gel from the popup window.
Use the Landmarking tool to account for distortions in the gels when matching up spots. The
landmark tool can be found in the Match Tools bar, which in turn is found on the tool bar. Spots
used for landmarking should be distinct and present in all gels. With the landmark tool active,
click a spot in the Standard. The cursor will automatically move to the next gel in a region
where it thinks the spot should be. Click the actual spot in this image and repeat for all of the
gels. If a spot is missing in one gel, simply click on the correct spot in the next gel, although this
should be avoided if possible. Continue to landmark several spots in various regions of the gel.
Green letters indicate spots that have been matched. Red circles in the Standard indicate spots
that only have matches in the template from which the Standard was constructed. Red circles on
the gel images represent spots not included in the Standard. Additional landmarks should
decrease the number of unmatched spots. To determine if landmarks are being made correctly,
select the Hide Overlays button in the tool bar and click on each image to remove excess
symbols. Now select the Show Offsets button in the Match Tools Bar and click the gels. Small
lines will appear. They should be roughly parallel, of the same length, and hey should not cross.
If this is not the case, make sure all spots have been landmarked correctly. Use the Show
Symbols button from the Match Tools bar to restore the standard symbols. Try to remove more
of the red circles, using the Member Images, with landmarks. Retransforming the gels might
help to clarify very faint spots. If they are retransformed, rematch the gels using the Match All
Gels button in the Match Tools Bar. To find mismatched spots, not just spots that may be
23
different between control and experimental results, got to Match -> Edit Matches -> Mark
Erratic Response in the menus bar and click the Standard. The fold factor in the pop-up
window will allow this function to mark spots with a yellow circle that vary more than the factor.
Rework the landmarks to avoid such spots.
The computer may misrepresent spots that are slightly merged or very large and dark. To correct
this, zoom on the trouble region and select the Combine button from the Edit Spots tool bar.
Drag a box around what should be a single spot and click inside the box. This will place a single
crosshair at the user-defined spot. Make sure to use the Member Images for this step for more
accurate results. When finished, use the Match All Gels button. Purple squares should appear
around the spots manually matched.
This step will attempt to counteract variations due to differences in preparation. This is called
normalization and will be done by creating an analysis set. Additional analysis sets will be made
latter so it is important to distinguish between them when giving the saved parameters a title.
Open the Analysis/Annotation Tools bar and select the Create Analysis Set button. Name the
set and select Arbitrary for the “Type.” Select “No Spots” and click “Go” to create the set. Add
spots to the set with the Add Spot to Set button from the Analysis/Annotation Tools bar. Use
spots that are well resolved, do not overlap, and are present in most gels. Add several spots then
go to Analysis -> Normalize in the menus bar. In the pop-up, click “Enable Normalization” and
use “Total of Spots in Analysis Set.” If several good spots were not found for the set, use the
“Total of All Valid Spots” option. Select the desired scaling and hit OK.
The following steps require molecular weight and pI data obtained from MALDI-TOF analysis
or some other means.
Setting MrpI standards allows for the calculation of molecular weight and pI of unknown spots
and further statistical analysis. Use the Configure Subwindows button to display the Standard.
From the MrpI Tools bar, select Enter MrpI Data and click on known spots. In the pop-up
window, enter the molecular weight, in kDa, followed by a space and then the pI data. If one of
the two parameters is unknown, type zero. Repeat this for several spots, more spots will yield
more accuracy, and press the Plot MrpI button. Select spots on the edges for increasingly
accurate pI information. Go to Spots -> Spots Parameter to display all MrpI data.
Analysis sets allow for further study of differences in spots through qualitative, quantitative,
statistical, and Boolean means.
Quantitative Analysis: Use the Configure Subwindows option to view the Standard. Select the
Create Analysis Set button form the Analysis/Annotations Tools bar. Select “Quantitative”
under Type and select gels to be A and B by pressing the corresponding buttons and choosing a
gel from the pop-up list. Next, select Increased, Decreased, or Increased or Decreased and set
the value in the window. This will show spots that have, for example, increased in set B by
greater than a factor of 3 over the corresponding spot in A. Highlighted spots appear in the
Standard’s window. To view the difference in spot expression with a histogram, select the
Margin Graph button and click any spot to display a pop-up graph of the difference in
expression. To remove graphs, use the Hide Overlays tool in the tool bar. The data in the
24
histogram is a comparison of spot quantities. Based on the previous settings, the computer
assigns a number to each spot that tells how much protein is present based on the pixels. With
the cursor over a gel image, press Alt + F6 to see all the quantities in that gel.
Qualitative Analysis: Used for detecting spots found in one gel (i.e. the experimental gel) and
not in another (i.e. the control), This type of analysis can be used by creating a new analysis set
and selecting “Qualitative” from the Type option. Reselect gels for study and select whether
spots are found in B and not A, On in B, or vice versa, On in A. The value for “Fold Over
Minimum Sensitivity” setting will allow the program to highlight spots that are the set number
times greater then on the gel lacking the spot.
Statistical Tests: If several gels were made from the same sample, they can be assigned to a
replicate group and a T-test can be preformed on them for comparison. In the menus bar, go to
Analysis -> Replicate Groups -> Create Group and select the gels to be included in the group.
The T-tests available can be found by making a new analysis set and selecting “Statistic” for the
type.
Boolean Test: This test compares statistically significant spots from other analysis sets (i.e. spots
that have increased in one gel and spots that are found in one gel but not the other).
Viewing Graphs: To see all of the created graphs, click the Spot review Tool from the Graph
Tools bar. Clicking on a graph will zoom on the corresponding spots and highlight it.
Exporting Data: Data can be sent to the clip board for use in Microsoft Excel or other spread
sheet programs and can be saved as a TIFF file by going to File -> Export and selecting the
desired option.
Printing: Images with overlays and graphs can be printed with File -> Print.
25
APPENDIX 2: MATERIALS & REAGENT PREPARATION
Equipment
Autoclave
Criterion™ Cell (Bio-Rad catalog number 165-6001)
Incubator Shaker (capable of providing variable temperature)
Orbital Shaker equipped with utility tray
PDQuest™ 2-D Analysis Software (for PC, Bio-Rad catalog number 170-8603)
Pipettors capable of delivering 2-1000 mL
Power Supply (capable of providing 200 V)
PROTEAN® IEF System (Bio-Rad catalog number 165-4000)
Refrigerated Centrifuge
Refrigerator, -20 °C Freezer, -80 °C Freezer
VersaDoc™ Imager (for PC, Bio-Rad catalog number 170-8010)
UV/Vis Spectrophotometer
Laboratory Consumables
Erlenmeyer flasks, wide mouth, 250 mL
Fernbalch flasks, 3 or 4 L
Centrifuge tubes, 50 mL
Glass pipets, disposable, sterile
Eppendorf Tubes, 0.5 mL and 1.5 mL, sterile
Pipet Tips, capable of delivering 2-1000 mL, sterile
Plastic containers with lids for gel staining
Rehydration/Equilibration Tray with lid, disposable, 17 cm (25 pack, Bio-Rad catalog number
165-4015)
Semi-micro cuvettes, disposable
Semi-micro cuvettes, quartz
Reagents
Acetic acid
Benzonase (Serratia marcescens Recombinant Endonuclease, 250 units/mL)
Bio-Lyte 3/10 Ampholyte, 100X (1 mL, Bio-Rad catalog number 163-2094)
Bromophenol Blue
CHAPS
Criterion™ XT Precast Gel, 4-12% Bis-Tris, 1 mm, IPG+1 comb, 11 cm (Bio-Rad catalog
number 345-0127)
DTT
Escherichia coli K-12 MG1655
Ethanol
Glycerol
Glycine
Hydrochloric Acid
Iodoacetamide
Magnesium Chloride
Methanol
26
Mineral Oil
Novex™ Colloidal Blue Staining Kit (Invitrogen™ catalog number 45-7101)
ReadyStrip™ IGP Strips, pH 4.7-5.9, 11 cm (12 pack, Bio-Rad catalog number 163-2025)
ReadyStrip Buffer, 100X, pH 4.7-5.9 (1 mL, Bio-Rad catalog number 163-2097)
SDS
SDS-PAGE Standards, Broad Range (200 _L, Bio-Rad catalog number 161-0317)
Sodium Chloride
Sulfosalicylic Acid
SYPRO® Ruby Protein Gel Stain (1 L, catalog number 170-3125)
Trichloroacetic Acid
Tris
Tryptone
Urea (Ultra Pure)
XT MOPS Running Buffer, 20X (Bio-Rad catalog number 161-0788)
Yeast Extract
The following solution recipes provide sufficient material for six student groups.
Stock Solution
1.5 M Tris-HCl, pH 8.8
For 500 mL, dissolve 90.75 g Tris in 400 mL ddH2O. Adjust pH to 8.8 with 6 M HCl.
Bring to a final volume of 500 mL.
Working Solutions, presented in order of use
Luria Bertoni (LB) Media
Starter cultures: Dissolve 5.0 g tryptone, 5.0 g NaCl, and 2.5 g yeast extract in
425 mL ddH2O. Adjust pH to 7.0 and bring to final volume of 500 mL. Divide into ten
50 mL aliquots in 250 mL wide-mouth Erlenmeyer flasks. Autoclave for 20 min.
Heat shock culture: Dissolve 10 g tryptone, 10 g NaCl, and 5 g yeast extract in 950 mL
ddH2O. Adjust pH to 7.0, bring to final volume of 1 L, and autoclave for 20 min in a 4 L
Fernbalch flask.
40 mM Tris-HCl, pH 8.8
For 100 mL of solution, dilute 2.67 mL of the 1.5 M Tris-HCl stock solution with 80 mL
ddH2O. Check pH; adjust to pH 8.8 if necessary. Bring to a final volume of 100 mL.
Lysis Buffer, containing 8 M urea (FW 60.06), 4% w/v CHAPS, and 40 mM Tris
For 50 mL of solution, dissolve 24.02 g urea in approximately 20 mL ddH20 with gentle
heating. Add 0.24 g Tris base and dissolve, followed by 2.00 g CHAPS. Bring to a final
volume of 50 mL. Store at 4 °C. To resuspend urea that precipitates during storage,
place the solution in a warm water bath (30-35 °C). To minimize protein carbamylation,
avoid excessive heating during urea dissolution, and be sure the lysis buffer is at room
temperature before addition to cells.
If streaking is observed during IEF, consider supplementing the lysis buffer with
1 mM MgCl2 and 500 units Benzonase per gram of wet cells. Once the enzyme is added,
incubate the lysis mixture on ice for 15 min, then add Na2EDTA to a concentration of
2 mM. Proceed with protein concentration determination.
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100 mM MgCl2 (required only if performing Benzonase digest)
For 100 of mL solution, dissolve 2.03 g MgCl2 • 6 H2O (FW 203.3) in 80 mL ddH2O.
Bring to a final volume of 100 mL.
200 mM Na2EDTA (required only if performing Benzonase digest)
For 100 mL of solution, dissolve 7.44 g Na2EDTA • 2 H2O (FW 372.2) in 80 mL ddH2O.
Bring to a final volume of 100 mL.
Rehydration Buffer, containing 8 M urea, 2% w/v CHAPS, 0.001% w/v bromophenol blue,
1X (equivalent to 0.2% w/v) ampholyte, and 28 mg/mL DTT
For 25 mL of solution, dissolve 12.01 g urea, 0.50 g CHAPS, and 0.3 mg bromophenol
blue in 10 mL ddH2O. Bring to a final volume of 25 mL. Store in 2.5 mL aliquots at
-20 °C. Just before use dilute the appropriate 100X Bio-Lyte IEF buffer (e.g., either
100X Bio-Lyte 3/10 Ampholyte or 100X ReadyStrip 4.7-5.9 Buffer) to 1X by adding
25 mL to each thawed 2.5 mL aliquot. Finally add 7.0 mg DTT per aliquot. Do not
refreeze or reuse.
Equilibration Buffer I, containing 6 M urea, 20% glycerol, 0.375 M Tris-HCl, pH 7.8,
2% w/v SDS, and 130 mM DTT
For 10 mL of solution, combine 3.6 g urea, 0.2 g SDS, 2.5 mL 1.5 M Tris-HCl, pH 8.8,
2 mL glycerol, and 200 mg DTT (FW 154.2). Bring to a final volume of 10 mL with
ddH2O. Prepare just before use.
Equilibration Buffer II, containing 6 M urea, 20% glycerol, 0.375 M Tris-HCl, 2% w/v SDS, and
135 mM iodoacetamide
For 10 mL of solution, combine 3.6 g urea, 0.2 g SDS, 2.5 mL 1.5 M Tris-HCl, pH 8.8,
2 mL glycerol, and 250 mg iodoacetamide (FW 185.0). Bring to a final volume of 10 mL
with ddH2O. Prepare just before use.
1X XT MOPS Running Buffer
Prepare 1 L for each Criterion electrophoresis unit. Dilute 50 mL 20X XT MOPS
Running Buffer to a final volume of 1 L with ddH2O. Do not adjust pH.
Fixative Solution for Fluorescent Staining (SYPRO® Ruby), containing 7% v/v glacial acetic
acid and 10% v/v methanol
For 1 L of solution, add 70 mL glacial acetic acid and 100 mL methanol to approximately
700 mL ddH2O. Bring to a final volume of 1 L. Reserve half of the solution for gel
fixation prior to staining, and save the remainder for gel storage.
Fixative Solution for Colloidal Blue Staining, containing 12% w/v trichloroacetic acid and
3.5% w/v sulfosalicylic acid
For 1 L of solution, add 120 g TCA and 35 g sulfosalicylic acid to approximately
700 mL dd H2O. Once the acids are dissolved, bring to a final volume of 1 L.
Colloidal Blue Staining Solution, containing 20% v/v methanol, 20% v/v Novex Stainer A, and
2% v/v Novex Stainer B
For 1 L of solution add 200 mL methanol, 200 mL Novex Stainer A, and 20 mL Novex
Stainer B to approximately 500 mL ddH2O. Bring to a final volume of 1 L. For best
results, shake Stainer B well before use. Novex Stainers A and B are available as a
colloidal blue staining kit from Invitrogen.
Ammonium Sulfate Storage Solution for Colloidal Blue Stained Gels, containing 20% w/v
ammonium sulfate
For each gel requiring storage, prepare 100 mL of this solution by dissolving 20 g
ammonium sulfate in 60 mL ddH2O. Bring to a final volume of 100 mL.
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